This article describes a method for detecting and isolating specific interacting proteins using bimolecular complementation affinity purification. The technique involves transfecting mammalian cells with plasmids encoding bait and prey proteins fused to complementary fluorescent protein fragments.
To detect and isolate a pair of specific interacting proteins using bimolecular complementation affinity purification, add a pair of bimolecular fluorescence complementation plasmids-transfection agent mixture into a culture plate containing mammalian cells.
One plasmid encodes one interacting protein — the bait protein — fused to a reporter fluorescent protein fragment. The other plasmid encodes for the other binding protein — the prey protein — fused to the fluorescent protein's complementary fragment.
Incubate. The transfection agent facilitates plasmid cellular uptake. Successfully transfected cells express cell membrane-localized proteins.
The bait-prey proteins' interactions bring the fluorescent protein fragments closer. This causes the fragments to fuse and refold, forming the functional fluorescent protein, whose fluorescence can be visualized under a fluorescence microscope.
Replace the media with a non-ionic detergent-containing lysis buffer to disrupt the cellular membranes, releasing the fusion proteins. Collect the cell lysate in a tube. Centrifuge.
Transfer the fusion protein-containing supernatant into a fresh tube. Add agarose beads conjugated with the fluorescent protein-targeting single-domain antibody, nanobody, which recognizes and binds to a three-dimensional epitope on the folded fluorescent protein.
Centrifuge. Resuspend the fusion protein-bound agarose beads in buffer. Heat to dissociate fusion proteins from the agarose beads. Perform SDS-PAGE to separate individual proteins, followed by western blotting with antibodies specific for the fluorescent protein fragments.
Only interacting proteins tagged with fluorescent fragments are detected, confirming their isolation.
First, seed HEK293T cells in 10-centimeter dishes containing 10 milliliters of DMEM. Then, dilute 2.5 micrograms of each bimolecular fluorescence complementation vector in 500 microliters of transfection buffer.
Add 10 microliters of the transfection reagent, and vortex the mixture for 10 seconds. Then, briefly centrifuge the samples, and incubate them at room temperature for 10 minutes. Next, add the DNA transfection mixture drop-wise to the dish. Then, incubate the samples for 8 to 24 hours.
Prepare cell lysis buffer and supplemented cell lysis buffer as outlined in the text protocol. Then, wash the cells with ice-cold PBS twice. Aspirate the PBS, and add 1 milliliter of ice-cold supplemented cell lysis buffer.
Next, place the dish on ice, and incubate for five minutes. Then, use a cell scraper to remove the cells and transfer them to a chilled microcentrifuge tube. Centrifuge the tube at 18,000 times gravity for five minutes at 4 degrees Celsius to remove the cellular debris. Then, transfer the clear supernatant to a fresh microcentrifuge tube.
Wash an appropriate volume of agarose beads in 1 milliliter of PBS. Then, centrifuge the beads at 300 times gravity, and carefully remove the supernatant. Next, add 20 microliters of agarose beads to each sample, and incubate at 4 degrees Celsius for two hours with end-to-end rotation.
Centrifuge the beads at 300 times gravity, and wash them in cell lysis buffer three times. Then, resuspend the washed beads in 50 microliters of diluted sample buffer. Finally, heat the samples at 95 degrees Celsius for two to three minutes.
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