Cochlear Surface Preparation in the Adult Mouse

All research protocols involving male adult CBA/J mice at the ages of 10–12 weeks and C57BL/6J mice at the ages of 6–8 weeks were approved by the Institutional Animal Care and Use Committee (IACUC) at the Medical University of South Carolina (MUSC). Animal care was under the supervision of the Division of Laboratory Animal Resources at MUSC.

NOTE: For the procedures presented below, mice are anesthetized with ketamine (100 mg/kg) and xylazine (10 mg/kg) via intraperitoneal injections. Mice are decapitated after the animal no longer responds to painful stimuli, such as a toe pinch.

1. Extraction of Temporal Bones

1. Decapitate the mouse immediately postmortem with surgical scissors (17 cm long) and cut the skull bone with scissors from the posterior aspect forward along the center line of the skull after exposing the skull bone by pulling the skin anteriorly.
2. Remove the brain tissue using forceps and manually remove the temporal bones with the thumb and index finger for mice three months of age or older. Use small surgical scissors (11 cm long) to cut the temporal bone of mice younger than three months old.
3. Put the temporal bone into a Petri dish (30 mm in diameter) containing ice-cold fresh 4% paraformaldehyde (PFA) solution dissolved in phosphate-buffered saline (PBS), pH 7.4.
   NOTE: Prepare fresh 4% PFA solution and adjust the pH to 7.4 just before fixing the temporal bones, because improper pH balancing of the PFA solution will decrease the quality of the immunolabeling.

2. Fixation and Perfusion

1. Remove the stapes from the oval window and puncture the round window membrane with size #5 forceps. Under a stereo dissection microscope make a small hole into the apex of the cochlea using a 27 G needle connected to a 1 mL syringe.
2. Gently and slowly perfuse the cochlea with 4% PFA solution via the round and oval windows until the solution washes out of the small hole at the apex. Transfer the cochlea (one or two cochleae per vial) to 20 mL volume scintillation vials containing 10 mL of 4% PFA solution.
3. Gently agitate the scintillation vials at room temperature (RT) for 2 h and leave overnight in a refrigerator (4 °C) on a rotator.
   NOTE: The time of fixation can be adjusted depending on the antibody used. For example, limiting the fixation to 1.5 h will allow for successful immunolabeling of post synaptic terminals when using the GluA 2 antibody.

3. Decalcification of the Temporal Bone

1. Remove the PFA solution and wash the cochleae with fresh PBS 3x for 5 min each.
2. Add 20 mL of 4% ethylenediaminetetraacetic acid disodium salt (EDTA) solution (pH 7.4) to the scintillation vials and keep in a refrigerator for 48–72 h on a rotator with gentle agitation. Change the EDTA solution daily until the cochleae are decalcified. Check if the cochleae are decalcified by touching the bony vestibular portion with forceps to assess for elasticity or simply cut a small piece from the edge of the vestibular portion. If such cut results in a crushed piece, the cochlea is not decalcified.
   NOTE: The EDTA solution may interfere with the immunoreaction of primary antibodies depending on the solution concentration. Prepare 4% EDTA in PBS and adjust the pH to 7.4. For consistent decalcification of temporal bones, fresh EDTA solution is used.
3. Change the solution to PBS for microdissection after decalcification is complete.
   NOTE: If the cochleae are not sufficiently decalcified, dissection of the cochleae cannot be performed.

4. Microdissection of the Cochlear Epithelium

NOTE: Once decalcification is complete, microdissection of cochlear epithelium for immunolabeling needs to be performed as soon as possible. In a clean 30 mm Petri dish containing PBS, the inner ear is oriented in the following manner: in reference to the Petri dish’s top (lid) and bottom, the cochlear round and oval windows face the top. In reference to dissector, the cochlear portion is oriented toward the front (away from dissector) and vestibular portion toward the back (near dissector). The following describes the steps in detail.

