Establishing Primary Mixed Glial Cell Cultures from a Mouse Spinal Cord

Working in a tissue culture hood, transfer four mouse spinal cords into a Petri dish of HBSS. Use sterile scissors and forceps to cut each of the spinal cords into small pieces. Then, transfer the pieces to a 50-milliliter conical tube containing papain DNase enzyme mixture. Avoid transferring HBSS to the enzyme mixture as this may result in decreased performance of the enzyme.

It is crucial that tissue pieces are well-digested to obtain a single-cell suspension. However, over-digestion will result in fewer viable cells. Each lab needs to perform pilot tests to determine the exact digestion time based on what works best for their cells.

Next, gently vortex the tube and then incubate at 37 degrees Celsius for one hour with orbital shaking at 150 rotations per minute. Following the incubation, vortex the tube, and then vigorously triturate the tissue using a 5-milliliter pipette to promote further dissociation. Then, transfer the cell suspension into a 15-milliliter tube, and centrifuge at 300 times g for 5 minutes at room temperature.

During the centrifugation, add 300 microliters of reconstituted albumin ovomucoid inhibitor solution to 2.7 milliliters of EBSS in a sterile tube and mix well. Then, add 150 microliters of DNase solution. Following the centrifugation, remove the supernatant and resuspend the cell pellet in the freshly prepared albumin ovomucoid inhibitor and DNase solution. Vortex well to break the cell pellet.

Then, add 3 milliliters of reconstituted albumin ovomucoid inhibitor solution without DNase to the cell suspension. Centrifuge the cells at 70 times g for 6 minutes at room temperature. After the spin, remove the supernatant which contains membrane fragments, and retain the pellet.

To remove myelin from the dissociated spinal cord cells, first, add 8 milliliters of room temperature 20% density gradient medium into the tube containing the cell pellet and vortex gently. Next, centrifuge the cells at 800 times g for 30 minutes at room temperature without breaking. Following centrifugation, carefully aspirate the top layer of debris, which contains mostly myelin, and the supernatant leaving the pellet.

To remove any remaining density gradient, wash the cells by resuspending the pellet with 8 milliliters of cDMEM diluted with HBSS. Centrifuge the cells at 400 g for 10 minutes at 4 degrees Celsius. Remove the supernatant and wash the cells with diluted cDMEM as before.

After removing the supernatant, the pellet can be stored on ice until seeding the cells. Once ready for plating, resuspend the cells in 14 milliliters of cDMEM supplemented with 2-Mercaptoethanol, and add 1 milliliter of the cell suspension to each well in a 12-well plate.

Plate extra wells that can be used to determine the average cell number per well, and the microglial content of the culture. Once the cells have been plated, incubate the cells at 35.9 degrees Celsius with 5% carbon dioxide. Change the medium on day 1, which is the day after plating.

Some cells are attached to the culture plate, but they are still mostly round. There are also many floating cells and significant debris. Repeat the media change three to four days thereafter.