The protein I am interested in have transmembrane region as well as a soluble region. I am more interested in how the soluble region comes together for function. Would appreciate if anyone can provide suggestions on what methods to use?
You could potentially use fluorescent labels or SPR to monitor the binding of the proteins in real time. Are you able to express the soluble domains of the protein subunits as soluble proteins without the transmembrane parts? This simplified system would make the experiments easier, although you would have to balance that simplification against the concern about whether the results accurately reflect the interactions of the whole proteins.
Look up the BioID method - quite interesting, and it's been used for detection of mitochondrial interactions. Essentially, you tag your protein with a Biotin ligase from E. coli (BirA*) which biotinylates anything within close proximity (20-30nm, can be brought down through say the Apex method) which will fish out both strong but also potentially weak interactions (suggestive of protein complexes). The biotinylated substrates can then be purified through a streptavidin pull-down.
One drawback is you will end up with potentially 100-200 proteins to sort through. Though I could imagine that if you repeated the experiment multiple times, you could reduce background and fish out the higher frequency (stronger interaction) guys.
Here are some papers to get you going
Varnaitė, Renata, and Stuart A. MacNeill. “Meet the Neighbors: Mapping Local Protein Interactomes by Proximity‐dependent Labeling with BioID.” Ed. Lucie Kalvodova. Proteomics 16.19 (2016): 2503–2518. PMC. Web. 2 Aug. 2018.
Rhee, Hyun-Woo et al. “Proteomic Mapping of Mitochondria in Living Cells via Spatially-Restricted Enzymatic Tagging.” Science (New York, N.Y.) 339.6125 (2013): 1328–1331. PMC. Web. 2 Aug. 2018.
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