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...

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.

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			<td>Friendship Medical Electronics Co., Ltd., Xi&#39;an,China</td>
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			<td>Tucker-Davis Technologies, Alachua, FL, USA</td>
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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</strong...

Watch this Scientific Journal Video about Morphological and Functional Evaluation of Ribbon Synapses at Specific Frequency Regions of the Mouse Cochlea at JoVE.com

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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11.6K Views.  Capital Medical University. 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.

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). ...

Telomerase Activity in the Various Regions of Mouse Brain: Non-Radioactive Telomerase Repeat Amplification Protocol (TRAP) Assay

Telomerase, a ribonucleoprotein, is responsible for maintaining the telomere length and therefore promoting genomic integrity, proliferation, and lifespan. In addition, telomerase protects the mitochondria from oxidative stress and confers resistance to apoptosis, suggesting its possible importance for the surviving of non-mitotic, highly active cells such as neurons. We previously demonstrated the ability of novel telomerase activators to increase telomerase activity and expression in the various mouse brain regions and to protect motor neurons cells from oxidative stress. These results strengthen the notion that telomerase is involved in the protection of neurons from various lesions. To underline the role of telomerase in the brain, we here compare the activity of telomerase in male and female mouse brain and its dependence on age. TRAP assay is a standard method for detecting telomerase activity in various tissues or cell lines. Here we demonstrate the analysis of telomerase activity in three regions of the mouse brain by non-denaturing protein extraction using CHAPS lysis buffer followed by modification of the standard TRAP assay.
In this 2-step assay, endogenous telomerase elongates a specific telomerase substrate (TS primer) by adding TTAGGG 6 bp repeats (telomerase reaction). The telomerase reaction products are amplified by PCR reaction creating a DNA ladder of 6 bp increments. The analysis of the DNA ladder is made by 4.5% high resolution agarose gel electrophoresis followed by staining with highly sensitive nucleic acid stain.
Compared to the traditional TRAP assay that utilize 32P labeled radioactive dCTP&#39;s for DNA detection and polyacrylamide gel electrophoresis for resolving the DNA ladder, this protocol offers a non-toxic time saving TRAP assay for evaluating telomerase activity in the mouse brain, demonstrating the ability to detect differences in telomerase activity in the various female and male mouse brain region.

Telomerase Activity in the Various Regions of Mouse Brain: Non-Radioactive Telomerase Repeat Amplification Protocol TRAP Assay

Telomerase is expressed in the neonatal brain and also in distinct regions of the adult brain. We present a non-toxic time saving TRAP assay for the analysis of telomerase activity in various regions of the mouse brain and detection of differences in telomerase activity between male and female mouse brains.

Telomerase, a ribonucleoprotein, is responsible for maintaining the telomere length and therefore promoting genomic integrity, proliferation, and ...

Functional and Morphological Assessment of Diaphragm Innervation by Phrenic Motor Neurons

This protocol specifically focuses on tools for assessing phrenic motor neuron (PhMN) innervation of the diaphragm at both the electrophysiological and morphological levels. Compound muscle action potential (CMAP) recording following phrenic nerve stimulation can be used to quantitatively assess functional diaphragm innervation by PhMNs of the cervical spinal cord in vivo in anesthetized rats and mice. Because CMAPs represent simultaneous recording of all myofibers of the whole hemi-diaphragm, it is useful to also examine the phenotypes of individual motor axons and myofibers at the diaphragm NMJ in order to track disease- and therapy-relevant morphological changes such as partial and complete denervation, regenerative sprouting and reinnervation. This can be accomplished via whole-mount immunohistochemistry (IHC) of the diaphragm, followed by detailed morphological assessment of individual NMJs throughout the muscle. Combining CMAPs and NMJ analysis provides a powerful approach for quantitatively studying diaphragmatic innervation in rodent models of CNS and PNS disease.

Compound muscle action potential recording quantitatively assesses functional diaphragm innervation by phrenic motor neurons. Whole-mount diaphragm immunohistochemistry assesses morphological innervation at individual neuromuscular junctions. The goal of this protocol is to demonstrate how these two powerful methodologies can be used in various rodent models of spinal cord disease.

