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

NOTE: An overview of the method is shown on Figure 1.

1. Planning of the Cloning Strategy

1. Select two putatively interacting proteins to be tested.
   NOTE: Each of the two proteins will be fused to one split-BioID fragment: either NBirA* or CBirA*. As a negative control, the CBirA* fusion proteins will be tested with NBirA* fused to the green fluorescent protein (NBirA*-GFP). As an additional control, the NBirA* fusions may be tested alone, without cognate CBirA* fusion or in combination with a CBirA* fused to an unrelated protein. It is advisable not to use CBirA*-GFP as a negative control as it was shown to consistently lead to major background when combined to any NBirA* fusion protein⁹. The cause of this observation may be the expression level of CBirA*-GFP, much higher than any other CBirA* fusion proteins we have used so far, that may lead to significant reassociation with the NBirA* fragment.
2. Once two proteins are selected, check the literature to find whether both have already been successfully tagged in functional studies (for example as a GFP-tagged fusion protein).
   1. If such studies exist, note the position of the tag (at the N- or C-terminus) and use the same orientation to tag the proteins of interest with the split-BioID fragments.
   2. If no such study exists, plan constructs coding for both proteins tagged at the N- and C-terminus and plan an assay to test the functionality of the fusion proteins (for example, a rescue experiment in a cell line depleted for the WT protein).
      NOTE: When pairing two fusion proteins using the plasmids described in Figure 2 that encode long flexible linkers, the orientation of the fusion proteins (BirA* fragments fused upstream or downstream of the proteins of interest) generally does not matter. Indeed using FRB and FKBP as model proteins, it was shown that all four possible iterations (both fragments at the N-termini, both at the C-termini, one at the N-terminus and the other the C-terminus, and vice versa) yield comparable biotinylation⁹.
3. Design primers to amplify the ORFs of the two proteins of interest for cloning into the split-BioID plasmids. Pay attention that the translation frames are the same as the BirA* fragments. If using the plasmids described in Figure 2, use the restriction enzymes PmeI and PacI to construct the CBirA* fusion, and the enzymes ClaI and MluI for the NBirA* fusion.
   NOTE: The plasmids depicted in Figure 2 carry a bi-directional tet-responsive element flanked by F3/FRT recombination sites. This allows the regulated co-expression at roughly the same level of both fusion proteins from the same plasmid¹⁰, and the possibility to rapidly construct stable cell lines by recombination-mediated cassette exchange (RMCE). In these plasmids, NBirA* has a myc tag while CBirA* has a FLAG tag. The ORFs can of course also be cloned on individual plasmids with constitutive promoters.
   CRITICAL STEP: When testing two interacting proteins, it is recommended to compare NBirA* fused to protein 1 combined with CBirA* fused to protein 2, and NBirA* fused to protein 2 combined with CBirA* fused to protein 1. Indeed, it is frequently observed that one iteration works better than the other.

2. Cloning the ORFs of the Genes of Interest into the Split-BioID Plasmid

NOTE: In this example, two proteins that can be tagged at the N-terminus are considered. Four conditions will be tested and compared to non-transfected cells (Table 1).

1. Use polymerase chain reaction (PCR) to amplify the ORFs of the two proteins to be tested with the appropriate primers. Design primers that introduce the ClaI and MluI restriction sites for the NBirA* fusion proteins, and the PacI and PmeI sites for the CBirA* fusion proteins.
   NOTE: The PCR can be performed with any commercial, preferentially proofreading, DNA polymerase. Follow the standard protocol from the handbook and adapt it to the melting temperature of the primers and the length of the ORF to be amplified according to the manufacturer guidelines.
2. Subclone the first ORF
   1. Digest both plasmid (about 2 μg) and PCR-amplified ORF1 (to be fused to NBirA*) with ClaI and MluI. Perform digestion reactions using 1 μL of each enzyme, mixed with the appropriate volume of DNA and reaction buffer in a total volume of 20 μL in a 1.5 mL tube for 1 h at 37 °C.
   2. Run both digestion samples on a 1% agarose-TAE DNA gel. Excise the bands corresponding to digested plasmid and ORF1 with a clean scalpel and transfer to 1.5 mL tubes.
   3. Purify both bands using a standard DNA extraction kit.
   4. Ligate digested plasmid and ORF1 using standard reagents.
      1. If using the ligation kit indicated in Table of Materials, ligate 100 ng of DNA containing three to five time molar excess insert over plasmid in a total volume of 4.5 μL. Add 5 μL of 2x T4 ligase buffer and 0.5 μL of T4 DNA ligase from the ligation kit. Perform the ligation reaction in a 1.5 mL tube for 10 min at room temperature (RT).
   5. Transform into standard DH-5α E. coli competent cells (prepared according to Inoue's method¹¹).
      1. Mix 3 μL of the ligation reaction with 50 μL of competent cells in a 1.5 mL tube on ice and incubate for 30 min. Transfer the cells to a heat block set to 42 °C for 30-45 s and then incubate back on ice for 2 min. Add 250 μL of pre-warmed (37 °C) lysogeny broth (LB) medium and plate 100 μL of the cells on a pre-warmed (37 °C) ampicillin-containing LB-agar plate. Incubate the cells overnight at 37 °C.
   6. On the next day, pick four to six colonies and incubate them at 37 °C, 180 rpm, overnight in 3 mL of LB medium containing 100 μg⋅mL^-1 ampicillin in a 15 mL tube.
   7. Isolate the plasmids from the picked colonies using a standard DNA MiniPrep kit.
   8. Verify the correctness of the plasmids by Sanger sequencing using the Cassette 2 reverse primer (Table 2).
3. Once a plasmid containing the first ORF has been identified, subclone the second ORF in that plasmid following the same steps as step 2.2 but using the PmeI and PacI enzymes.
4. Sequence the resulting plasmids using the Cassette 1 reverse primer (Table 2).

