Hi there,

I am trying to optimise an immunoprecipitation experiment, where I am attempting to pull down protein X and see if protein Y comes with it. My problem is that I am detecting protein Y in my control IgG IP.

I am using the Pierce magnetic protein A/G beads, lysis buffer [TBS, 150mM NaCl, 1% Triton, protease and phosphatase inhibitors] and wash buffer [TBS, 500mM Nacl, 0.05% Tween]. Lysates are pre-cleaned with blank beads prior to IP. For the experiment I perform a negative control (beads + no antibody), a positive control (beads + rabbit IgG) and an experimental IP (beads + rabbit antibody against protein X). After the IP I perform three washes (1 min each), followed by a water wash, and elution in 60'C 1X laemmli.

When I probe the membrane with a mouse antibody against protein Y, I see a band of the correct molecular weight in the WCE lane and my experimental IP lane, but also in the rabbit IgG lane. The bead-only negative control is clear, so it is not binding to the beads or the tube.

I have validated that the band the protein Y antibody detects by WB (approx. 110 kDa) is specific, using siRNA knockdown. I am limited in that I cannot use another protein Y antibody for detection, so I need to try and fix the issue if possible.

Things I have tried:

• Increasing the Tween concentration in the wash buffer to 0.2% did not fix the issue
• Using other rabbit antibodies against eg. HA, Myc and irrelevant proteins all pull down protein Y.
• Other proteins (such as GAPDH and protein X) do not seem to be coming down with the rabbit IgG to the same extent. I can see very faint bands if I overexpose the membrane, but nowhere near as strong as the band I see for protein Y. Protein Y is particularly sticky it seems!

Has anyone encountered this problem before? Any suggestions on what to change to remove this non-specific binding to antibodies?

Thanks for any advice you can offer.