Hello.
I'm having quite some trouble getting cells to adhere to my cell culture slides after fixation/permeabilization/washing/antibody incubation steps in my immunofluorescence protocol.
Firstly, I work with mouse embryonic stem cells which are relatively fragile and normally grow on "feeder" fibroblast cells for stability, attachment, and support. In our lab, when we do sensitive experiments like IF or RNA extraction, we "pre-plate" our ES cells to get rid of feeder fibroblasts (which have much faster attachment). So I do this preplating, and plate cells directly on glass slides coated with gelatin (not coverslips). I let them attach for at least 24h before starting the IF. Even though I start with slides which have a confluency of around 80%, when I get to imaging I see about 10-20% of the cells remaining, really just patches of cells that have managed to stick around.
I follow a general IF protocol, so I don't think it's necessary to list the details of all the steps here, but here are the important bits:
- I do the washes in coplin jars so this is quite gentle (I just dip the slides in and out of the jars; there is not aspiration or shearing pressure)
- I do the antibody incubations by pipetting the antibody mixture on coverslips and putting the coverslip on the slide (saves antibody). I then dip the slide into a PBS solution to remove the coverslip automatically without pulling it or sliding it across the layer of cells.
Is the problem that I need to let the cells grow on the slides for longer than 24h? If I let them grow for too long, mESCs tend to grow vertically and on top of each other, which is no bueno for IF imaging.
I don't know what could be going wrong, but I lose way too many cells and literally need to search for fields containing cells for hours during imaging.
Thank you!