1. Hold the vestibular portion of the temporal bone with forceps and cut the apical turn with the scalpel at a 45° angle as indicated by the red line (Figure 1a).
2. Cut vertically along the faint line between the round window and the oval window, as indicated by the red line (Figure 1b) to separate the cochlea from the vestibular portion (Figure 1c).
   NOTE: This cochlear portion contains the middle, basal, and hook portions of the epithelium.
3. Place the cochlear portion with the basal turn toward the bottom and the middle turn toward the top of the Petri dish and cut the bony capsule and lateral wall of the middle turn toward the end from which the apical section was removed, as indicated by the red line (Figure 1d,e).
4. Continue cutting to completely separate the middle portion from the basal and hook regions (Figure 1f,g).
   NOTE: The hook region of the cochlea is the end of the cochlear epithelium corresponding to sensitivity to tones 48 kHz and higher in mice.
5. Put the basal and hook portion with the basilar membrane site oriented toward the bottom of the dish and vertically cut the modiolus off the hook region to remove the modiolus as indicated by the vertical red line (Figure 1g).
   NOTE: The modiolus is the conical-shaped central axis of the cochlea that consists of spongy bone and cochlear nerve as well as the spiral ganglion. It may be preferable to perform this cut with scissors.
6. Cut the basal and hook regions as the other red line indicates in Figure 1g to separate the hook portion (Figure 1h).
   NOTE: The cochlea has now been separated into apical, middle, basal, and hook portions. The next steps will be the final dissection of each of the individual turns, using the middle turn as an example.
7. Cut away the relatively large portions of bony capsule and lateral wall tissue of the middle turn as indicated by the red line (Figure 1i).
   NOTE: The lateral wall of the cochlear duct is formed by the spiral ligament and the stria vascularis.
8. Hold the lateral wall with forceps to align the bony capsule and lateral wall with the bottom of the Petri dish and cut these tissues from the basilar membrane side. Then flatten the specimen to orient the sensory hair cell surface side up. Trim away the rest of the bony capsule and lateral wall (Figure 1j).
9. Remove the tectorial membrane using forceps to completely separate the middle region (Figure 1k).
10. Repeat steps 4.7−4.9 for the final dissections of the remaining turns or regions as shown in Figure 1l.
11. Prepare a 10 mm round coverslip for adhesion of the sensory epithelium. Use a pipette to hand-spread 0.5 µL of cell and tissue adhesive on the center of the round coverslip. After drying (3–5 min at RT), put the coverslips in PBS in Petri dishes.
12. Stick all four pieces of the sensory epithelium on the 10 mm round coverslip in the Petri dish. Then hold one edge of the coverslip to transfer it to a four-well dish for immunolabeling or immunohistochemistry (Figure 1m).
    NOTE: Figure 2 illustrates the location of the major cuts.

5. Immunolabeling for Cochlear Synapses

NOTE: The protocol for immunolabeling for cochlear synapses followed in this study has been previously described¹³. Presynaptic ribbons were labeled with a CtBP2 antibody (mouse anti-carboxyl-terminal binding protein 2 IgG1, labeling the B domain of the RIBEYE scaffolding protein). Post synaptic terminals were labeled with GluA2 antibody (mouse anti-glutamate receptor 2 IgG2a, labeling subunits of the AMPA receptor).

1. Wash the sensory epithelium with PBS 3x for 5 min each wash in a four-well dish and then add 2 mL of 2% nonionic surfactant (i.e., Triton-X 100) into the dish for 30 min at RT on a rotator.
2. Remove the nonionic surfactant solution from the dish using a pipette and add 100 µL of blocking solution containing 10% normal goat serum to each well for 1 h on a rotator with gentle agitation at RT.
3. Remove the blocking solution from each well, wash with PBS 3x for 5 min each wash under gentle agitation at RT.
4. Add 100 µL of the primary antibodies diluted with PBS to each well: mouse anti-GluA2 IgG2a (1:2,000), mouse anti-CtBP2 IgG1 (1:400). Cover each four-well dish with its lid and place into a large humidified container that protects from light. Incubate at 37 °C for 24 h.
   NOTE: As an option, rabbit anti-myosin VIIa (1:400) for immunolabeling sensory hair cells can be added.
5. Wash 3x with PBS for 5 min each wash with gentle agitation at RT.
6. Add 100 µL of the secondary antibodies Alexa 594 goat anti-mouse IgG1a (1:1,000), Alexa 488 goat anti-mouse IgG2a (1:1,000) diluted with PBS to each well. Cover each four-well dish with its lid and place into a large humidified container that protects from light. Incubate at 37 °C for 2 h.
   NOTE: If myosin VIIa is used, add Alexa 350 goat anti-rabbit (1:200).
7. Wash 3x with PBS for 5 min. Then, transfer the 10 mm round coverslip onto a slide with the samples on top.
8. Carefully add 8 µL of Fluoro-Gel with Tris buffer into the center of the coverslip. Then hold the edge of another 10 mm round coverslip with forceps to mount on top, sandwiching the two coverslips together.
9. Use nail polish to seal the slides, put them in a cardboard slide folder, and store in the refrigerator.
   NOTE: If some Fluoro-Gel leaks out between the coverslips during mounting, clean the edges of the coverslips before sealing the slide with nail polish. Confocal images need to be taken within 7 days.