Isolating Vestibular Ganglion Neurons From a Mouse Inner Ear

All procedures involving sample collection have been performed in accordance with the institute's IRB guidelines.

1. Preparations

NOTE: The solutions and tools described in this section can be made well ahead of time and used on the day of dissection and recording.

1. Prepare Liebovitz media supplemented with 10 mM HEPES (L-15 solution). The following steps describe making 1 L of L-15 solution.
   1. Pour 990 mL of deionized H₂O into one beaker. Measure and add 2.386 g (10 mM) of HEPES. Add one bottle of 1 L of L-15 solution powder.
      ​NOTE: Powdered L-15 has a longer shelf life and takes up less storage space. Alternatively, purchase L-15 solution and supplement with HEPES. The latter option also has the option of using a phenol-free solution, which is useful for fluorescence imaging.
   2. Stir the solution on a stir plate. Using a pH meter, slowly add 1 N sodium hydroxide (NaOH) to obtain a pH between 7.34 and 7.36.
   3. Add deionized H₂O until the solution reaches a volume of 1,000 mL. Filter the solution using a 0.22 µm pore size membrane.
      NOTE: L-15 solution can be stored at 4 °C for 1-2 weeks. If using L-15 made with phenol-red, the solution will turn pink if too basic (pH > 7.4) or orange if too acidic (pH < 7.34).
2. Prepare the culture medium for vestibular ganglion neurons (VGNs). Follow the steps below to make 50 mL of culture medium:
   1. Add 47.5 mL of GluMAX-I minimum essential medium to a clean glass beaker.
   2. Measure and add 0.1194 g (10 mM) of HEPES. This additional buffer resists pH changes while the cells settle before being moved to the incubator.
   3. Add 2.5 mL of fetal bovine serum (FBS). This makes 5% by volume FBS in a total volume of 50 mL of medium.
   4. Stir the solution on a stir plate. Using a pH meter, slowly add 1 N NaOH to obtain a pH between 7.38 and 7.4.
   5. Add 0.5 mL of penicillin-streptomycin. Filter the solution through a 0.22 µm sterile filter. Store the solution at 4 °C.
   6. At least 30 min before use, transfer the culture media to a tissue culture flask with a vented cap. Place the flask with culture media in an incubator with 5% CO₂ concentration at 37 °C.

2. Fabricating trituration pipettes

NOTE: Trituration pipettes can be reused for multiple experiments. Flush the pipettes with ethanol, water, and L-15 solution before each use and clean them with water and ethanol after each use. Carefully store between uses.

1. To prepare trituration pipettes for cell dissociation, hold a glass Pasteur pipette to the flame of a Bunsen burner. Once the glass tip begins to melt, stretch the glass out to the desired tip diameter. Pull one end of the pipette away from the flame to create a small bend.
2. Bring the bent pipette near to a microscope. Using a scoring tile, score and break the glass at the desired diameter.
3. Pass the tip of the pipette over the top of the Bunsen burner flame. This will quickly polish the rough edges near the tip. Check to ensure that the tip is not sealed by the flame.
4. Repeat the process until four or five pipettes have been prepared with varying sizes.