3. Testing of the Fusion Proteins

NOTE: The following instructions are for the dual inducible expression plasmids (Figure 2) and HeLa-11ht cells, a subclonal HeLa-CCL2 cell line, stably expressing the reverse tetracycline-controlled transcription activator rtTA-M2 and containing a locus for RMCE¹². The growth medium for these cells is Dulbecco's Modified Eagle Medium (DMEM) containing 10% tetracycline-free fetal bovine serum (FBS). When using another cell type, exact seeding conditions and growth medium will need to be adapted.

1. Transient transfection
   1. Seed cells at a concentration of 1 x 10⁵ cells in 2 mL per well of a six-well plate the day before transfection and incubate the cells overnight at 37 °C, 5% CO₂ in a cell culture incubator.
   2. On the day of transfection, remove the medium of each well and replace with 2 mL of fresh medium.
   3. Prepare the four transfection reactions according to Table 1. For each well to transfect, add 6 µg of polyethylenimine to 3 µg of plasmid DNA in a 1.5 mL sterile tube and fill to 500 µL with DMEM medium without serum.
   4. Incubate each transfection mix for at least 5 min at room temperature before adding drop wise to each well. Incubate the cells overnight at 37 °C, 5% CO₂.
2. Induction and proximity-labeling
   1. The day after transfection, remove the medium and replace with medium supplemented with biotin at 50 µM to stimulate biotinylation and doxycycline at 200 ng⋅mL⁻¹ to induce the expression of the fusion proteins. Incubate the cells for at least 20 h at 37 °C, 5% CO₂.
      1. To make a stock solution of biotin, dissolve biotin at 50 mg⋅mL^-1 (correspond to ca. 200 mM) in 2 M ammonium hydroxide. Once it is completely dissolved, dilute it to 50 mM in 500 mM Hepes, pH 7.4, then adjust the pH to 7.4 with HCl. Aliquot and store the resulting 1,000x stock solution at -20 °C. Dissolve doxycycline at 10 mg⋅mL^-1 in 70% ethanol and store in a screw cap microtube at -20 °C in the dark.
3. Cell lysate preparation
   1. Wash the cells once with 1 mL of cold (4 °C) phosphate-buffered saline (PBS).
   2. To each well, add 100 µL of lysis buffer (50 mM Tris pH 7.4, 150 mM NaCl, 2 mM EDTA, 0.5% NP-40, 0.5 mM DTT and protease inhibitors).
   3. Harvest the cells with a cell scrapper and transfer to 1.5 mL tubes.
   4. Centrifuge the samples at 14,000 x g for 10 min at 4 °C to remove cell debris.
   5. Transfer the supernatants to fresh tubes and place on ice.
   6. Determine protein amounts with a Bradford assay.
4. SDS-polyacrylamide gel electrophoresis (PAGE) and Western blotting
   1. Prepare a SDS-polyacrylamide gel.
   2. For each cleared lysate, prepare a PAGE sample of 30 µg (minimum 15 µg) protein in a total of 30 µL of SDS-loading buffer. Then load equal amounts of each sample (20 µL/well) on the SDS-polyacrylamide gel and proceed to electrophoresis.
   3. After electrophoresis, transfer the fractionated proteins to a low fluorescence PVDF blot membrane using any standard protocol.
      NOTE: For analyzing protein biotinylation, a 10 min transfer time with the "high MW" program of a fast transfer semi-dry Western blotting device usually works well.
   4. After the transfer, block the membrane in 5% dry milk in PBS for 30 min at RT.
   5. Incubate the membrane for 30–60 min at RT with fluorescent streptavidin conjugate diluted 1:15,000 in PBS containing 2% BSA and 0.1% Tween-20.
   6. Wash the membrane three times, each for 10 min, with PBS containing 0.1% Tween-20, and then once more with PBS.
   7. Scan the membrane on a fluorescence scanner imaging system.
      NOTE:A typical Western blot is shown on Figure 3. The above procedure describes a fluorescent-based detection but a luminescence-based detection using HRP-coupled streptavidin works equally well.

