Name: whole mount dissection and immunofluorescence of the adult mouse cochlea
Uploaded: 2016-01-01T15:00:00.000+00:00
Duration: 12 min 2 s
Description: Watch this Scientific Journal Video about Whole Mount Dissection and Immunofluorescence of the Adult Mouse Cochlea at JoVE.com

This work was supported in part by a grant from the Office of Naval Research (N000141310569). The Southern Illinois University School of Medicine Research Imaging Facility equipment was supported by the National Center for Research Resources-Health (S10RR027716).

Ethics Statement: Procedures involving animal subjects have been approved by the Institutional Animal Care and Use Committee at Southern Illinois University School of Medicine.
1. Extraction of Temporal Bones
<ol>
	<li>Identify temporal bones in the base of the mouse skull13 and scrape away the cranial&#160;nerves using standard pattern forceps.</li>
	<li>Place standard pattern forceps at the tip of the otic capsule and with the thumb of the opposite hand press down on the posterior semicircular canal to dislodge the encapsulated cochlea.</li>
	<li>Free the bottom half of the temporal bone from the skull manually with the thumb and index finger or by using 10.5 cm fine scissors.</li>
</ol>
2. Post-fix Temporal Bones
<ol>
	<li>Place temporal bones into 2 ml microcentrifuge tubes containing 250 - 500 &#181;l 4% paraformaldehyde (PFA) diluted in 10 mM phosphate buffered saline (PBS) pH 7.4 and incubate at RT&#160;for 2 - 20 hr. 
	NOTE: No opening of the apical cap or injection of PFA into the round or oval window is needed.&#160;Recommend using methanol free, ultra-pure, electron microscopy (EM) grade PFA that can be purchased at a 16% concentration in glass vials. Once a vial is opened and diluted 1:4 in PBS, making a 4% solution, it can be used for up to 2 weeks when stored at 4 oC&#160;(Some antibodies require short fixation and others can tolerate O/N (O/N)&#160;fixation).</li>
</ol>
3. Decalcify Temporal Bones
<ol>
	<li>After fixation, remove PFA using a pipette and replace with 120 mM ethylenediaminetetraacetic acid (EDTA), slightly more than 2 ml. Make sure to fill the entire 2 ml microcentrifuge tube to prevent air bubbles when the tube is closed.
	<ol>
		<li>If not decalcifying immediately, replace PFA with 10 mM PBS pH 7.4 and store samples at 4 oC. 
		NOTE: After fixation, temporal bones can be stored for variable amounts of time before decalcification depending on the antigens being examined.</li>
	</ol>
	</li>
	<li>Place tubes on end-over-end rotator and rotate at 4 rpm at RT. Change EDTA solution daily using a pipette. Length of decalcification depends on the age of the sample and the preference of the scientist doing the dissection.
	<ol>
		<li>Incubate temporal bones in EDTA using the following guidelines for incubation times: 
		For samples postnatal day (P) 8 to P15: 2 - 4 hr; For samples P15 to P21: O/N; For samples P21 to P30: 2 O/N; For samples older than P30: 3 or more O/N. 
		NOTE: Decalcification times are subject to users&#39; preference and can be extended as needed. Decalcification time can be reduced by changing the EDTA solution twice a day, about 8 hr apart. Some labs add 1% PFA to the EDTA solution when decalcifying for longer than 3 days to prevent contamination.</li>
	</ol>
	</li>
	<li>To determine if the sample is adequately decalcified, place temporal bones on a silicon elastomer-coated dissection dish and gently press forceps onto the snail-shaped cochlea. If the tissue is spongey, then decalcification is complete.</li>
	<li>Once decalcification is complete, remove EDTA using a pipette and add 500 -1,000 &#181;l of 10 mM PBS pH 7.4. Store samples at 4 oC until ready to dissect.</li>
</ol>
4. Create Silicone Elastomer-coated Dissection Dish
<ol>
	<li>Combine the base and curing agent of a silicone elastomer encapsulant kit according to the manufacturer&#39;s instructions and mix thoroughly.</li>
	<li>Add approximately 2 - 3 tablespoons of powdered charcoal until the solution is black and mix well.&#160; 
	NOTE: Liquid ink or liquid charcoal will not mix with the solution and cannot be used.&#160;Silicone elastomer encapsulant kits can be purchased in black, allowing step 4.2 to be omitted.</li>
	<li>Pour the solution into 60 mm glass or plastic petri dishes, filling enough to coat the bottom. If bubbles are present, puff with air on the surface. Let stand at least 24 hr at RT to set. 
	NOTE: Silicone elastomer-coated dishes can be used repeatedly for years. However do not use ethanol for cleaning, as this will cause the silicone elastomer to crack.</li>
