Hello all,

I am attempting to compare degradation of a GPCR, specifically the Dopamine D2 receptor, with a mutant form of the receptor I have created, with extended agonist treatment. Both receptors are N-terminally FLAG-tagged.

These are HEK293 cells stably expressing either the D2 or mutant receptor, grown to confluency in 6-well plates and treated. To lyse the cells, I use 150ul ice cold RIPA buffer (with 1% Triton X-100 and 2% SDS), which I then shear through a 30g needle, sonicate and centrifuge at 4oC at 14,000rpm for 10 minutes. I then transfer the supernatant into an Eppendorf tube containing 4x sample buffer supplemented with 0.05M DTT. I do not at any point heat/boil the samples, as I know boiling GPCRs causes them to aggregate and makes them incapable of moving through the acrylamide gel.

When I resolve the lysates via SDS-PAGE, I seem to get very weak signal at the level I would expect (about 50kDa) and some weak bands just above this, which I assume is PTMs (glycosylation etc.). My biggest problem is that I get a lot of signal on my blots around the 100kDa and 120-ish kDa mark, which is indicative that my sample contains mostly receptor dimers. Due to the high concentration of detergent in my lysis buffer and reducing agent in the sample buffer, I'm unsure why I am still getting so much dimerisation. I have attached a picture of a representative blot I get, for reference. I use the M1 monoclonal antibody directed against the FLAG tag. Can anybody suggest what I might do to reduce the signal at these higher bands?

Kind regards,

Kyle