Hi
I am recently doing Co-immunoprecipitation experiment. And I can always find two bands at 35kD and 40kD (I use rabbit polyclonal antibody as primary antibody of WB) which are strong enough and prevent me from checking whether my target Co-IP protein exist at 37kD. I used protein a/g plus agarose beads of Santa cruz (sc-2003). To find the reason, I boiled the beads in loading buffer (with dtt, beta ME, sds) directly after washing with PBS-triton-x. Strikingly, no matter what kind of primary rabbit polyclonal antibody I use, it always gives these two strong bands after WB. I searched online and find they would be protein A/G that come off after boiling. So I tried to elute my protein with loading buffer at RT, then I got supernatant, boiled then for western blot. However, these two bands still exist as strong as before! Now I think maybe I could try to elute with loading buffer without beta ME, dtt or I should try ph 2.5 glycine elution? Are they protein a/g and how do I eliminate them in CoIP experiment? You can see the files attached below, two red arrows point 43 and 34 kD respectively.
Hi there,
As you get the signal whatever the primary antibody, the problem seems linked to the reactivity of the secondary antibody. What are the dilutions? Do you saturate beads prior use? What is your WB protocol? I sounds weird to me that covalently conjuguated proteina/g would come off the beads and react in WB...
Dilution of my secondary is at 1:10000 in 5%milk. Though protein a/g is covalently linked to agarose beads，possibly some reducing agent like dtt can destroy the bond? Actually I dont know what these two bands are. But I am sure that they come from beads. By the way I dont understand "saturate beads". Before coip. i just wash my beads three times with pbs-tritonx
Well, I have used protein a/g agarose for coIP without having any problem with boiling/reduction treatment. About saturation nevermind I've come to read that your product is pretreated with BSA to saturate non specific sites on the beads.