</ol>
5. Whole Mount Dissection of the Cochlea (for P7 and Older Samples)
<ol>
	<li>Separate the basal turn from middle/apical turns
	<ol>
		<li>Place one decalcified temporal bone in a silicone elastomer-coated dissection dish filled two-thirds full with 10 mM PBS pH 7.4 and use a stereo dissection microscope for the following steps.</li>
		<li>Hold the temporal bone in the vestibular region with #4 or #5 straight jeweler&#39;s forceps and using 5 mm Vannas-Tubingen spring scissors, cut away excess otic capsule tissue along the sides and above the apex.</li>
		<li>Using 2.5 mm Vannas spring scissors, insert one blade into the oval window and make several small cuts along the spiral ligament/lateral wall of the basal turn.</li>
		<li>Using 5 mm Vannas-Tubingen spring scissors, insert one blade into the region just cut and place the other blade on the outside the temporal bone, medial to the oval window. This cut separates the basal turn from middle and apical turns.</li>
	</ol>
	</li>
	<li>Complete dissection of the basal turn.
	<ol>
		<li>Using 2.5 mm Vannas spring scissors, cut the spiral ganglion nerve fibers that connect to the modiolus to release the tension in the basal turn and cut below the basal turn to separate from vestibular organs.</li>
		<li>Using 2.5 mm Vannas spring scissors, make a series of small cuts to remove the spiral ligament/lateral wall from both above and below the organ of Corti. Use #4 or #5 straight jeweler&#39;s forceps to guide the tissue. Pin the spiral ganglion nerve fibers to the silicone elastomer-coated dissection dish, but do not hold onto this region as the tissue will tear.</li>
		<li>During the previous steps, some of Reissner&#39;s membrane will often be removed. If any still remains, grasp the Reissner&#39;s membrane with #4 or #5 straight jeweler&#39;s forceps and pull away from the organ of Corti. 
		NOTE: The tectorial membrane is rarely visible and typically floats away without requiring a specific step for removal.</li>
		<li>Finally make several cuts to reduce the thickness of the spiral ganglion axons, to make the turn as flat as possible and use #4 or #5 straight jeweler&#39;s forceps to transfer the dissected basal turn, by grasping the remaining axons of the spiral ganglion, to a 48-well plate (or chamber slide) containing ~500 &#181;l of 10 mM PBS pH 7.4.</li>
	</ol>
	</li>
	<li>Separate middle and apical turns
	<ol>
		<li>Place the remaining two-thirds of the cochlea, apical side down.</li>
		<li>Using 2.5 mm Vannas spring scissors, insert one blade into the scala media where the middle turn used to be connected to the basal turn, and make several small cuts along the spiral ligament/lateral wall of the middle turn.</li>
		<li>Using 5 mm Vannas-Tubingen spring scissors, insert one blade into the region just cut with the middle turn placed on top of the blade, and place the other blade on the outside the bony labyrinth, at a 90o angle from the apical tip. This separates the middle turn from the apical turn.</li>
	</ol>
	</li>
	<li>Complete dissection of the middle turn in a similar manner to the basal turn (see steps 5.2.2 to 5.2.4).</li>
	<li>Complete dissection of the apical turn.
	<ol>
		<li>Using 2.5 mm Vannas spring scissors, open the cap that covers the apical turn. Complete dissection in a similar manner to the basal turn (see steps 5.2.2 to 5.2.4). 
		NOTE: Dissected cochlear turns can be stored in 10 mM PBS pH 7.4 in a 48-well plate (or chamber slide) at 4 oC for several weeks before immunostaining. However PBS will evaporate and needs to be periodically replenished and storage longer than 2 - 3 weeks can result in bacterial or fungal growth on the tissue which can decrease the quality of the image. We recommend long-term storage of samples as undissected temporal bones.</li>
	</ol>
	</li>
</ol>
6. Immunostaining with Fluorescently Tagged Antibodies
<ol>
	<li>After the dissection, store each cochlear turn in a separate well in a 48-well plate (or chamber slide),&#160;submerged in ~500 &#181;l of 10 mM PBS pH 7.4. For each of the following steps the cochlear turn should be submerged in liquid, not floating on top or stuck to the side of the well.&#160; 
	NOTE: When removing liquid from each well, it is easy to lose the cochlear turns or suck it up in the pipette tip. Changing solutions with a 200 &#181;l pipette tip using a dissection scope will help prevent this.</li>
	<li>Using a pipette, remove PBS from each well and replace with ~200 - 300 &#181;l per well of blocking/permeabilization solution (1% Triton X-100, 1% bovine serum albumin (BSA), and 10% normal goat serum (NGS) diluted in 10 mM PBS pH 7.4). Incubate 1 hr at RT on a 3D rotator. 
