Dissection and Isolation of Murine Glia from Multiple Central Nervous System Regions

NOTE: All animal studies were authorized and approved by the Cleveland Clinic Lerner Research Institute Institutional Animal Care and Use Committee.

1. Prepare media and supplies for dissection

NOTE: All buffer and media recipes are provided in Table 1. This procedure is done under sterile conditions in a tissue culture designated biosafety cabinet.

1. Prepare and sterile filter mixed glia media (MGM). This can be done the day before and stored at 4 °C.
2. Dilute fibronectin at a 1:100 concentration with sterile H₂O. The volume of diluted fibronectin will depend of the number of pups and CNS region used for the experiment. Use one T25 flask for 1 cortex, one T25 flask for 2-4 cerebella, one T25 flask for 2-4 brain stems and one T25 flask for 2-4 spinal cords.
   NOTE: Combining tissue from the same region using the criteria listed above will allow adequate dissociation without altering the protocol described below. Combining more tissue than described in Step 1.2 will require further optimization of dissociation reagent concentrations.
3. Pipette 3 mL of diluted fibronectin into T25 flasks at room temperature and allow to sit for 2 min.
4. Aspirate the fibronectin and allow flasks to dry overnight with flask lid loosened.

2. Cortex, cerebellum, brainstem, and spinal cord dissection

NOTE: This procedure can be done on the benchtop and requires a dissection scope. Use strict aseptic technique for all steps of the procedure and minimize tissue exposure to the room air. Keep all media chilled on ice during dissection to ensure maximal tissue preservation. Alternatively, this procedure could be done in a hood that allows the use of an internal dissection scope.

1. Wipe all areas with 70% ethanol.
2. Pipette 10 mL of PBS/antibiotic solution (PAS) into a 10 cm Petri dish. Prepare one Petri dish for each pup to be dissected. Set on ice to keep the solution chilled.
3. In a disinfected tissue culture hood, pipette 9 mL of DMEM (with no additives) into 4 separate 15 mL conical tubes, one for each CNS region, replace tube caps, and put on ice. Place ice bucket with tubes within reaching distance from the dissection scope.
4. Place 10 cm Petri dish prepared in step 2.2 on disinfected dissection scope stage.
5. Anesthetize and euthanize mouse pup according to institutional protocol and remove the head by rapid decapitation with sharp scissors.
   NOTE: Postnatal day (P) 3-5 pups are used. Older pups have a more developed CNS and may not be an adequate source of expanding glial cells. Younger pups (P) 0-2 may yield a higher number of glia and can be used; however, spinal cord tissue is very difficult to dissect at this young age due to size.
6. Clean the pup’s skin using 70% ethanol.
7. Using fine scissors, cut the cutaneous layer along the midline of the head of the animal, starting caudally and moving rostral, until reaching the snout. Avoid cutting deeply into the skull to avoid any tissue damage.
8. Angling the head down, pull cutaneous layer to each side of the skull and using spring scissors, make an incision along the skull midline, starting at the foramen magnum, again cutting caudal to rostral.
9. With fine-tipped forceps, pull the skull halves to the right and left sides, exposing the cortex, cerebellum, and brainstem.
10. Once exposed, gently lift the brain out of the skull and into the 10 cm Petri dish prepared in Step 2.2. Ensure the brain remains undamaged to preserve anatomical structure, with the hindbrain attached.
11. Using fine, curved-tipped forceps with the point of the forceps facing upward, pinch off the cerebellum. Remove the meninges and place in designated 15 mL conical tube in ice bucket.
12. Ensure that the brainstem is directly ventral to the cerebellum and is visible after removal of the cerebellum. Remove it with fine-tipped forceps, remove meninges, and place in designated 15 mL conical tube in ice bucket.
13. Separate the midbrain from the cortex, remove cortical meninges, and place in designated 15 mL conical tube in ice bucket.
14. To remove the spinal cord, place the decapitated mouse pup in a supine position (lying face upward) with the severed vertebral column elevated towards the investigator.
15. Spray again with 70% ethanol.
16. Cut along the lateral sides of the vertebral column, in a rostral to caudal direction, through the rib cage until reaching the hind limbs. While cutting, push back the internal organs until the vertebral column is visible.
17. Cut along each lateral side of the vertebral column until it is isolated and place in the 10 cm Petri dish prepared in Step 2.2.
18. With ventral side up, using fine spring scissors, alternate cutting the right and left sides of each vertebrae until reaching the lumbar region to expose the spinal cord tissue.
19. Gently remove the spinal cord and the meninges with fine-tipped forceps under the dissecting microscope. Place in designated 15 mL conical tube in ice bucket.
20. Repeat steps 2.5.-2.19 for each pup, combining tissue from the same region to fit the criteria outlined in step 1.2 for each prepared T25 flask.
    NOTE: Remove the meninges as completely as possible. If a significant amount of meninges remain, the fibroblast-like phenotype of meningeal cells will outgrow and overwhelm the cell culture. Multiple spinal cords, of equal phenotype, can be combined in order to generate a specific cell culture. Take care to ensure that overcrowding of cells does not occur, which may lead to glial apoptosis and differential phenotypes.

