Hi there,
I have a problem with a lot of non-specific binding in WB using AT8 (P-tau (Ser202, Thr205)) monoclonal antibody. The sample is mouse brain (hippocampal and cortical areas). Surprisingly, I don't have issues with other antibodies on the same membrane (after stripping), but AT8 always turns out like that. The protocol is the following:
1. After transfer block with 5% milk in TBS-T for 2 hours at RT on a rotating table.
2. Rinse for 10 min with TBS-T on a rotating table.
3. Apply primary antibody overnight at 4C.
4. Rinse 3X10 min with TBS-T on a rotating table.
5. Apply secondary antibody for 1 hour at RT on a rotating table.
6. Rinse 3x10 min with TBS-T on a rotating table.
7. ECL and imaging.
If anybody has suggestions about antibody optimization - please don't hesitate to share them!
Thank you!
Your protocol looks fine to me. Consider reducing the concentrations of the primary and secondary antibodies.
Hello Nina Semenova
AT8 is a phosphorylation-dependent anti-tau antibody which is used to identify specific amino acids that are phosphorylated in tau at both serine 202 and threonine 205.
So, try using BSA as a blocking agent instead of milk. Milk contains casein, a phosphoprotein that will bind to the primary antibody causing non-specific binding and high background.
For detection of phosphorylated protein by Western Blot you should use 5% BSA in TBST for blocking, primary antibody in 3% BSA in TBST and secondary antibody in 3% BSA in TBST.
Also, decrease your secondary antibody concentration. Use a dilution of 1: 15000 or 1: 20000 for the secondary antibody. This will reduce the non-specific bands. Your primary antibody dilution should be in the range of 1:1000 to 1:5000. You can keep your blocking time to 1-1.5 hrs. The rest of the protocol seems okay.
Good Luck.