I’ve been looking at several different protocols for antibody immunostaining in worms, and it seems many of these protocols involve performing all the fixation and staining steps on poly-l-lysine slides.
I am only interested in looking at embryos, and I was wondering if it really is necessary to perform all these washes on slides. Since I am working with embryos, I was thinking it would be more practical to fix them in formaldehyde, freeze crack them, then perform the majority of the staining and wash steps in microfuge tubes.
Does anyone have a good standard protocol for doing antibody staining in microfuge tubes?
Someone with more experience staining embryos may chip in.
My advice would be to bleach worms and rinse and collect the embryos in small tubes (0.5 ul), discarding any corpses.
First try the Bouin’s tube fix protocol from WormBook: http://www.wormbook.org/chapters/www_immunohistochemistry/immunohistochemistry.html#sec2_8
Check with DAPI and some known antibody (anti-DNA, anti-histone, etc.) for good permeabilization.
If you have any abundant epitope, you may not need a collagenase treatment.
If you do not get good staining, you can add collagenase treatment of the embryos before the first antibody incubation.
Our lab routinely used Tube staining for our staining needs.
The protocol you provided seems standard. They don’t freeze crack because they are using the Bme to get through the cuticle.
For our tube staining, we just do our fixing, staining, and washes in the tube.