It sounds like your main challenges are high background in unstimulated controls and excessive signal from the isotype control. One possible reason your unstimulated cells are showing the same phosphorylation levels as the stimulated ones is that bone marrow B cells might already have some baseline ERK activation, especially if they were exposed to cytokines during isolation. Resting them in low-serum media for longer, like overnight, is a good approach, but you might also try washing them with a mild phosphatase inhibitor to prevent any artifacts from dephosphorylation. Another thing to consider is your fixation timing—fixing immediately after stimulation can sometimes lock in phosphorylation from stress responses. You could try briefly washing the cells in PBS before fixation to see if that helps lower the background. Also, if the cells are at a high density, they might be signaling to each other and activating ERK even without stimulation, so lowering the cell concentration during resting might help.

For the high signal in your isotype control, one issue could be that polyclonal antibodies sometimes bind nonspecifically, so it might be worth testing a different isotype control, ideally from the same species and host as your primary antibody. If possible, switching to a fluorophore-conjugated pERK antibody could help rule out any problems caused by the secondary antibody. Blocking could also make a difference—adding an Fc-blocking step with anti-CD16/CD32 before staining might help reduce background. Another thing to try is switching your secondary antibody since some anti-rabbit secondaries can be quite sticky. If none of these work, you might want to consider using a directly conjugated primary antibody to remove the secondary step altogether. Let me know if any of these ideas sound promising or if you’ve already tried some of them. :)

Hi Attapong Sinkitjasub thanks for your reply and advice!

Just to let you know what I have /haven't tried:

I do rest my cells in low serum media and tested various timescales for this, when I leave overnight I find that there is a high proportion of dead cells and I would'nt say the phosphorylation results look much improved.

I always fix immediately after stimulation so I may try your suggestion of washing first. I will also try lowering the cell concentration in case that affects the activation/phosphorylation. I also do Fc block all my samples already.

I'm finding I get the same issue across multiple antibodies. I've also tried just assessing phosphorylation ex vivo straight away without any stimulation (as I suspected that IGF1 was for some reason not stimulating any activity) with a variety of antibodies to different phosphorylated species, and I again find that they all give the same level of signal, as if there is a systematic issue - as I find it hard to believe the phosphorylation levels would be the same for all the targets and all genotypes of mice I am testing?

The isotype control is the same species and host as my primary antibody, I've also tried a couple of antibodies from different vendors with the same result. I was suspicious that it was a sticky secondary, however my "secondary only" FMO control looks clean and entirely negative. I also did try a different fluorophore conjugated secondary and achieved the same results.

I would ideally use direct fluorophore-conjugated antibodies but the targets I'm interested in do not have a wide selection of available antibodies, and most seem to be unconjugated.

It seems to me there is a systematic issue I'm not aware of, as I continue to achieve the same results no matter if I adjust resting time, stimulation time, phospho-antibody etc. I just can't think what else to try and troubleshoot!

Thanks for your tips, I will try and see if delaying fixation and the cell density change improves anything at all. If you have any more ideas I am keen to hear any! thank you.