for a while now I've been trying to optimize my ICC procedure, a technique with which I'm not that familiar. Basically, I'm starting with just trying to stain PC12 cells for bèta-actin. Colleagues of mine have tried this before, but as the PC12 cells are a semi-adherent cell line they tend to detach from the culture surface fairly easily during the washing steps. To avoid this as much as possible I've been trying some different fixation methods in the hopes of keeping my cells in place.
In general, I seed and culture my PC12 cells in collagen-coated 96-wells plates. One day post-seed, I differentiate them with NGF for three days. I've tried fixing them with 4% PFA, ice-cold ethanol and a mixture of 4% PFA/ethanol/acetic acid. I wash the cells with PBS, blocking is done with PBS containing a low concentration of Triton-X100 (between 0.1 and 0.3%) and Normal Goat Serum. The antibodies are diluted in PBS + Normal Goat Serum.
The first time I tried this procedure, the cells stayed in place nicely with all fixation methods but the overall staining of actin didn't go so well. The second time I tried I only used 4% PFA to fix with, as the fixative didn't make much difference the previous time, but in the end I discovered not a single cell had withstood the protocol this time - washed away completely, even though the protocol I used was basically the same, save for some small alterations (nothing that could have resulted in the disappearance of the cells).
I'm thinking of incorporating an extra fixation step after the primary antibody incubation, but I'm not sure to what extend this will prevent the cells from being washed away. Does anybody have any experience with ICC on differentiated PC12 cells?
Edited by SusieQ, 14 January 2014 - 06:58 AM.