By Santosh Patnaik, June 2004

Typical method for CHO cells

Incubation periods may be shortened to 15 minutes

Antibody concentrations will vary from case to case

Cells incubated in Hank's balanced salt solution (has glucose), HBSS with 2% BSA

1. Use 5x10e5 cells per '300-400 ul tube' - 1.5ml eppendorf tubes used.

2. 'Control' tubes with no primary antibody but only the secondary antibody. If primary antibody is conjugated to a fluorescent dye, then use a 'no antibody' tube for control. If doing multiple staining (say, two colors [antibodies] for two epitopes), there should be multiple control tubes, each with only one of the secondary antibody.

Steps 3 and 4 done in 15 ml tubes (cells for all 'tubes' together) or 1.5 ml tubes

3. Wash enough number of cells in PBS or HBSS before setting up the 'pre-incubation.'

4. Preincubate at 4 deg with head-on-head rotation for 30 min to 1 hr.

5. Add primary antibody and incubate as before.

6. Microfuge at 600g for 2 minutes at room temperature.

7. Resuspend pellet in 1 ml HBSS/BSA and repeat step 6.

8. Resuspend in 300-400 ul HBSS/BSA and add secondary antibody.

9. Repeat 6 and 7.

10. Resuspend in 300-500 ul HBSS without BSA, transfer to FACS tubes, put on ice, cover ice bucket for darkness and take for analysis.

Note - for viability staining, in order to gate out dead cells, one can add propidium iodide solution to final 1-2 ug/ml just before putting the tube on the FACS machine, or 7-AAD (actinomycin D) to final 0.5 ug/ml about 10 minutes before FACS analysis. 7-AAD is better than PI as its signal does not overflow into the FL2 channel.

See FACS Scan or Calibur using method.