Differentiating Immortalized Multipotent Otic Progenitors into Spiral Ganglion Neurons and Evaluating the Differentiation
Transcript
First, clean round glass coverslips by placing them in a sterile 100-millimeter plate, and submerging them in 70% ethanol. Gently agitate the coverslips to ensure they are covered and leave the dish at room temperature for 10 minutes. Next, rinse the coverslips three times in sterile 1x PBS and then once with sterile water.
Then, let the coverslips dry by exposing them to UV light in a tissue culture hood for 15 minutes. For immediate use, place one sterile coverslip into each well of a 24-well plate, and gently shake the plate to assure that all coverslips lie flat. Then, coat the coverslips by adding 0.25 milliliters of 1x PBS containing 10 micrograms per milliliter of poly-D-lysine to each well.
After the plate has incubated for one hour at 37 degrees Celsius, remove the poly-D-lysine solution and wash the wells three times with sterile 1x PBS. Once the liquid from the last wash has been aspirated, introduce 0.25 milliliters of 1x PBS containing 10 micrograms per milliliter laminin into each of the 24 wells. After incubating the plate overnight at 37 degrees Celsius, aspirate the laminin solution. Then, wash each well with 1 milliliter of 1x PBS three times, but leave the PBS from the last wash in the wells until the cells are ready to be plated.
To begin neuronal differentiation, count the cells from dissociated otospheres as previously described and then resuspend them in prewarmed neuronal differentiation medium. Proceed by seeding between 1 and 1.5 x 105 iMOP cells into each well of the plate containing the coated coverslips. Incubate the plate at 37 degrees Celsius with 5% carbon dioxide, replacing the neuronal differentiation medium every other day.
After seven days, aspirate the medium and fix the cells, which should be attached to the coverslips, by adding 4% formaldehyde into each well and leaving them for 15 minutes at room temperature. Then, remove the formaldehyde and wash the cells once in 1x PBS containing 0.1% Triton X-100. Next, remove the wash buffer and add blocking buffer consisting of 1x PBS containing 10% normal goat serum and 0.1% Triton X-100 to each well.
Leave the cells for one hour at room temperature and then remove the liquid and replace it with blocking buffer containing the appropriate dilution of Cdkn1b or Tubb3 antibody. Following an overnight incubation at 4 degrees Celsius, wash the cells once with 1x PBS containing 1% Triton X-100 and then proceed with immunostaining as previously described. After staining has been completed, perform a final wash by adding 1x PBS to the wells.
To prepare them for analysis, remove the coverslips to which stain cells are attached from the 24-well plate and place them onto mounting medium on a glass slide. After letting the mounting medium dry overnight at 4 degrees Celsius, acquire epifluorescence images as previously described.
