New member; and in the few hours I've had to explore this forum, I have to say I'm very impressed with the wealth of knowledge that's available here. This is a wonderful resource.
Anyways, to the point:
I, like many other struggling grad students, am having difficulty optimizing a co-ip.
I'm trying to confirm a specific interaction between Protein A (ProA, bait) and Protein B (ProB, prey) in washed platelets. ProB was identified by LC MS/MS of excised bands from an affinity ligand pull down using ProA. IF shows that ProA and ProB co-localize relatively strongly. The only thing that remain is getting a solid Co-IP to confirm the interaction.
Lysis Buffers I've Tried:
25mM Tris, 150mM NaCl, 1mM EDTA, 1% NP-40, 5% Glycerol
25mM Tris, 150mM NaCl, 1mM EDTA, 1% CHAPS, 5% Glycerol
Tyrode's Buffer + 2mM CaCl2 + 2mM MgCl2 + 10mM Glucose + 0.5% Triton X-100
The IP works consistently for each of these buffers
Input (In): Platelet lysate in lysis buffer
- Eluate: Mouse IgG
+ Eluate: Mouse monoclonal anti-ProA
In - +
x - x (these x's represent bands on a western, dashes represent a lack of one )
However, the Co-IP seems to pull out ProB in the negative eluate, and I can't figure out why
In - +
x x x
Initially, I figured it was idiosyncratic for ProB. But I've repeated the above experiment with the listed buffers to test Protein C and Protein D (both of which are known binding partners cited in many papers). Same results. I can't for the life of me figure why. It couldn't be possible that ProB, C, and D all bind non-specifically to mouse IgG?
Thanks in advance for the advice guys
Edited by vh04x, 26 August 2012 - 12:51 PM.