Hi researchers,
I have been running a western blot setup optimisation with beta actin as a loading control protein. I am getting non-specific bands at the higher MW range in addition to my target beta-actin band.
These were my key steps:
1. Gel was made at 16.5% (as I was probing a smaller MW protein at 5kDa as well) and samples were run at 120V for 90 mins.
2. Transfer of gel was done at 10mA overnight in the cold room (4 degrees celsius).
3. Transfer done as per bio rad protocol, and following remove of PVDF membrane, blocking was done in 5% BSA in TBS-T for 1 hour and r.t., following which anti-beta actin was added in a 1:10000 ratio (antibody from protein tech, mouse monoclonal). But primary antibody was in 1% BSA in TBS-T.
4. Pri incubation was overnight in cold room as well.
5. Next day, after 3-5 washes in TBS-T (each was 5 minutes), secondary incubation with 1:20000 of anti-mouse antibody (diluted in 1% BSA in TBS-T as well.
Is it something to do with antibody dilution? I have read article that used even more dilute concentrations for beta actin (1:100,000 for eg.), and I am not sure that the higher concentration could have resulted in the extra bands. Or the fact that I incubated my antibodies in 1% BSA, as opposed to maintaining the BSA % used for blocking.
Kindly refer to the attached image for the blot.
If someone has experience in this aspect, it would be great to hear from you!
Thanks and Regards,
Mathangi