I am trying to optimize an EMSA to look at DNA binding by NF-kB. I started with 293T cells treated with TNF for either 30 or 60 minutes. I extracted the nuclear fraction using high salt buffer (420mM) with glycerol and checked for cytoplasmic contamination by western prior to performing the EMSA. The protein concentration of each sample was normalized using high salt buffer (i.e. the salt concentration in the binding reaction for each sample is the same). 5, 2.5 or 1ug of protein was incubated in binding buffer (20mM Tric-Cl, 50mM NaCl, 1mM EDTA, 10% glycerol, 0.1% NP-40, 1mM DTT) + 1ug poly(dI:dC) + protease inhibitors for 10 minutes at room temperature. Subsequently, 1ul radiolabeled probe was added and the reaction was incubated at room temperature for 15 minutes. The sample were loaded onto a 5% acrylamide gel (29:1 ratio) made with TGE buffer and 10% glycerol, which was prerun at 25mA for 1 hour prior to loading the samples. The gel was run until Orange G dye (free probe, FP, lane only) was a few cm from the bottom. The gel was dried and expose to film on a screen at -80 for two hours.
I'm not super please with how it turned out. It look quite messy to me and the shift is very close to what appears to be a non-specific band. I suspect that I'm either loading too much protein (since I can only see the shift with 1ug protein) or using too much probe. Does anyone have any other tips for cleaning this up?
Edited by DrSnood, 27 June 2017 - 08:37 AM.