Split-BioID — Proteomic Analysis of Context-specific Protein Complexes in Their Native Cellular Environment

We provide a step-by-step protocol for split-BioID, a protein fragments-complementation assay based on the proximity-labeling technique BioID. Activated on the interaction of two given proteins, it allows the proteomics analysis of context-dependent protein complexes in their native cellular environment. The method is simple, cost-effective and only requires standard laboratory equipment.

The overall goal of this procedure is to probe the composition of protein complexes that specifically assemble around a pair of interacting proteins of interest using split-BioID, a protein fragment complementation assay that induces proximity dependent protein biotinylation within living cells. This method can help answer key questions in cell biology such as how protein complexes dynamically remodel to regulate different cellular functions. The main advantage of this technique is that it allows the easy identification of context-specific protein complexes in their native cellular environments and with very high resolution.

For the final mass spectrometric analysis, the following steps are to be performed in keratin-free conditions and all material and reagents should be as keratin-free as possible. After cloning the ORFs of two proteins of interest into the split-BioID plasmid, carrying out transient transfection then adding biotin supplemented medium according to the text protocol, use PBS to wash the cells twice. Add 1.5 milliliters of PBS to each plate and use a scraper to harvest the cells, then transfer the cells for each condition to a separate 15 milliliter tube and spin the tubes at 1, 200 times gravity and four degrees Celsius for five minutes.

Remove the supernatants and snap freeze the pellets in liquid nitrogen. Then store the tubes at negative 80 degree Celsius until further processing. To prepare cell lysates use one milliliter of RT lysis buffer to resuspend the pellets.

Then pass the cells through a 25 gauge needle 10 to 20 times. Next sonicate the samples. Then add 100 microliters of 20%Triton X-100 per 900 microliters of recovered sonicated lysate for a final concentration of 2%Add 2.3 milliliters of 50 millimolar Tris, pH 7.4, per milliliter of lysate to adjust the NaCl concentration to 150 millimolar for binding to streptavidin beads.

Then distribute the adjusted lysates into 1.5 milliliter tubes and centrifuge them at 16, 000 times gravity and four degrees Celsius for ten minutes. Transfer the supernatants to a 15 milliliter tube and keep 50 to 100 microliters as input material. To perform a streptavidin pulldown, for each condition transfer 200 microliters of streptavidin coupled magnetic bead suspension to a 1.5 milliliter tube.

Place the tubes on a magnetic rack and wait approximately one minute before removing the storage buffer. With one milliliter of equilibration buffer wash the beads by gently mixing. Then evenly dispatch the equilibrated beads in the necessary number of tubes and place them back on the magnetic rack.

After removing the equilibration buffer, resuspend each set of beads with equal amounts of the corresponding cell lysates. Then incubate the samples at four degrees Celsius on a rotating wheel overnight. The next day place the tubes on a magnetic rack.

Wait until the beads stick to the side of the tubes and transfer the supernatants to a 15 milliliter tube labeled as flow through. With 200 microliters of wash buffer one, resuspend the beads in each tube and combine each set of resuspended beads corresponding to one condition in 1.5 milliliter tubes. Use one milliliter of wash buffer one to wash the beads twice on a rotating wheel for eight minutes.

Then perform two washes each for wash buffers two, three and four. To make sure the wash buffer is completely eliminated following the last wash step, after removing most of the supernatant spin down the samples. Then put them back on the magnetic rack and remove the remaining buffer.

Add 30 microliters of elution buffer to the beads then incubate the samples at 98 degrees Celsius for fifteen minutes and immediately move the tubes to the magnetic rack. Transfer the eluted sample to a fresh tube and store the tubes at negative 20 degrees Celsius until further processing. For each input sample prepare a PAGE sample by mixing equal amounts of protein with the appropriate volume of 3X SDS loading buffer in a total volume of 28 microliters.

Then prepare PAGE samples by mixing five microliters of each elution sample with 2.5 microliters of 3X SDS loading buffer. Load the samples on an SDS polyacrylamide gel. Then proceed to electrophoresis and western blotting according to the text protocol.

To carry out SDS-PAGE for MS analysis add 6.25 microliters of 4X sample buffer to 18.75 microliters of each elution sample and run the samples on a four to 20%pre-cast SDS gel until they migrate two to three centimeters into the gel. In a 15 centimeter petri dish, stain the gel with colloidal Coomassie Brilliant Blue G250 stain. Use a clean scalpel to excise the whole lanes for each sample, excluding the streptavidin band, and transfer the excised bands to 1.5 milliliter tubes for MS analysis.

In addition to endogenous biotinylated proteins identified in a BioID, split-BioID experiment, the additional major bands observed by western blot are the fusion proteins that self-biotinylated, which indicates that the two tested proteins, in this example Ago2 and TNRC6C or Ago2 and Dicer interacted in the cells. In addition the results of the western blot indicate that having an NBirA*Ago2 fusion protein paired with CBirA*fusions to TNRC6C or Dicer is more efficient than the opposite combinations in which CBirA*Ago2 is paired to NBirA*fusions of the other two proteins. Moreover, the activation was specific, as none of the CBirA*fusions could activate the NBirA*GFP control fusion protein to appreciable levels.

When performing the isolation for the first time all the steps of the purification should be analyzed by western blotting. As illustrated here, binding to the beads should be almost quantitative and virtually no leak through should be observed in the washes. When the eluted material is run on a Coomassie stained gel after protein expression and induced biotinylation the strongest band observed runs at about 17 kilodaltons and corresponds to monomeric streptavidin.

The area of the sample lane above the streptavidin band up the loading well is typically excised for MS analysis. While applying this procedure to two given proteins it's important to test different combination of fusion proteins. Indeed we have often observed that the overall biotinylation efficiency can closely depend on which protein is appended to the N or C terminal BirA fragments.

It is equally important to add at least one negative control split-BioID experiments. For example, on a pair of interacting proteins that are not related to the pair of protein of interest. Following this procedure and mass spectrometry, computational analysis should be applied to score the identified peptides against the negative control experiments.

We use, for example the MaxQuant and Perseus software that were developed by the Cox and Mann labs in Munich, Germany.