I’m facing a big problem with microscopy and would like to hear your suggestions. My aim is to visualize gamma-H2A.X foci in hameatopoietic stem cells under the microscope. When I try this or any other intracellular stain using antibodies, such as Lamin A/C staining, I simply get extremely high background everywhere without any specific signal. I don’t see foci or structures of what I’m trying to visualize. With both gH2AX or Lamin all I see is a green nucleus with green everywhere else. I’ve attached examples of what I see.

I use 4% PFA for fixation. I’ve tried different ways of permeabilising (different Triton concentrations) as well as different mounting slides, for example I’ve attached cells to slides with wells in them, and I’ve tried attaching cells to cover slips and doing my washes in a well-plate, which turned out slightly better (lower background), but the problem persists. I still don’t see foci for g.H2A.X or a clear nuclear boundary for Lamin. All I see is green everywhere (I conjugate with FITC, but I also get a high background signal for Cy3 even without any Cy3 antibody or any antibody at all, the same for FITC/GFP).

I always use 4% paraformaldehyde to fix, for 10 to 15 minutes. Other people do this with no problems. The Triton concentrations I use vary from 0.1% to 0.3% depending on the step or antibody used. I’ve tried blocking with 1 to 2% BSA or 5% donkey serum. I have tried different concentrations of primary antibody as well as secondary antibody but nothing changes. Just signal everywhere. This problem is driving me mad.

I’ve done Phalloidin Alexa Fluor 647 and DAPI staining the same way, which works fine. I see red cytoplasm, blue nuclei against a nice dark background. It is my antibody stains I am having problems with. I am also using non-adherent cells and this can’t be changed.

Any help would be greatly appreciated.