	NOTE: If any primary antibody used was made in a goat host, then a secondary anti-goat antibody will be needed and NGS should NOT be used for any of the steps. Normal horse serum can be used as a replacement for NGS.</li>
	<li>Remove blocking/permeabilization solution using a pipette and replace with ~100 &#181;l per well of primary antibody solution (0.1% Triton X-100, 1% BSA, and 5% NGS diluted in 10 mM PBS pH 7.4). The dilution factor for each primary antibody varies. Incubate O/N (minimum 14 hr) at 4 oC on a 3D rotator. 
	NOTE: If more than one primary antibody is used, all can be combined into the same solution for incubation, just make sure that each primary antibody has a different host.</li>
	<li>Remove primary antibody solution using a pipette and perform 3 washes of 10 mM PBS pH 7.4 at ~500 &#181;l per well. Each wash incubates a minimum of 5 min at RT on a 3D rotator.</li>
	<li>Remove last PBS wash using a pipette and replace with ~100 &#181;l per well of secondary antibody solution (0.1% Triton X-100, 1% BSA, and 5% NGS diluted in 10 mM PBS pH 7.4). The dilution factor for each fluorescently tagged secondary antibody is usually 1:500 or 1:1,000. Place 48-well plate in a black box to protect fluorescently tagged secondary antibodies from light. Incubate 2 - 3 hr at RT on a 3D rotator. 
	NOTE: If more than one secondary antibody is needed, they can be combined into the same solution for incubation. Make sure there is no possibility of cross-labeling (e.g., using goat anti-rabbit and chicken anti-goat secondary antibodies)</li>
	<li>Remove secondary antibody solution using a pipette and perform 3 washes of 10 mM PBS pH 7.4 at ~500 &#181;l per well. Incubate each wash for a minimum of 5 min at RT on a 3D rotator. Keep 48-well plate in a black box to protect from light.</li>
	<li>Remove last PBS wash using a pipette and replace with ~100 &#181;l per well of Hoechst 33342 (diluted 1:2,000 in 10 mM PBS pH 7.4) to label nuclei. Incubate 15 - 20 min at RT on a 3D rotator. Keep 48-well plate in a black box to protect from light. Do NOT incubate longer than 20 min.</li>
	<li>Remove Hoechst solution using a pipette and perform 3 washes of 10 mM PBS pH 7.4 at ~500 &#181;l per well. Incubate each wash for a minimum of 5 min at RT on a 3D rotator. Keep 48-well plate in a black box to protect from light.</li>
	<li>All steps can be extended, except Hoechst incubation, by several hours if needed. To pause the reaction at any stage, just submerge samples in ~500 ul of 10mM PBS pH7.4 and store the 48 -well plate (or chamber slide) at 4 oC until ready to resume the protocol hours or 1 - 2 days later. 
	&#160;</li>
</ol>
7. Mount Cochlear Turns on Slides
<ol>
	<li>Label slides with pertinent information about the sample and antibodies used.</li>
	<li>Pipette ~50 &#181;l of mounting media onto each slide and be careful to prevent bubbles. Centrifuge the tube containing the mounting media to remove&#160;any bubbles.</li>
	<li>Using #4 or #5 straight jeweler&#39;s forceps, grasp the axons of the spiral ganglion to gently transfer one cochlear turn from the 48-well plate to the slide and place in the mounting media. Mount one cochlear turn per slide to prevent light exposure and photobleaching during the imaging process.</li>
	<li>Use a stereo dissection microscope to ensure that cochlear turn is not folded, twisted, or near an air bubble.&#160;If any of these conditions occur, use #4 or #5 straight jeweler&#8217;s forceps to reposition the cochlear turn.</li>
	<li>Place one end of a coverslip on the slide and gently release to let the coverslip fall.</li>
	<li>Use a stereo dissection microscope to ensure that the cochlear turn is not folded, twisted, or near an air bubble. If any of these conditions occur, gently move the coverslip back and forth to reposition the cochlear turn.</li>
	<li>Place slides in a slide folder so that they lay flat. Let mounting media cure O/N at RT (keep in the DARK)</li>
	<li>Seal coverslips with clear nail polish and store at RT or -20&#160;oC until imaged. Slides can be stored in a slide folder or slide box. 
	NOTE:&#160;Slides can be stored long term at -20 oC or -80 oC and fluorescence will be maintained for several months.</li>
	<li>Image slides using a confocal microscope with the appropriate wavelength based on the secondary antibodies used during the immunostaining procedure. Brightness and contrast adjustments can be performed using the imaging software provided by the confocal vendor.</li>
</ol>