3. Tissue dissociation

NOTE: All the following procedures are carried out in a sterile tissue culture designated biosafety cabinet using aseptic technique and sterile materials.

1. Add 1 mL of 0.05% trypsin containing 0.53 mM EDTA to each 15 mL conical tube of 9 mL DMEM and tissue to begin tissue lysis.
   NOTE: DMEM contains a high amount of calcium chloride, which can act as an inhibitor of trypsin. If dissociation is not complete following the outlined steps, Hanks’ Balanced Salt Solution without calcium or magnesium can be used. After trypsinization, the enzyme can be neutralized by adding a trypsin inhibitor, although calcium should be added back to the solution as it is a cofactor for DNase I, which is used in subsequent steps.
2. Triturate with a 10 mL pipette approximately 20x.
3. Transfer the cell suspensions to empty 50 mL conical tubes.
4. Incubate the solution at 37 °C, 5% CO₂ for 15 min, gently agitating the lysates after 8 min.
5. Add 5 mL of MGM and 200 µL (5 mg/mL) DNase I to each tube for a final concentration of 50 µg/mL.
6. Triturate each lysate with a 10 mL pipette 10x.
7. Let the cell suspensions sit for 3 min at room temperature to allow non-dissociated tissue to settle at the bottom of the tubes.
8. Transfer the cell suspensions to new 50 mL conical tubes, leaving behind the non-dissociated tissue.
   NOTE: The lysis and trituration steps described above significantly limits the amount of non-dissociated tissue.
9. Centrifuge the tubes at 300 x g for 3 min at 4 °C without brake.
10. Aspirate the supernatant and resuspend the remaining cell pellets in 5 mL of MGM.
11. Triturate the pellet with a 5 mL pipette 20x.
12. Plate the 5 mL cell suspensions on coated T25 flasks.
13. Incubate the cells at 37 °C, 5% CO₂ and change media initially after 24 h to remove cell debris.
    NOTE: Some protocols recommend an initial media change after 72 h. Optimization of this step may be required.
14. Perform a 100% media change with MGM every 48-72 h until cells are 80% confluent (approximately 5-7 days).
    NOTE: All media must be warmed to 37 °C before media changes.

4. Microglia isolation

1. Once mixed glia cultures have reached 80% confluency, prepare flasks for shaking by tightening lids and sealing with paraffin film.
2. To remove microglia from mixed glial cultures, secure flasks horizontally on an orbital shaker inside of a 37 °C incubator. Shake flasks at 15 x g for 1 h.
3. Remove media and pipette into a 15 mL conical tube. Rinse flasks twice with 3 mL warm MGM, adding wash to the 15 mL conical tubes. These are the microglia.
4. Add 5 mL of warm, fresh MGM to the culture flasks.
5. Reseal flasks with paraffin film, secure flasks horizontally on shaker, and shake at 15 x g at 37 °C for 15 h for separation of OPCs from astrocytes.
   NOTE: This step can be done overnight, but the 15 h shake time is critical as excess time may result in cell death.
6. Centrifuge the supernatant from step 4.3. at 300 x g for 3 min and culture microglia according to standard protocols¹⁶ or use for a biological assay.

5. Oligodendrocyte precursor cell isolation

NOTE: When plating OPCs following initial isolation, they must be plated on a poly-D-lysine-coated surface (sterile plate or cover slip). Prepare these materials prior to the completion of this section.

1. Following 15 h shake, remove the supernatant from flasks and plate on sterile 100 mm Petri dish.
2. Incubate the supernatant at 37 °C, 5% CO₂ for 30 min, swirling after 15 min to remove remaining microglia, as these will very quickly adhere to the dish. Non-tissue culture-treated Petri dishes may be used for this step.
3. Remove non-adherent cell supernatant, count, and plate on a poly-D-lysine-coated surface. Typically, 7,500-10,000 OPCs are plated/cm².
4. Incubate at 37 °C, 5% CO₂ for at least 1 h (up to 6 h), then gently aspirate 95% of media, and slowly add warm OPC Media, pipetting media against the wall of the well to minimize disruption of OPCs. Change media every 48 h until cells are ready for use.
   NOTE: It is critical that only one well is changed at a time. OPCs are sensitive and are especially intolerant to dry conditions. The addition of PDGF-AA in OPC media is to delay OPC maturation into oligodendrocytes. This factor may be excluded from culture media if the experimental focus is mature oligodendrocytes.