4. Split-BioID for Proteomics Studies

CRITICAL NOTE: For the final mass spectrometric analysis, all the following steps are to be performed in keratin-free conditions, all material and reagents should be as keratin-free as possible.

1. Transient transfection
   1. The day before transfection, seed three to four 10 cm plates for each condition at a concentration of 8 x 10⁵ cells in 10 mL per plate, and incubate the cells overnight at 37 °C, 5% CO₂ in a cell culture incubator.
   2. On the next day, prepare a master transfection mix for each transfection condition: for three plates per condition, 36 µg of polyethylenimine, and 18 µg of plasmid DNA dissolved in 900 µL serum-free DMEM. Incubate each master mix for at least 5 min at RT.
   3. In the meantime, replace the medium of each dish with fresh medium, and then add 300 µL of the transfection mixtures drop wise to each plate.
   4. Incubate the cells overnight at 37 °C, 5% CO₂.
2. Induction and proximity-labeling
   1. The day after transfection, transfer the cells to 15 cm dishes. For each dish, remove the medium, wash the cells with 7 mL of PBS, add 1.5 mL of a trypsin-EDTA solution and incubate for 5 min at RT. Add 3.5 mL of growth medium to resuspend the cells and transfer the cell suspension to a 15 cm dish filled with 20 mL of growth medium supplemented with biotin at 50 µM to stimulate biotinylation and doxycycline at 200 ng⋅mL⁻¹ (final concentrations) to induce the expression of the fusion proteins.
   2. Incubate the cells for at least 20 h at 37 °C, 5% CO₂.
3. Harvesting and storage of the cells
   1. Wash the cells twice with PBS, then add 1.5 mL of PBS to each plate and harvest the cells with a scrapper.
   2. Transfer the harvested cells corresponding to one condition to a 15 mL tube and harvest them by centrifugation at 1,200 x g, 5 min, 4 °C.
   3. Remove the supernatants and snap freeze the pellets in liquid nitrogen, then store at 80 °C until further processing.
      NOTE: Alternatively, cells can also be detached by trypsinization, harvested in growth medium, transferred to a 15 mL tube and washed three time with PBS before freezing.
4. Preparation of cell lysates
   1. Resuspend the cell pellets in 1 mL of lysis buffer (50 mM Tris pH 7.4, 500 mM NaCl, 0.4% SDS, 5 mM EDTA, 1 mM DTT, 1x complete protease inhibitor) at RT. Pass the cells 10–20 times (five to ten strokes) through a 25 G needle.
   2. Sonicate the samples with a sonification device.
      NOTE: With the sonification device indicated in Table of Materials, the following program can be used: four cycles at high intensity, 30 s per cycle in a cold (4 °C) water bath. Any other sonification device is suitable but parameters might need to be adjusted accordingly.
   3. Add Triton X-100 to the recovered sonicated lysate to reach a final concentration of 2% (typically, add 100 μL of 20% Triton X-100 to 900 μL sonicated cell lysate), and then 2.3 mL of 50 mM Tris, pH = 7.4 per 1 mL of lysate to adjust the NaCl concentration to 150 mM before binding to streptavidin-coupled beads.
      Critical NOTE: Higher salt concentrations often result in much less efficient binding to the beads.
   4. Distribute the adjusted lysates in 1.5 mL tubes (ca. 1.1 mL in three tubes) and centrifuge them at 16,000 x g, 10 min, 4°C.
   5. Transfer the supernatants (ca. 3.2 mL) to a 15 mL tube, and keep 50–100 μL as INPUT material.
   6. Measure the concentration of each sample with a Bradford assay and use equivalent of 3 to 3.5 mg protein content for the streptavidin pulldown.
5. Streptavidin pulldown
   NOTE: An overview of the pulldown procedure is shown on Figure 4. It is almost identical to the original protocol published by Roux and colleagues². The first time the pulldown is performed, samples of flow through and wash 1–4 may be put aside for Western blot analysis to ensure the procedure worked properly (Figure 5).
   1. For each condition, transfer 200 μL of streptavidin-coupled magnetic beads suspension to a 1.5 mL tube. Place the tubes on a magnetic rack, wait until the beads stick to the side of the tubes (ca. 1 min) and remove the storage buffer.