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The authors have nothing to disclose.

There are several critical steps for successful whole mount dissection and immunostaining. However before either of these methods are performed, proper fixation of the cochlear tissue is needed. We recommend using methanol free, ultra-pure, EM grade PFA. PFA made from powder can have traces of methanol and an unstable pH which decreases the quality of immunofluorescence. Other groups have also shown that similar dissections are possible using fixatives that do not contain formaldehyde14-16. The length of fixat...

The spiral-shaped cochlea of the inner ear, contained within the temporal bone, houses the organ of Corti, the auditory sensory end organ in mammals. The cochlea is tonotopically organized and commonly divided into apical, middle, and basal turns corresponding to different frequency regions with high frequency sound detection in the base and low frequency detection in the apex1. Hair cells, the mechanosensory cells of the organ of Corti, run the length of the cochlea, which is approximately 6 mm long in mice2,3. These cells convert the mechanical energy of sound waves, which are transmitted through the fluid-filled membranous labyrinth, into neural signals that are processed by central auditory structures. The technique described here provides a method for preparing whole mounts of the organ of Corti after calcification of the cochlea is complete (for samples ranging from one week of age to adulthood). We also present a method for immunostaining the whole mounted cochlear tissue. Cochlear whole mounts are crucial for visualization of all hair cells and surrounding supporting cells in their natural spatial arrangements and allow for analysis in three dimensions with the use of confocal microscopy.
Drs. Hans Engstrom and Harlow Ades originally described a whole mount cochlear dissection method in 1966. They detailed a technique to rapidly fix and dissect calcified cochleae submerged in liquid from a variety of mammals, preserving short intact segments of the organ of Corti for microscopic analysis4. The dissection of an unfixed, calcified rat cochlea has also been illustrated in an instructional video5. Drs. Barbara Bohne and Gary Harding at Washington University made several important modifications to this method. In their version of the cochlear whole mount method, the temporal bone was decalcified, embedded in plastic, and five half-turns or ten quarter-turns were dissected6,7. Dr. Charles Liberman and colleagues at Eaton Peabody Laboratories, Massachusetts Eye and Ear Infirmary, modified this technique so that plastic embedding was not required8. Further modification of the technique occurred in Dr. Jian Zuo&#39;s lab at St. Jude Children&#39;s Research Hospital9-12 which informed the dissection method presented here. We use a different strategy to gain access to the organ of Corti than Bohne and Liberman, which allows isolation of complete apical, middle, and basal turns. Thus the dissected tissue is larger and less likely to be lost or damaged during the dissection or immunostaining processes. In addition, the current method facilitates measurement of the distance from the apical tip or basal hook to identify a frequency region.
Although many labs perform immunostaining of cochlear tissue, it is unclear where this method originated. As a result there are various recipes for blocking buffers and antibody incubation buffers that may affect the performance of individual primary antibodies. Here, we present one method for immunostaining with fluorescently tagged antibodies that is applicable to most commonly used antibodies in the auditory field.
The complex shape of the cochlea, delicate structure of the organ of Corti, and bony encasement provide a challenge for histological and biochemical analysis. A variety of techniques are currently used in the hearing field to surmount these difficult features and visualize the cells within the organ of Corti, each technique with its own advantages and disadvantages. The protocol presented here allows for whole mount dissection of the adult mouse cochlea and, with slight modification, can potentially be used to examine the critical structures within the cochleae from a variety of other model organisms used in the field.

