Protocol for Indirect Mouse Cell Surface Staining of White Blood Cells

1. Carefully layer the blood over 3ml of Lympholyte-M (specific for mouse). Do not allow the layers to mix since the cells will be retained at the interface of the two layers.
2. Centrifuge at 500rcf for 30 minutes.
3. Remove cells from the white buffer coat layer which will probably appear to be a slightly diffuse layer of cells at the interface of the Lymphocyte-M and media.
4. Bring the cells to 8ml with the same type of media used in step 1 and centrifuge 10 minutes at ~180rcf to wash and pellet the cells.
5. Discard supernatant.
6. Resuspend the cell pellet in 10ml of the same type of media used in step 1.
7. Make 1ml of a 1:10 dilution of the suspension for step 10 and count cells using a hemocytometer or cell counter.
8. Centrifuge cells from step 10 for 10 minutes at ~180rcf.
9. Discard supernatant.
10. Resuspend cells at working concentration (typically 2 x 10⁶ / 100ul).
11. Block immunoglobulin Fc receptors (reduces non-specific staining) by incubation with antibodies against mouse Fc(gamma)II/III (CD16/CD32). Typical conditions are 10 ug/ml blocking antibody for 20 minutes at 4^oC. It is not necessary to wash the cells after the blocking step.
12. Incubate cells in cell staining buffer containing optimal amount of biotinylated monoclonal antibody (this should be titrated in your target population) for 45 minutes at 4^oC.
13. Wash the cells 2x in an equal volume of cell staining buffer.
14. Incubate cells in cell staining buffer containing optimal amount (around 0.6 ug/ml) of fluorescent streptavidin for 30 minutes at 4^oC.
15. Wash the cells 2x in an equal volume of cell staining buffer.
16. Analyze on cytometer.

Note: Cells should be kept away from light and at 4oC during as much of the procedure as possible. Remember to include negative, isotype, and blocking antibody controls!