My protein is a secretory protein and I would like to find out what other proteins does it interact with (i.e. membrane proteins in the target cells). Ideally, I would like to screen for both strong and weak/transient binding. What is the best way to do it?
Methods that come to my mind:
1. Co-IP
2. Pull-down assay
3. Tandem affinity purification
4. Above coupled with cross-linking to find weak/transient binding?
5. Label Transfer Protein Interaction Analysis
What do you think?
Hi Garshasb Rigi . Weak protein-protein binding (Kd~100nM to 1uM) is difficult to detect in crude cell extracts. The quickest and easiest way I can think of is to do native PAGE of the extracts, blot and look for band shifts.
If you have an antibody against the bait protein (or if the bait protein is labelled, say, with biotin), you can incubate the bait with 35S-labelled cell extract and then do an immune precipitation (or the analogue experiment with labelled proteins). The antibody-bait-target complex is then captured with protein A agarose beads (or, better and cheaper, protein A expressing dead bacteria (Pansorbin)), washed and subjected to SDS-PAGE. Bound proteins are detected by autoradiography of the gel, nowadays usually with an electronic detector rather than X-ray film.
The difficult thing is to optimise the washing (RIPA-) buffer: the more denaturing detergents (SDS, cholate) are present, the cleaner your gels will be, but the stronger the bond between bait and target needs to be to survive the washing step.
apart from the Co-IP and pull down traditional assays, after finding the partners if possible, surface plasmon resonance or kinetics can be done to find affinity. This will validate the results. Also different acidic and basic pH combinations would answer the binding change under different environment. Thank you.
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