<table><tbody><tr><td>Standard&amp;nbsp; pattern forceps</td><td>Fine Science Tools</td><td>11000-12</td><td>can be purchased from other vendors</td></tr><tr><td>10.5 cm fine scissors</td><td>Fine Science Tools</td><td>14060-11</td><td>can be purchased from other vendors</td></tr><tr><td>2 ml microcentrifuge tubes</td><td>MidSci</td><td>AVSS2000</td><td>can be purchased from other vendors</td></tr><tr><td>16% formaldehyde, methanol free, ultra pure, EM grade</td><td>Polysciences</td><td>18814</td><td>TOXIC --wear gloves and cannot be disposed of in the sink. Can be purchased from other vendors.&amp;nbsp;</td></tr><tr><td>PBS pH 7.4&amp;nbsp;</td><td>Sigma</td><td>P3813-10PAK</td><td>can be purchased from other vendors</td></tr><tr><td>EDTA</td><td>Fisher</td><td>BP118-500</td><td>can be purchased from other vendors</td></tr><tr><td>end-over-end tube rotator</td><td>Fisher</td><td>05-450-127</td><td>can be purchased from other vendors</td></tr><tr><td>60 mm petri dish</td><td>Fisher</td><td>50-202-037</td><td>can be purchased from other vendors</td></tr><tr><td>Dow Corning Sylgard 184 silicone encapsulant kit</td><td>Ellsworth Adhesives</td><td>184 SIL ELAST KIT 0.5KG</td><td></td></tr><tr><td>activated charcoal</td><td>Fisher</td><td>AC134372500</td><td>can be purchased from other vendors</td></tr><tr><td>stereo dissection microscope</td><td>Zeiss</td><td>Stemi 2000</td><td>can be purchased from other vendors</td></tr><tr><td>Dumont #4 jeweler&#039;s forceps</td><td>Fine Science Tools</td><td>11241-30</td><td></td></tr><tr><td>Dumont #5 jeweler&#039;s forceps</td><td>Fine Science Tools</td><td>11251-20</td><td></td></tr><tr><td>2.5 mm Vannas spring scissors</td><td>Fine Science Tools</td><td>15001-08</td><td>curved&amp;nbsp;</td></tr><tr><td>5 mm Vannas-Tubingen spring scissors</td><td>Fine Science Tools</td><td>15003-08</td><td>straight</td></tr><tr><td>48-well plates</td><td>Fisher</td><td>08-772-52</td><td>can be purchased from other vendors</td></tr><tr><td>8-well chamber slides</td><td>Fisher</td><td>1256518</td><td>can be purchased from other vendors</td></tr><tr><td>Triton X-100</td><td>Sigma</td><td>X100-500</td><td>can be purchased from other vendors</td></tr><tr><td>BSA, fraction V</td><td>Fisher</td><td>BP1605</td><td>can be purchased from other vendors</td></tr><tr><td>NGS</td><td>Vector labs</td><td>S-1000</td><td>can be purchased from other vendors</td></tr><tr><td>NHS</td><td>Vector labs</td><td>S-2000</td><td>can be purchased from other vendors</td></tr><tr><td>3D rotator</td><td>Midsci</td><td>R3D-710</td><td>can be purchased from other vendors</td></tr><tr><td>Western blot incubation box XL</td><td>Licor</td><td>929-97401</td><td></td></tr><tr><td>Hoechst 33342</td><td>Life Technologies</td><td>H3570</td><td>can be purchased from other vendors</td></tr><tr><td>Prolong gold antifade mounting media</td><td>Life Technologies</td><td>P36930</td><td>can be purchased from other vendors,but mounting medias vary in their ability to protect against photobleaching</td></tr><tr><td>Superfrost Plus Slides</td><td>Fisher</td><td>12-550-15</td><td>can be purchased from other vendors</td></tr><tr><td>coverslips 22 x 22 x 1</td><td>Fisher</td><td>12-548-B</td><td>can be purchased from other vendors</td></tr><tr><td>clear nail polish</td><td>Local drug store</td><td></td><td>can be purchased from other vendors</td></tr><tr><td>cardboard slide folder</td><td>Fisher</td><td>12-587-10</td><td>can be purchased from other vendors</td></tr><tr><td>plastic slide box</td><td>Fisher</td><td>03-448-10</td><td>can be purchased from other vendors</td></tr></tbody></table>

