From Julian Guttman

USE GLOVES WHEN HANDLING PHALLOIDIN, it’s a poison

Purchase either ALEXA 488 (green) or ALEXA 568 (red) Phalloidin from MOLECULAR PROBES
It comes in methanol.  Store it at –20ºC (no need to aliquot it)

Methanol interferes with phalloidin’s binding to filamentous actin, so we will have to evaporate off the methanol prior to using it. To do that do the following:

Use a final dilution of 1:2
Therefore if you are going to need 40μl of solution to add to your slides, take 20μl of the phalloidin from the stock bottle in the –20ºC freezer, and quickly put it into a small GLASS test tube (they work better than plastic).  The phalloidin will try to get out of your pipette as you are doing this due to the methanol that is currently in it.

Next, evaporate off the methanol using air blowing along the inside of the test tube. And using the heat from your hand on the outside of the tube. This will take 2-5 minutes usually.

When completely evaporated down you will probably see an empty tube.  (I can’t stress enough how you need ALL OF THE METHANOL GONE for this to work well) 

Then add your 40 μl of buffer (PBS, TPBS/BSA, etc…) whatever buffer you like to the tube.
Vortex it well
Place this on ice until use
Add some of this solution to your slides (you don’t need much, just enough to cover the sample.
Place this on the bench-top for 15-20minutes (you’ll have to play with the incubation times depending on the intensity of labeling)  the longer it is on the sample, the more intense the staining
Wash very very well with either PBS or TPBS/BSA wash buffer, there is no such thing as over washing with this stuff
Add a drop of vectashield (or other anti-fade containing media)
Coverslip, nail polish
Look

NOTE: this will work on fresh or fixed tissue but do not fix with methanol!!!
It seems to work best on acetone treated samples as described on the staining protocol