Cultured mouse brain slices

[Zorndeig]

Does anyone here have experience with cultured brain slices?
I am working with 300 um slices from young mice (p3 – p8). Unfortunately, it is very difficult to detach them from the membrane and transfer them to an OTC-filled vessel after PFA-fixation and sucrose treatment.
Because my staining does not work, I considered trying acetone fixation instead of PFA, but as far as I have understood, this is normally used for specimens that are already sectioned and ready for staining? Do you think that acetone could work for ~ 300 um specimens as well?

Another idea would be trying to detach unfixed slices, fill OTC in the insert and freeze them in liquid nitrogen, but I have not yet tried this and presumably the outcome will be even worse…

[richard03]

You will have penetration issue even you used acetone fixed approach. If you have to use 300um sections, you may need to use strong permeabilization step (high cenc. of triton, etc), with longer incubation time and with increased temperature.

[Audiobeadz]

I too work with 300 um thick mouse brain slices, but from 9 PND animals.  How long do you culture for in the insert and what kind of membrane is it?  I have found that RT PBS works well to remove my brain slices that have been cultured for 24 hours.  I permeabilize with 1% PBS-Triton for 30-60 minutes although I've seen other protocols that use lower concentrations for longer periods of time which may keep your tissue more fully intact.  Otherwise try cutting the membrane out from it's plastic rim and moving the section that way.  I perform free float staining in a 12 well plate and have had good results but use PFA and usually lose a few sections when I mount which is a major concern for myself.  Hope you find something useful.