   2. Wash the beads by gently mixing with 1 mL of equilibration buffer (50 mM Tris pH 7–4, 150 mM NaCl, 0.05% Triton X-100, and 1 mM DTT).
   3. Dispatch the equilibrated beads in equal amount in the necessary number of tubes (usually when starting with 3–3.5 mg protein content, the lysates from each condition can be dispatched in two to four 1.5 mL tubes) and place back on the magnetic rack.
   4. Remove the equilibration buffer and resuspend each set of beads with equal amounts of the corresponding cell lysates from step 4.4.7. Incubate overnight at 4 °C on a rotating wheel.
   5. On the next day, place the 1.5 mL tubes on a magnetic rack, wait until the beads stick to the side of the tubes and transfer the supernatants to a 15 mL tube labeled as Flow Through.
      NOTE: From now on, all the steps are performed at room temperature unless indicated otherwise.
   6. Resuspend the beads in each tube with 200 μL of wash buffer 1 (2% SDS in water), and combine each set of resuspended beads corresponding to one condition in 1.5 mL tubes.
   7. Wash the beads twice for 8 min on a rotation wheel with 1 mL of wash buffer 1.
   8. Wash the beads twice for 8 min on a rotation wheel with 1 mL of wash buffer 2 (50 mM HEPES pH 7.4, 1 mM EDTA, 500 mM NaCl, 1% Triton X-100, and 0.1% Na-deoxycholate).
   9. Wash the beads twice for 8 min on a rotation wheel with 1 mL of wash buffer 3 (10 mM Tris pH 8, 250 mM LiCl, 1 mM EDTA, 0.5% NP-40, and 0.5% Na-deoxycholate).
   10. Wash the beads twice for 8 min on a rotation wheel with 1 mL of wash buffer 4 (50 mM Tris pH 7.4, 50 mM NaCl, 0.1% NP-40).
   11. To make sure the wash buffer is completely removed after the last wash step, remove most of the supernatant, and then spin down the samples. Put them back on the magnetic rack, wait until the beads stick to the side of the tubes and then remove the remaining buffer.
   12. Add 30 μL of elution buffer (10 mM Tris pH 7.4, 2% SDS, 5% β-mercaptoethanol, and 2 mM Biotin) to the beads. Incubate at 98 °C for 15 min, then immediately remove the beads on a magnetic rack.
   13. Transfer the eluted sample to a fresh tube and store at -20 °C until further processing.
6. SDS-PAGE and Western blotting
   NOTE: Prior to mass spectrometry analysis, it is recommended to assess the success of the biotinylation and pulldown by SDS-PAGE and Western blot. If no biotinylation pattern is observed one may consider that either dox or biotin was not added to the medium or that one of the two stock solutions is compromised.
   1. For each input sample, prepare a PAGE sample by mixing equal amounts of protein sample with the appropriate volume of 3x SDS-loading buffer in a total of 28 μL. Prepare PAGE samples by mixing 5 µL of each elution sample with 2.5 µL of 3x SDS-loading buffer.
   2. Load the samples (25 µL/well for inputs, 7 µL/well for elution samples) on an SDS-polyacrylamide gel. Proceed to electrophoresis and Western blotting as described in section 3.4.
   3. If performing the pulldown for the first time, prepare PAGE samples for each step of the pulldown (Input, flow through, Wash 1–4, elution) by mixing 20 µL of each sample (Input, flow through, Wash 1–4) with 10 µL of 3x SDS-loading buffer or 5 µL (elution) with 2.5 µL of 3x SDS-loading buffer and proceed as on step 4.6.4 (Figure 5).
7. SDS-PAGE for MS analysis
   NOTE: To minimize potential keratin contamination, precast gels and commercial sample loading buffer may be used.
   1. Add 6.25 μL of 4x sample buffer to 18.75 μL of each elution sample and run the samples on a 4–20% precast SDS-gel until they migrate 2–3 cm into the gel.
   2. Stain the gel with colloidal Coomassie Brilliant Blue G250 staining¹³ in a 15 cm petri dish.
   3. Use a clean scalpel to excise the whole lanes for each sample, excluding the streptavidin band (running at ca. 17 kDa, Figure 6), and transfer the excised bands to 1.5 mL tubes.
   4. Send these samples to a proteomics facility for further analysis.
      NOTE: Typical MS results can be seen in reference 9 (Figure 3 and Figure 7, and supplementary tables). The whole datasets are available on the PRIDE repository (accession number PXD005005).