This protocol specifically focuses on tools for assessing phrenic motor neuron (PhMN) innervation of the diaphragm at both the electrophysiological and ...

High-Resolution Quantitative Immunogold Analysis of Membrane Receptors at Retinal Ribbon Synapses

Retinal ganglion cells (RGCs) receive excitatory glutamatergic input from bipolar cells. Synaptic excitation of RGCs is mediated postsynaptically by NMDA receptors (NMDARs) and AMPA receptors (AMPARs). Physiological data have indicated that glutamate receptors at RGCs are expressed not only in postsynaptic but also in perisynaptic or extrasynaptic membrane compartments. However, precise anatomical locations for glutamate receptors at RGC synapses have not been determined. Although a high-resolution quantitative analysis of glutamate receptors at central synapses is widely employed, this approach has had only limited success in the retina. We developed a postembedding immunogold method for analysis of membrane receptors, making it possible to estimate the number, density and variability of these receptors at retinal ribbon synapses. Here we describe the tools, reagents, and the practical steps that are needed for: 1) successful preparation of retinal fixation, 2) freeze-substitution, 3) postembedding immunogold electron microscope (EM) immunocytochemistry and, 4) quantitative visualization of glutamate receptors at ribbon synapses.

The postembedding immunogold method is one of the most effective ways to provide high-resolution analyses of the subcellular localization of specific molecules. Here we describe a protocol to quantitatively analyze glutamate receptors at retinal ribbon synapses.

Retinal ganglion cells (RGCs) receive excitatory glutamatergic input from bipolar cells. Synaptic excitation of RGCs is mediated postsynaptically by NMDA ...

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 ...

Rapid and Specific Immunomagnetic Isolation of Mouse Primary Oligodendrocytes

The efficient and robust isolation and culture of primary oligodendrocytes (OLs) is a valuable tool for the in vitro study of the development of oligodendroglia as well as the biology of demyelinating diseases such as multiple sclerosis and Pelizaeus-Merzbacher-like disease (PMLD). Here, we present a simple and efficient selection method for the immunomagnetic isolation of stage three O4+ preoligodendrocytes cells from neonatal mice pups. Since immature OL constitute more than 80% of the rodent-brain white matter at postnatal day 7 (P7) this isolation method not only ensures high cellular yield, but also the specific isolation of OLs already committed to the oligodendroglial lineage, decreasing the possibility of isolating contaminating cells such as astrocytes and other cells from the mouse brain. This method is a modification of the techniques reported previously, and provides oligodendrocyte preparation purity above 80% in about 4 h.

We describe the immunomagnetic isolation of primary mouse oligodendrocytes, which allows the rapid and specific isolation of the cells for in vitro culture.

The efficient and robust isolation and culture of primary oligodendrocytes (OLs) is a valuable tool for the in vitro study of the development of ...

Functional Evaluation of Olfactory Pathways in Living Xenopus Tadpoles

Xenopus tadpoles offer a unique platform to investigate the function of the nervous system. They provide multiple experimental advantages, such as accessibility to numerous imaging approaches, electrophysiological techniques and behavioral assays. The Xenopus tadpole olfactory system is particularly well suited to investigate the function of synapses established during normal development or reformed after injury. Here, we describe methodologies to evaluate the processing of olfactory information in living Xenopus larvae. We outline a combination of in vivo measurements of presynaptic calcium responses in glomeruli of the olfactory bulb with olfactory-guided behavior assays. Methods can be combined with the transection of olfactory nerves to study the rewiring of synaptic connectivity. Experiments are presented using both wild-type and genetically modified animals expressing GFP reporters in central nervous system cells. Application of the approaches described to genetically modified tadpoles can be useful for unraveling the molecular bases that define vertebrate behavior.

Functional Evaluation of Olfactory Pathways in Living Xenopus Tadpoles

Xenopus tadpoles offer a unique platform to investigate the function of the nervous system in vivo. We describe methodologies to evaluate the processing of olfactory information in living Xenopus larvae in normal rearing conditions or after injury.

Xenopus tadpoles offer a unique platform to investigate the function of the nervous system. They provide multiple experimental advantages, such as ...