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 organ of Corti as three intact cochlear turns (apex, middle, and base) from cochlear tissue that is calcified, with key dissection steps presented in Figure 1. During the first postnatal week of development, calcification of the mouse cochlea is incomplete and a more simple dissection method can be used13. Using the neonatal whole mount dissection method with cochlea from P7 and older mice results in tears and shredding of the...

Watch this Scientific Journal Video about Whole Mount Dissection and Immunofluorescence of the Adult Mouse Cochlea at JoVE.com

Whole Mount Dissection and Immunofluorescence of the Adult Mouse Cochlea

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 ...

whole-mount-dissection-immunofluorescence-adult-mouse

Research

JoVE Journal

Neuroscience

54.8K Views.  Southern Illinois University, School of Medicine. 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.

Video: Whole Mount Dissection and Immunofluorescence of the Adult Mouse Cochlea

Vibratome Sectioning for Enhanced Preservation of the Cytoarchitecture of the Mammalian Organ of Corti

The mammalian organ of Corti is a highly ordered cellular mosaic of mechanosensory hair and nonsensory supporting cells (reviewed in 1,2).Visualization of this cellular mosaic often requires that the organ of Corti is cross-sectioned. In particular, the nonsensory pillar and Deiters' cells, whose nuclei are located basally with respect to the hair cells, cannot be visualized without cross-sectioning the organ of Corti. However, the delicate cytoarchitecture of the mammalian organ of Corti, including the fine cytoplasmic processes of the pillar and Deiters' cells, is difficult to preserve by routine histological procedures such as paraffin and cryo-sectioning, which are compatible with standard immunohistochemical staining techniques.
Here I describe a simple and robust procedure consisting of vibratome sectioning of the cochlea, immunohistochemical staining of these vibratome sections in whole mount, followed by confocal microscopy. This procedure has been used widely for immunhistochemical analysis of multiple organs, including the mouse limb bud, zebrafish gut, liver, pancreas, and heart (see 3-6 for selected examples). In addition, this procedure was sucessful for both imaging and quantitificaton of pillar cell number in mutant and control organs of Corti in both embryos and adult mice 7. This method, however, is currently not widely used to examine the mammalian organ of Corti. The potential for this procedure to both provide enhanced preservation of the fine cytoarchitecture of the adult organ of Corti and allow for quantification of various cell types is described.

A simple procedure of vibratome sectioning the organ of Corti, followed by immunohistochemistry and confocal microscopy is described. This procedure allows for improved preservation of the fine cytoarchitecture of the mammalian organ of Corti, and consequently allows for accurate quantification of cell types.

The mammalian organ of Corti is a highly ordered cellular mosaic of mechanosensory hair and nonsensory supporting cells (reviewed in 1,2).Visualization of ...

