Isolation of Brain and Spinal Cord Mononuclear Cells Using Percoll Gradients

Preparation of reagents

Prepare stock isotonic percoll (SIP), 4 ml per brain, by mixing 9 parts of percoll with one part of 10X HBSS without Ca++ and Mg++.

Prepare SIP at 70% in 1X HBSS without Ca++ and Mg++, 2 ml per brain dispensed into a 15 ml polypropylene conical tube.

Note: It is recommended that percoll should be used at room temperature, if used cold, cells tend to clump and cell separation is less efficient.

Tissue collection and homogenization

Anesthetize mice and perfuse through the left ventricle with ice-cold 1X HBSS. Dissect brain and spinal cord and maintain in RPMI without phenol red until all mice are sacrificed.

Place tissues in 7 or 15 ml dounce homogenizer containing 3 ml of RPMI, gently make a cell suspension with a loose-fitting pestle (A size) followed by a tigh-fitting pestle (B size) to further dissociate the tissues. Complete the volume of the cell suspensions to 7 ml with RPMI.

Gradient set up

Add 3 ml of SIP to the cell suspension to make a final 30% SIP

Slowly layer the 10 mL cell suspension on top of the 70% SIP. This is the most critical step of the procedure, use a pipette-aid set in the gravity mode avoiding mixing of the 70% and 30% solutions. A very clear flat line should be visible at the 70%-30% junction.

Centrifuge 30 min at 500G 18°C. Make sure centrifuge will stop with minimal or no brake so that the interphase is not disturbed.

Using a transfer pipet, gently remove the layer of debris from the top of the tube and collect 2.0-3.0 ml of the 70%-30% interphase into a clean conical tube containing 8 ml of 1X HBSS. Ensure that the percoll containing the interphase is diluted about three fold, mix a few times by inversion and centrifugre 7 min at 500G at 18°C.

Resuspend pellet in 1 ml of cell staining buffer and transfer it to a 1.5 ml tube and wash one more time using a micro-centrifuge at 10,000G 1 min at 4°C.

Antibody staining

Resuspend pellets in 50 μl of purified antimouse CD16/CD32 diluted 1:200 in cell staining buffer to block the Fc binding sites. Incubate over ice for 10 min. Count cells using a hemocytometer.

Add 50 μl of antibody cocktail mix and incubate on ice for 30 min. Each antibody must be first titrated to assess optimal dilution. Use appropriate fluorochrome combinations chosen according the capabilities of laser and filter settings of your particular flow cytometer.

Wash cells in cell staining buffer, resuspend pellets in 100-200 μl of cell staining buffer and analyze immediately in the cytometer, or alternatively cells can be resuspended in 2% paraformaldehyde (PFA) prepared in PBS and stored over-night at 4°C.

Representative Results:

After spinning the gradients, the 70%-30% interphase should have a defined white halo, and quantitation of the cells recovered from a naíve mouse should yield 3-5 x10⁵ cells per mouse (brain plus spinal cord). Inflamed brain might have ranging numbers of inflammatory cells depending of the CNS insult or disease model (Figure 1).

Figure 1. Isolation of mononuclear cells, from naíve and EAE-brain at peak disease. Brain cells suspensions were separated over discontinuous 70%/30% percoll gradients. Cells stained were incubated with Fc-block followed by anti-CD45 antibodies for 30 min on ice. Cells were rinsed in cell staining buffer, and fixed in 2% PFA prior acquisition in an LSR-II. CD45 intensity is measured in the Y-axis, where three distinct populations are visualized. Further characterization of the immune phenotype of CD45^hi infiltrating cells is implemented using additional antibodies and subsequently analyzed by flow cytometry.