I am running an EMSA with a myogenic TF (myogenin with N-terminal GST tag expressed in BL21 E coli) and its wildtype binding site (labelled with Alexa647 for visualisation).

I use 100 fmol binding site (and 1 pmol unlabelled DNA for competition reactions) and 7 µg crude protein (dialysed in PBS after protein extraction).

Binding reaction contains: 1X binding buffer (10 mM Tris, 50 mM KCl, 10 mM DTT, pH 7.5), 2.5% glycerol, 5 mM MgCl2 50 ng/µL poly(dI.dC), 0.05% NP-40

The reactions are set up and incubated at room temperature for 40 min then I add 5X loading buffer (compatible with fluorescence) and run the gel at 100V for 1.5 hours on ice. I pre-run the gel at 100V for at least 1 hour before loading the samples.

The gel is 5% (29:1) PAA gel 0.5X TBE with 2.5% glycerol, run in 0.5X TBE.

Attached is an image of my results, where PBC=positive binding control (wildtype binding site from promoter of Mef2C) and NBC= negative binding control (Foxa1 exonic sequence containing no binding site for MYOG).

I can see a band shift when the DNA is incubated with the protein but I am unable to get the shift to leave the well. I need it to enter the gel as I want to eventually cut it out and extract the DNA. Does anyone have advice for how I might improve the migration of the DNA-protein complex through the gel? I have tried running it on a larger gel for longer with no success. I have also tried running it at 4°C. I have also tried with purified myogenin (using the GST tag for purification) but then the protein does not cause a DNA shift.