In my recent experiments I have been trying to quantify physiological concentrations of PRMT1 (the protein I study) in mammalian cell lines using western blot analysis. I run known concentrations of PRMT1 (for a standard curve) alongside three different microgram amounts of HEK 293 T17 cell lysate on an SDS gel. I pre-run the 12% 0.75mm 10-well gel at 110V for 20 minutes. I load my samples and run at 100V until the samples reach the separating gel. Then I crank up the voltage to 180V. The entire time the SDS Gel apparatus is surrounded in a bucket of ice. Then I transfer to PVDF at 100V for 70 min while also on ice. I use a 1:1000 anti-PRMT1 rabbit antibody (in TTBS + 5% BSA) and incubate overnight at 4 degrees Celsius. Then i use a 1:10,000 secondary anti-rabbit-HRP antibody (in TTBS + 5% non-fat dry milk) and incubate at room temp for one hour. Every time I have done this the bands have had a strange dumbbell shape and I have tried searching western blot troubleshooting advice on how to prevent this... the only thing I can think of is that I may be transferring to PVDF too long.

I also do not know why I am getting higher bands in my HEK cell lysate. The HEK cell lysate samples do not seem to stack well in the gel and I am not sure why. I only load between 12 and 2 uL of sample. Any help would be greatly appreciated! I have been struggling for a while with this. Thanks!