I have been trying to get my ELISA running for a while now and at the moment don't know what else to try.
I am trying to detect Flotilin-1 in lysates of human monocytes.
Primary antibody is from cell signaling https://www.cellsignal.de/products/primary-antibodies/flotillin-1-d2v7j-xp-rabbit-mab/18634
Cells were lysed with 1% Triton. Cell lysates were put onto 96-well plate with high-protein-binding capacity (after cooking them for 5 minutes at 95°C) overnight at 4°C. After washing three times with PBST and blocking for one hour at 37°C using protein free blocking (https://www.thermofisher.com/order/catalog/product/37584#%2F37584) plate was incubated with primary antibody 1:25 in 1% BSA in PBST overnight at 4°C. After washing three times for 5 minutes with PBST secondary antibody was put onto the plates for one hour at 37°C. Secondary buffer is 3% BSA in PBST. So far I tried different secondary antibody concentrations (1:2500, 1:5000, 1:10 000). I then washed 5 times 5 minutes. TMB kit was used for detection.
The problem is, that only in the 1:2500 dilution I see any change of colour but the signal-to-noise ratio is pretty poor and I get only 1,5-fold change in my cell lysates vs background (defined as signal in wells with cell lysates treated with only secondary). In the higher dilutions I get very few signal.
As 1:2500 is quite a lot of secondary, I suspect that all the signal I see is just unspecfic binding of the secondary. My guess at the moment would be that the primary antibody is not working in ELISA, although I previously tested the antibody successfully in Western Blots.
Any help is appreciated