6. Astrocyte isolation

1. Following 15 h shake, remove supernatant and rinse flasks 2x with warm 1x PBS.
2. Add 4 mL of 0.05% trypsin containing 0.53 mM EDTA and incubate at 37 °C, 5% CO₂ for 5 min or until cells have lifted. To ensure astrocytes have lifted, visualize them using a standard wide-field microscope. Astrocytes will appear spherical following trypsinization.
3. Once astrocytes have detached, stand flask vertically and add 4 mL MGM. Triturate to mix.
4. Pipette astrocytes into a 15 mL conical tube, centrifuge at 300 x g for 5 min.
5. Resuspend astrocytes in Astrocyte Media and plate on fibronectin-coated surface as described in step 1.2. or use for biological assay.
   NOTE: 20 ng/mL murine fibroblast growth factor can be added during the first media change to help establish the astrocyte culture. Additionally, standard gelatin coating may be a low-cost alternative to fibronectin.

7. Identification and isolation of microglia, OPCs, astrocytes, and mature oligodendrocytes using immunocytochemistry

1. Once plated cells have reached appropriate confluency, gently remove media and fix adherent cells using 4% paraformaldehyde (PFA) for 10 min. Do this step in a biosafety cabinet.
   NOTE: When aspirating or adding solutions, pipette with gentle pressure to prevent cell detachment.
2. Slowly aspirate PFA then wash cells 3x with 1x PBS for 5 min.
3. Prepare appropriate blocking solution using 10% serum and 0.1% Triton X-100 in 1x PBS.
   NOTE: Serum source reflects the host animal in which the secondary antibody was raised. For example, if the secondary antibody is goat anti-rabbit, the appropriate blocking serum is normal goat serum. Likewise, if the secondary antibody is donkey anti-rabbit, the blocking serum should be normal donkey serum.
4. Add blocking solution until cells are completely covered. Block for 1 h at room temperature.
5. Prepare an antibody diluent solution (9 mL of 1x PBS, 0.01 g bovine serum albumin, 30 µL Triton X-100). Alternatively, antibodies can be diluted in the blocking solution described in step 6.3.
6. Dilute primary antibodies specific for the following in antibody diluent or blocking solution:
   1. Microglia: ionized calcium binding adaptor molecule 1 (Iba1) at a 1:250 dilution (2.4 μg/mL).
   2. OPCs: neural/glial antigen 2 (NG2) at a 1:200 dilution (5 μg/mL).
   3. Mature oligodendrocytes: myelin basic protein (MBP) at a 1:400 dilution (concentration is lot-dependent and optimization may be necessary).
   4. Astrocytes: glial fibrillary acidic protein (GFAP) at a 1:400 dilution(0.25 μg/mL)¹⁷.
      NOTE: Do not mix primary antibodies raised in the same species.
      NOTE: GFAP will reliably label white matter astrocytes. For grey matter astrocytes, an alternative marker may need to be used.
7. Incubate primary antibody overnight at 4 °C (with gentle agitation is preferable).
8. Wash 3x with 1x PBS for 5 min to remove the primary antibody.
9. Incubate cells with appropriate secondary antibodies at a 1:400 dilution in antibody diluent or blocking solution protected from light.
   NOTE: Secondary antibodies are conjugated to a fluorophore which may be interchanged depending on available microscope parameters. Once fluorescent secondary antibodies have been applied, protect from light as much as possible.
10. Incubate secondary antibody for 1 h at room temperature, limiting exposure to light.
11. Wash 3 times with 1x PBS for 5 min to remove excess secondary antibody.
12. To label nuclei, use a 1:1,000 DAPI/PBS solution and incubate cells for 5 min at room temperature in the dark.
13. Wash 3x with 1x PBS for 5 min in the dark.
14. To mount, apply mounting media and allow to dry in the dark before imaging.
    NOTE: Mounted slides can be stored at room temperature for 2-3 days. For long term storage, move slides to 4 °C. Image within one week for maximal signal. All representative images have been imaged on a confocal microscope; however, inverted fluorescent microscopes are also recommended.