Whole-mount Confocal Microscopy for Adult Ear Skin: A Model System to Study Neuro-vascular Branching Morphogenesis and Immune Cell Distribution

Here, we present a protocol of a whole-mount adult ear skin imaging technique to study comprehensive three-dimensional neuro-vascular branching morphogenesis and patterning, as well as immune cell distribution at a cellular level. The analysis of peripheral nerve and blood vessel anatomical structures in adult tissues provides some insights into the understanding of functional neuro-vascular wiring and neuro-vascular degeneration in pathological conditions such as wound healing. As a highly informative model system, we have focused our studies on adult ear skin, which is readily accessible for dissection. Our simple and reproducible protocol provides an accurate depiction of the cellular components in the entire skin, such as peripheral nerves (sensory axons, sympathetic axons, and Schwann cells), blood vessels (endothelial cells and vascular smooth muscle cells), and inflammatory cells. We believe this protocol will pave the way to investigate morphological abnormalities in peripheral nerves and blood vessels as well as the inflammation in the adult ear skin under different pathological conditions.

Here, we describe a high resolution whole-mount imaging method in the entire adult mouse ear skin, which enables us to visualize branching morphogenesis and patterning of peripheral nerves and blood vessels, as well as immune cell distribution.

Here, we present a protocol of a whole-mount adult ear skin imaging technique to study comprehensive three-dimensional neuro-vascular branching ...

Developmental Biology

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

Cryosectioning and Immunostaining Mouse Inner Ear Tissue: From Embryonic to Adult Stages

This protocol details general histology methods for preparing inner ear samples from embryonic, neonatal, and adult mice. The purpose of this protocol is to provide a straightforward and standardized method for inner ear tissue processing for researchers, possibly new to the field, interested in working with the mouse cochlea. Included here are protocols for dissection, fixation, embedding, cryosectioning, and immunostaining. Section immunostaining is one of the best methods for examining individual cells within the context of the entire cochlea.
Key steps include dissecting the inner ear away from the skull, fixing the tissue using 4% paraformaldehyde, embedding with Optimal Cutting Temperature compound, cryosectioning, and immunostaining with antibodies targeting specific proteins expressed by the cochlea.
Included in our methods are special considerations made for the cochlea given its 
shape and structure.&#160;Reasonably good focus and dissection skills are needed, as tissue damage can affect the quality of immunostaining and consistency among samples. However, with the procedures we outline, most users can be trained in a few weeks to make beautiful preparations. Overall, this protocol offers a valuable way to promote research to understand auditory development, function, and disease in mouse models.

This protocol, intended for novice users of mouse inner ear tissue, comprehensively details steps for processing mouse inner ears at different developmental stages for section immunostaining.

This protocol details general histology methods for preparing inner ear samples from embryonic, neonatal, and adult mice. The purpose of this protocol is ...

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.

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 ...

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 Time-lapse Live-Cell Imaging to Explore Cell Migration toward the Organ of Corti

To study the effects of mesenchymal stem cells (MSCs) on cell regeneration and treatment, this method tracks MSC migration and morphological changes after co-culture with cochlear epithelium. The organ of Corti was immobilized on a plastic coverslip by pressing a portion of the Reissner's membrane generated during the dissection. MSCs confined by a glass cylinder migrated toward cochlear epithelium when the cylinder was removed. Their predominant localization was observed in the modiolus of the organ of Corti, aligned in a direction similarly to that of the nerve fibers. However, some MSCs were localized in the limbus area and showed a horizontally elongated shape. In addition, migration into the hair cell area was increased, and the morphology of the MSCs changed to various forms after kanamycin treatment. In conclusion, the results of this study indicate that the coculture of MSCs with cochlear epithelium will be useful for the development of therapeutics via cell transplantation and for studies of cell regeneration that can examine various conditions and factors.

In this study, we present a real-time imaging method using confocal microscopy to observe cells moving toward damaged tissue by ex vivo incubation with the cochlear epithelium containing the organ of Corti.

To study the effects of mesenchymal stem cells (MSCs) on cell regeneration and treatment, this method tracks MSC migration and morphological changes after ...

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 ...