I am confused;
just can’t understand, when I stained adults with phaloidin, I detected puncta, like dense body, instead of filament;
did anyone have such experience?
Maybe you are accidentally staining cuticle? The cuticle is VERY sticky and looks spotty. It may be possible that you aren’t permeating the cuticle in a good way (acetone fixation).
Here is a protocol that worked well for me while I was a post-doc in the Ruvkun lab. Hope this helps.
Phalloidin Staining worms
by Michael Koelle
modified from Beth Bucher and Andrew Chisholm
Phalloidin binds to filamentous actin. This stain allows visualization of mainly of muscles, but some other actin structures are visible.
- Rinse worms off a plate with S basal, spin briefly in a clinical centrifuge, rinse once with more S. basal to remove bacteria.
- Put 10 ul of worms in an eppendorf, freeze in liquid nitrogen, then immediately place in a speedvac to lyophilize the worms (takes ~5 min). Add 3-4 drops ice cold acetone, wait 5 min, remove as much of the acetone as possible and air dry/speedvac off the remaining acetone. Worms can be stored this way prior to staining.
- Put 2 U fluorescein conjugated phalloidin (Molecular Probes, Inc. #F-432) in an eppendorf tube. Speedvac off the methanol to dryness. Add 20 ul “S mix” to dissolve the phalloidin, and add this to the dry worms.
S mix (1 ml)
743 ul dH20
250 ul 0.8 M Na phosphate pH 7.5 (recipe: 8.1 ml 1M Na2HPO4, 1.9 ml 1M NaH2PO4, 2.5 ml H20)
1 ul 1M MgCl2
4 ul 1% SDS
Let the worms stain at room temp (in the dark to prevent bleaching) for 0.5-1 hour.
- Wash the worms 2X in 1 ml of PBBT (PBS + 0.5% BSA+0.5% Tween-20).
- Resuspend the worms in ~20 ul 2 ug/ml DAPI in PBS (DAPI is kept frozen as a 1 mg/ml stock). Let the worms sit >5 min.
- For viewing mix a few ul of worm suspension with an equal volume of mounting solution (90% glycerol, 10% PBS, 1 mg/ml phenlyenediamine).
- Viewing: illuminating for DAPI fluorescence bleaches the FITC rapidly, so be careful.