I am starting a new project using 300um slices obtained from postnatal rat brains. We're specifically interested in hypothalamic development and axonal growth. Nobody in my lab has used slice cultures before, so I'm trying to teach myself the technique.
After a few days in vitro, I fix my slices in 4% PFA for 1h at room temp, and then proceed to treat them like normal brain slices using our labs standard IHC techniques. The problem is that the slices aren't really holding together, and they readily disintegrate during processing. I have a suspicion that's because the slices were DOA (and perhaps fixing necrotic tissue does not make it any more durable). Is fragile tissue a sign of a failed culture?
My question above is probably so general that you wouldn't need to look at my protocol. But if it indicates a severe problem with what I'm doing, then it's worth having a look at what I do and seeing where the problem is.
I embed PND10 rat brains in low melting agarose, and generate 300um coronal sections through the diencephalon on a vibratome. I use chilled HBSS as my dissection medium. Slices are transferred onto millicell membranes in a 6 well plate. Here's the recipe I use for my medium:
DMEM (phenol-red free): 50%
I also add 1M HEPES (to 1.25%) and 100X pen/strept in the appropriate volumes.
Does this make any sense?
Edited by marmot, 17 December 2012 - 09:46 AM.