Isolating Murine Primary Oligodendrocytes Using Immunomagnetic Beads

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

1. Mouse Brain Cortex Dissociation

1. Sacrifice postnatal 5-7-day old C57Bl/6N mouse pups by quick decapitation with scissors previously cleaned in 70% ethanol.    
   NOTE: Cerebral cortices from 5-6 neonatal mice pups were used for each preparation. On average, one mouse brain yields 1-1.5 x 10⁶ primary oligodendrocytes (Ols).
2. Cut the scalp skin along the midline with small dissecting scissors and retract to expose the skull.
3. Cut the skull carefully along the midline starting from the opening in the back of the skull towards the frontal area, lifting up with the scissors to avoid damaging the brain. Then, using the opening in the back of the skull cut toward each eye socket along the base of the skull.
4. Use fine forceps to gently tease the cortices away from the midbrain and transfer them to a 60-mm tissue culture dish containing 7 mL of B27NBMA. Repeat steps 2.2 through 2.4 for each brain, pooling the dissected cortices.
5. Transfer the cortices to a new 60-mm tissue culture dish containing 5 mL of dissociation buffer (B27NBMA, 20-30 U/mL Papain, and 2,500 U DNase I).
6. Dice the cortices into small pieces of about 1 mm³ using a #15 scalpel blade and incubate for 20 min in a 37 °C, 5% carbon dioxide (CO₂) incubator.
7. Add 1 mL of bovine growth serum (BGS) to stop the enzymatic reaction.
8. Transfer the cortices along with the dissociation media to a 15-mL conical tube using a 10 mL serological pipet.
9. Gently begin to dissociate the brain tissue by slowly pipetting up and down 6-8 times using a 10-mL serological pipet, using care to minimize bubbles.
10. Allow the tissue chunks to settle for 2-3 min and transfer the supernatant to a fresh tube.
11. Add 3 mL of B27NBMA containing 10% BGS/2500 U DNase I to the tissue pellet.
12. Use a 5 mL serological pipette to gently dissociate the brain tissue while pipetting up and down 6-8 times. Be careful to minimize bubbles.
13. Allow the tissue chunks to settle for 2-3 min.
14. Transfer the supernatant to a fresh tube. Add 3 mL of B27NBMA containing 10% BGS/DNase I to the tissue pellet.
15. Gently dissociate the brain tissue using a P1000 pipet tip, while pipetting the mix of media and brain tissue up and down. Repeat Steps 2.13 and 2.14 until no large chunks of tissue remain or until B27NBMA containing 10% BGS/DNase I is exhausted, whichever comes first. Be careful to minimize bubbles.
16. Pool cell suspension with previous supernatants.
17. Place 70-µm cell strainer in a 50-mL tube. Using a 10-mL serological pipet, pass the pooled cell suspension gently over the 70-µm cell strainer.
18. Wash the cell strainer by adding 1 mL of B27NBMA containing 10% BGS/DNase I.
19. Discard the cell strainer. Bring the volume to 30-mL with B27NBMA containing 10% BGS/DNase I.
20. Transfer the cell suspension to two 15-mL conical tubes. Centrifuge the cell suspension for 10 min at 200 x g.
21. Remove the supernatant without leaving the cells exposed to the air. The supernatant will be cloudy due to cell debris. Add 3 mL of B27NBMA containing 10% BGS to the pellet.
22. Carefully dissociate the cell pellet using a P1000 pipet. Bring the volume to 15 mL with B27NBMA containing 10% BGS.
23. Pass the cell suspension over a fresh 40 µm cell strainer.
24. Wash the cell strainer by adding 1mL of B27NBMA containing 10% BGS. Discard the cell strainer.
25. Bring the volume to 30 mL with B27NBMA containing 10% BGS.
26. Transfer the cell suspension to 2 x 15-mL conical tubes. Centrifuge the cell suspension for 10 min at 200 x g.
27. Remove most of the supernatant without leaving the cells exposed to the air. Resuspend the cell pellets in 5 mL of ice cold magnetic cell sorting (MCS) buffer.

2. Determination of Cell Count and Viability

1. Dilute 100 µL of the cell suspension with 400 µL of Trypan Blue Solution in a 1.5-mL microcentrifuge tube to achieve a 1:5 dilution in 0.4% (w/v).
2. Center a cover glass over a hemocytometer chamber and fill the two chambers with 10 µL of the cell dilution using a P10 pipette and avoiding overfill. The solution will pass under the cover glass by capillary action.
3. Place the hemocytometer on the microscope stage and adjust focus using 40X magnification.
   NOTE: Cell counts are recorded using a hand-held counter in each of five squares (four corners and one center). Only non-viable cells absorb the dye and appear blue, while live and healthy cells appear round and refractive and do not absorb the blue-colored dye, allowing for determination of the number of viable and total cells per milliliter.

3. Isolation of O4⁺ Oligodendrocytes

1. Centrifuge the cell suspension at 200 x g for 10 min.
2. Carefully discard the supernatant by using vacuum aspiration, avoiding exposure of the cells to the air.
3. Resuspend the pellet in 90 µL of MCS buffer per 1 x 10⁷ total cells followed by the addition of 10 µL of anti-O4 beads per 1 x 10⁷ cells.
4. Mix the cell suspension and beads by gently flicking the 15-mL conical tube with the finger 4-5 times.
5. Incubate the mix for 15 min at 4 °C, flicking the 15-mL conical tube with the finger 4-5 timesevery 5 min.
6. Wash the mix by gently adding 2 mL of MCS buffer per 1 x 10⁷ cells to the tube. Centrifuge the mix at 200 x g for 10 min.
7. Discard the supernatant by carefully using vacuum aspiration, avoiding exposure of the cells to the air. Resuspend the cell pellet in 500 µL of MCS buffer for every 1 x 10⁷ cells.
8. Attach a magnetic separator to a magnetic separator stand. Attach a selection column to the magnetic separator and place a 40 µm cell strainer on top of the column. Place two 15-mL or one 50-mL conical tube below the separation column to collect the flow through.
9. Pre-rinse the 40-µm cell strainer and separation column with 3 mL of MCS buffer and let the buffer run through the column without letting it dry.
10. Add the mix of cell suspension and beads to the cell strainer and into the selection column.
11. Wash the 40 µm cell strainer with 1 mL of MCS buffer.
12. Discard the cell strainer and let the mix of cells, beads and buffer run through the column without letting it dry.
13. Wash the separation column 3 times with 3 mL of MCS buffer and 1 time with OL proliferation media.
14. Remove the separation column from the magnetic separator, quickly place it into a 15-mL conical tube, and immediately add 5 mL of OL proliferation media.
15. Place a plunger on top of the column and firmly push to flush out the labeled cells into the 15-mL conical tube.
16. Remove 100 µL of cell suspension to determine cell count and viability using Trypan Blue and hemocytometer as indicated in section 3.
    NOTE: Cell viability above 80% is considered acceptable to proceed with the culture of OLs, but viability greater than 90% is optimal.