Rhodamine-Phalloidin Staining

Last updated 7/11/97 JAC

1. Rinse cells or tissue in PBS briefly to remove media components.

2. Fix cells or tissue in 1-2% paraformaldehyde in PBS (freshly prepared) for 15 min at RT. The time and concentration of fixative can vary, depending on the thickness of the specimen (time for fixative to diffuse into the sample) and on how sensitive the antigen may be to the fixative. For an unknown sample, it is worthwhile to vary the conditions widely in a pilot experiment.

Prepare 10% para. stock like this: 1g paraformaldehyde in 10 ml dH2O Heat to ~60°C Add 1 drop of 1M NaOH and continue heating until most is dissolved. Filter stock thru 0.2 µm filter

*** Phalloidin binding requires the F-actin to have a protein structure near native. Methanol or acetone used to fix and / or permeabilize essentially abolishes phalloidin binding. ****

3. Quench excess aldehyde with 10 mM ethanolamine in PBS (or 0.1 M glycine in PBS) for 5 min.

4. Permeabilize cells in 0.1% Triton-X100 in PBS for 1 min.

5. Incubate cells in Rhodamine-phalloidin (Molecular Probes) diluted 1:100 in PBS for 15 min.

6. Rinse 3 times in PBS, 5 min/wash.

(Alternative: For vertebrate cells, whose actin binds phalloidin very tightly, one can add Rh-ph to the final PBS wash at 1:1000 and then simply mount in that medium

7. Mount for microscopy using PBS-buffered 50% glycerol as mounting medium. Use an anti-bleaching agent, p-phenylene diamine or N-propyl gallate. (I would not use glycerol unless necessary for some other reason. PBS should be fine.)