Lyndall, you need to cross link the Ab and the surface Ag together
BEFORE the EtOH permeabilization. EtOH isn't really a fixative in the
sense that paraformaldehyde is. A protocol that can be used with 10^6
cells/sample in 12x75 mm tubes follows. There are similar protocols for
microfuge tubes. You may also find that you need to start with more or
less cells, depending on how many you loose in the washes.
(1) Stain cells with FITC-Ab as usual. You should probably do a
titration to determine the minimal working Ab concentration.
This will make the PI/FITC compensation easier.
2) Wash twice with PBS or media + 0.1% NaN3.
(3) Resuspend cells in 1 ml cold 1% paraformaledehyde in PBS.
Vortex immediately.
(4) Incubate 60 minutes on ice in the dark.
(5) Wash cells once for 5 minutes @ 1800 RPM, 4C.
(6) Pour off supernatant and add 1 ml cold 70% EtOH dropwise while
vortexing. This will minimalize clumping. The cells can be kept
for several days in the dark at 4C.
(7) Pour off EtOH and resuspend in 1 ml PBS or 50 mM citrate with
0.1 mg/ml RNase A. Incubate 5 minutes at room temperature.
(8) Add PI stock (1 mg/ml), mix and incubate 10 minutes in the dark
at room temperature. 10 or 20 ul should do the trick, for a
final concentration of 10 to 20 ug/ml. As in step 1, use the
lowest effective concentration for best compensation. If you can
keep the PI/cells ratio the same in all tubes, the G1 peak should
appear at the same fluorescence intensity for each sample.
(9) Analyze by flow cytometry with pulse processing/dublet
discrimination.
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