I am growing human brain endothelial cells (hCMEC/D3) on transwell membranes and due to limited amount of reagents need to process the transwells for immunocytochemistry after my experiment. To do this I have been fixing my cells in 4%PFA and cutting the membranes out with a scalpel then performing ICC as per usual. I am having difficulty getting good clear images of ZO-1 staining though and wondered if anyone had any optimised transwell ICC protocols for ZO-1 or other TJ proteins or suggestions to make it better?
Thanks
Years ago I had done this. The key is that you should ensure that you have conducted a broad assessment of primary antibody concentrations (titrations) and that you use will fixed tissue, incubation with the primary antibody for 48h, and I like to recommend staining with ABC immunoperoxidase methods using NiDAB as the chromogen. Here are the protocols. The key is rinse, rinse, rinse since any residual reagents will mess of the clarity of staining. Staining times are 12-15 minutes! if the primary antibody is correct you should get no background staining in that time.
One more comment. The paper I attached used freely floating fixed tissue. I found that cultures can use the same protocol. Having many many rinses really improved staining of cells in culture. What may not be obvious is that when tissue is attached to a well, it may be more difficult to get all the reagents in and out. That is why using longer incubation times and multiple rinses really works. Use of fluorescence without knowing how the antibody will function with peroxidase-based staining can make it more difficult to achieve optimal staining. See my comparisons of techniques in the paper.
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