3. Extraction of vestibular ganglion and plating of vestibular neurons

1. Preparation for dissection
   1. Prepare an enzyme mixture of L-15 solution with 0.05% collagenase and 0.25% trypsin. For example, add 0.001 g of collagenase and 0.005 g of trypsin to 2 mL of L-15 solution into a beaker, add a small magnetic stirrer, and stir until fully mixed. Set aside at room temperature.
   2. Prepare a commercially obtained guillotine by placing paper towels to the side and underneath the guillotine to minimize exposure of the benchtop and other surfaces to blood and other biological materials.
   3. Lay out large stainless-steel scissors, large forceps, and large spring scissors. Keep a large spray bottle with 70% ethanol handy for head cleaning.
   4. Fill a 100 mL beaker with L-15 solution and place it on ice. Oxygenate the L-15 solution.
   5. Lay out the spring scissors, surgical scissors, forceps, scalpel, and transfer pipette (see Table of Materials).
   6. Prepare a 60 mm Petri dish (for the gross dissection) and two 35 mm Petri dishes (for the cleaning/fine dissection of the ganglia). Fill the 60 mm dish with oxygenated L-15 solution using a 30 mL syringe with a 0.22 µm filter tip.
2. Euthanasia, head-cleaning, and hemisection
   1. Weigh the pups to compute the appropriate dosage of fatal-plus dilution to administer intraperitoneally (~0.01 mL per 10 g).
   2. Once a deep plane of anesthesia is reached, as assessed by a lack of response to the toe pinch, decapitate using the guillotine.
   3. Rinse the head thoroughly with 70% ethanol and then thoroughly with L-15 solution.
   4. Completely remove the skin from the skull. Bisect the head using large spring scissors by starting at the entry point of the spinal cord to the brain and making two cuts, the first cut through the top of the skull and the second through the bottom of the jaw. Finish bisecting the head using surgical scissors.
   5. Place both halves of the head brain side down in a 60 mm Petri dish filled with L-15 solution. Place the fresh tissue not currently being dissected on ice.
3. Extracting the superior vestibular ganglion
   1. Scoop out the brain using closed surgical scissors. Sever and remove the cranial nerve to completely detach the brain from the skull.
      NOTE: At this point, the otic capsule (shaped like the number 8) should be visible at the back of the head.
   2. Using forceps and starting at the back of the head, pull clear membranous material off.
   3. Cut the top of the skull out using surgical scissors. Remove excess tissue from the back of the head and neck to make the dissection area and otic capsule cleaner and easier to access.
      NOTE: Avoid making the dish too cloudy by discarding excess tissue throughout the procedure.
   4. Transfer the tissue to a second Petri dish with fresh L-15 solution. Locate the otic capsule and the auditory, superior vestibular, and inferior vestibular nerves. Cut away the auditory nerve and separate the superior and inferior ganglia using small spring scissors.
      NOTE: Although the somata of the vestibular nerve are found in both the inferior (thinner) and superior (thicker) ganglia, cells extracted from the superior ganglion have better survival.
   5. Using a scalpel, gently shave off the bony ridge to weaken the bony area under which the nerve dives into the bony capsule. Carefully remove debris with fine forceps, exposing the entire swollen portion of the ganglion.
   6. Use the fine spring scissors to cut and separate the ganglion from the peripheral nerve branch that is diving toward the utricle.
   7. Remove the superior ganglion using fine forceps, making sure not to pinch too close to the ganglionic tissue. Transfer to a 35 mm Petri dish with fresh L-15 solution.
   8. Before proceeding, pre-heat the enzyme solution: pour the enzyme mixture into a 35 mm Petri dish and place in a 37 °C incubator for 10-15 min.
   9. Clean the ganglion using fine forceps and small spring scissors by removing bone (which appears white and crystallized in structure), excess tissue, nerve fibers, and any other superfluous structures. Take care to minimize the removal of any ganglionic tissue.
4. Tissue dissociation and plating
   1. Transfer cleaned ganglia into the pre-heated enzyme solution and place it back in the incubator for 10 to 40 min. Enzyme treatment is complete once the tissue starts to break apart but remains in one piece. Overtreating the tissue with enzymes will result in complete dissolution of the ganglia before trituration.
      ​NOTE: The amount of time that the tissue undergoes enzymatic treatment depends on the age of the animal. For example, ganglia from P9 rats are treated with enzyme for 25 min, and ganglia from P15 rats are treated with enzyme for 35 min.
   2. Transfer the ganglia to the 35 mm Petri dish with fresh L-15 solution and incubate for 2-3 min.
   3. Transfer the ganglia to another 35 mm Petri dish filled with filtered culture media.
   4. Using a 200 µL micropipette, pipette a ~150 µL drop of filtered culture media onto a coated glass-bottom dish. Do not add too much solution, as it is important to maintain surface tension to form a bubble of solution that remains over the glass coverslip.
   5. Transfer the desired number of ganglia (one to four) to the coverslip.
   6. Draw a small amount of culture medium from the culture dish to rinse the trituration pipette with medium to prevent the tissue from sticking to the sides of the glass pipette. Triturate by gently and repeatedly passing the tissue through the pipette until the ganglia are sufficiently dissociated.
      NOTE: Do not overwork the tissue to attempt single-cell suspensions or allow the cells to crash to the bottom of the dish using excessive positive pressure. Also, avoid forming air bubbles, as this will reduce cell survival. Gentle trituration is the key to achieving successful cell survival.
   7. Let the cells rest for 5 min. Check under a light microscope to see if the cells have settled on the coverslip.
   8. Carefully place a culture dish into a 37 °C incubator for 12-24 h. Again, make sure to keep the bubble of solution intact.
      NOTE: When culturing for longer than 24 h, refresh the culture media daily. This can be a possible pause point.