Ex Vivo Imaging of Cell-specific Calcium Signaling at the Tripartite Synapse of the Mouse Diaphragm

The electrical activity of cells in tissues can be monitored by electrophysiological techniques, but these are usually limited to the analysis of individual cells. Since an increase of intracellular calcium (Ca2+) in the cytosol often occurs because of the electrical activity, or in response to a myriad of other stimuli, this process can be monitored by the imaging of cells loaded with fluorescent calcium-sensitive dyes.&#160; However, it is difficult to image this response in an individual cell type within whole tissue because these dyes are taken up by all cell types within the tissue. In contrast, genetically encoded calcium indicators (GECIs) can be expressed by an individual cell type and fluoresce in response to an increase of intracellular Ca2+, thus permitting the imaging of Ca2+ signaling in entire populations of individual cell types. Here, we apply the use of the GECIs GCaMP3/6 to the mouse neuromuscular junction, a tripartite synapse between motor neurons, skeletal muscle, and terminal/perisynaptic Schwann cells. We demonstrate the utility of this technique in classic ex vivo tissue preparations. Using an optical splitter, we perform dual-wavelength imaging of dynamic Ca2+ signals and a static label of the neuromuscular junction (NMJ) in an approach that could be easily adapted to monitor two cell-specific GECI or genetically encoded voltage indicators (GEVI) simultaneously. Finally, we discuss the routines used to capture spatial maps of fluorescence intensity. Together, these optical, transgenic, and analytic techniques can be employed to study the biological activity of distinct cell subpopulations at the NMJ in a wide variety of contexts.

Ex Vivo Imaging of Cell-specific Calcium Signaling at the Tripartite Synapse of the Mouse Diaphragm

Here we present a protocol to image calcium signaling in populations of individual cell types at the murine neuromuscular junction.

The electrical activity of cells in tissues can be monitored by electrophysiological techniques, but these are usually limited to the analysis of ...

Stimulus-specific Cortical Visual Evoked Potential Morphological Patterns

This paper presents a methodology for the recording and analysis of cortical visual evoked potentials (CVEPs) in response to various visual stimuli using 128-channel high-density electroencephalography (EEG). The specific aim of the described stimuli and analyses is to examine whether it is feasible to replicate previously reported CVEP morphological patterns elicited by an apparent motion stimulus, designed to simultaneously stimulate both ventral and dorsal central visual networks, using object and motion stimuli designed to separately stimulate ventral and dorsal visual cortical networks.&#160; Four visual paradigms are presented: 1. Randomized visual objects with consistent temporal presentation. 2. Randomized visual objects with inconsistent temporal presentation (or jitter).&#160; 3. Visual motion via a radial field of coherent central dot motion without jitter.&#160; 4. Visual motion via a radial field of coherent central dot motion with jitter.&#160; These four paradigms are presented in a pseudo-randomized order for each participant.&#160; Jitter is introduced in order to view how possible anticipatory-related effects may affect the morphology of the object-onset and motion-onset CVEP response.&#160; EEG data analyses are described in detail, including steps of data exportation from and importation to signal processing platforms, bad channel identification&#160;and removal, artifact rejection, averaging, and categorization of average CVEP morphological pattern type based upon latency ranges of component peaks. Representative data show that the methodological approach is indeed sensitive in eliciting differential object-onset and motion-onset CVEP morphological patterns and may, therefore, be useful in addressing the larger research aim. Given the high temporal resolution of EEG and the possible application of high-density EEG in source localization analyses, this protocol is ideal for the investigation of distinct CVEP morphological patterns and the underlying neural mechanisms which generate these differential responses.&#160; &#160; &#160;&#160;

In this paper, we present a protocol to investigate differential cortical visual evoked potential morphological patterns through stimulation of ventral and dorsal networks using high-density EEG. Visual object and motion stimulus paradigms, with and without temporal jitter, are described. Visual evoked potential morphological analyses are also outlined.&#160;&#160;

This paper presents a methodology for the recording and analysis of cortical visual evoked potentials (CVEPs) in response to various visual stimuli using ...

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

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Functional Evaluation of Olfactory Pathways in Living Xenopus Tadpoles

Ex Vivo Imaging of Cell-specific Calcium Signaling at the Tripartite Synapse of the Mouse Diaphragm

Stimulus-specific Cortical Visual Evoked Potential Morphological Patterns