D.R.C is a Cancer Institute NSW Fellow and D.N.S was previously a Cancer Institute NSW Fellow. The research findings presented in this manuscript were funded by the Cancer Institute NSW (13/FRL/1-02 and 09/CDF/2-39), NHMRC (Project Grant GNT1052963), Science Foundation Ireland (11/SIRG/B2157), NSW Office of Science and Medical Research, Guest Family Fellowship and Mostyn Family Foundation. J.F.H. and R.S. were recipients of an Australian Postgraduate Award.

1. Plasmid Cloning
NOTE: To generate plasmid vectors with the V1 or V2 tags fused to either the N-terminus or C-terminus of the gene of interest, BiFC destination vectors have been deposited with Addgene [N-terminal tag: pDEST-V1-ORF (#73635), pDEST-V2-ORF (#73636). C-terminal tag: pDEST-ORF-V1 (#73637), pDEST-ORF-V2 (#73638)]. The gene/s of interest will need to be in specific recombination cloning compatible entry vectors (i.e., pDONR223 or pENTR221), without stop codons, to proceed with cloning. Many compatible clones are already available within various plasmid collections, including the Mammalian Genome Collection (https://genecollections.nci.nih.gov/MGC/).
<ol>
	<li>Perform a recombination reaction to insert the gene of interest between the attR sites of the appropriate BiFC destination vector:
	<ol>
		<li>In a 0.2 mL tube, add 150 ng entry vector, 150 ng BiFC destination vector, and 8 &#181;L TE buffer (pH 8.0).</li>
		<li>Add 2 &#181;L recombinase enzyme mix (see Table of Materials). Mix well and centrifuge briefly.</li>
		<li>Incubate the reaction for 1 h at room temperature or at 16 &#176;C overnight.</li>
		<li>Stop the reaction by adding 1 &#181;L Proteinase-K solution and incubating at 37 &#176;C for 10 min.</li>
	</ol>
	</li>
	<li>Transform heat-shock-competent cells (see Table of Materials) with the LR reaction product: 
	NOTE: Any basic strain can conceivably be used at this point, as long as it does not contain the Ccdb antitoxin, which would prevent the specific selection of plasmids containing the insert.
	<ol>
		<li>Thaw competent cells on ice and transfer 50 &#181;L of cells into a 14 mL round-bottomed polypropylene tube.</li>
		<li>Add 1 &#181;L of reaction product to the cells and gently mix. Incubate for 20 min on ice.</li>
		<li>Heat shock in 42 &#176;C water bath for 45 s. Immediately return to ice for 2 min.</li>
		<li>Add 1 mL of LB media and incubate with shaking at 37 &#176;C for 1 h.</li>
		<li>Plate the transformation onto a 10 cm agar plate containing Ampicillin (100 &#181;g/mL) and incubate at 37 &#176;C overnight.</li>
	</ol>
	</li>
	<li>Purification of BiFC plasmid.
	<ol>
		<li>To purify plasmid from an individual colony, add 100 mL of LB media to a large conical flask and add Ampicillin (100 &#181;g/mL). To screen multiple colonies, add 5 mL of LB media to 14 mL round-bottomed polypropylene tubes and add Ampicillin (100 &#181;g/mL).</li>
		<li>Using a sterile inoculation loop, pick a single colony from the agar plate. Place the inoculation loop into the LB media and mix briefly.</li>
		<li>Cover the top of the conical flask with aluminum foil and incubate overnight at 37 &#176;C with shaking.</li>
		<li>Produce transfection grade plasmid DNA using a standard maxiprep or midiprep plasmid DNA purification kit (see Table of Materials)10. Assess quality of DNA by absorbance spectra. 
		NOTE: DNA should have an A260/A280 ratio &#62;1.8 and an A260/A230 ratio &#62;2.0. At this point, it is recommended to screen multiple individual colonies to check for efficient expression of the insert and tag.</li>
	</ol>
	</li>
</ol>
2. Cell Culture and Transfection
NOTE: For transfection of the BiFC vectors it is important to achieve a high efficiency, and relatively homogenous transfection. The vectors will likely be compatible with any standard transfection reagent, and the transfection conditions should be optimized accordingly. To perform mass spectrometry, we usually culture cells within 10 cm dishes, although this can also be proportionally scaled down to smaller dishes or plates for experiments that require less material.
<ol>
	<li>Seed 1.0 &#215; 106 HEK293T cells in a 10 cm dish with 10 mL of DMEM media, supplemented with 10% FBS and Penicillin/Streptomycin (1:100).</li>
	<li>Dilute 2.5 &#181;g of each BiFC vector into 500 &#181;L of transfection buffer (see Table of Materials) in a 1.5 mL microcentrifuge tube.</li>
	<li>Add 10 &#181;L of transfection reagent (see Table of Materials).</li>
	<li>Vortex mixture for 10 s, then briefly centrifuge. Incubate for 10 min at room temperature.</li>
	<li>Add the DNA transfection mixture to the plate, dropwise. Incubate the cells for a sufficient length of time for the BiFC fusion proteins to interact and for Venus folding and fluorophore maturation, generally ~8 - 24 h. 
	NOTE: The peak excitation for Venus is 515 nm, and its peak emission is 528 nm, although this is readily viewed using a standard fluorescent microscope set up to visualize GFP fluorescence.</li>
</ol>
3. Sample Preparation
<ol>
	<li>Harvesting lysates
	<ol>
		<li>Prior to lysate collection, prepare Cell Lysis Buffer [50 mM Tris-HCl (pH 7.4), 150 mM NaCl, 1 mM EDTA, 1% (v/v) non-ionic detergent (see Table of Materials)]. Store at 4 &#176;C.</li>
		<li>Immediately prior to harvesting, supplement 10 mL of Cell Lysis Buffer with protease inhibitor and phosphatase inhibitor (see Table of Materials). Keep supplemented Cell Lysis Buffer on ice.</li>
		<li>Wash cells twice in ice-cold PBS. Aspirate the PBS and add 1 mL of ice-cold supplemented Cell Lysis Buffer. Place the dish on ice, ensuring the buffer is spread over the entire surface area.</li>
		<li>Incubate on ice for approximately 5 min, then use a cell scraper (see Table of Materials) to scrape the cells and transfer to a pre-chilled 1.5 mL microcentrifuge tube.</li>
		<li>Remove cellular debris by centrifuging the collected lysate at 18,000 &#215; g for 5 min at 4 &#176;C and transfer cleared supernatant into fresh microcentrifuge tubes. 
		NOTE: At this point the lysate can be used immediately, or stored at -80 &#176;C. It is also recommended to keep an aliquot of crude lysate so that the efficiency of transfection and the affinity purification can be validated.</li>
	</ol>
	</li>
	<li>Affinity purification 
	NOTE: The BiCAP isolation step is performed using the GFP nanobody conjugated to agarose beads (see Table of Materials).
	<ol>
		<li>Prepare the agarose beads by washing an appropriate volume (20 &#181;L per sample + 10 &#181;L excess) in 1 mL PBS. Centrifuge the beads at 300 x g and remove the supernatant.</li>
		<li>Add 20 &#181;L to each lysate sample.</li>
		<li>Incubate the samples for 2 h at 4 &#176;C with end-to-end rotation. 
		NOTE: At this point the samples can be prepared for either SDS-PAGE and western blotting, or analysis by mass spectrometry.</li>
	</ol>
	</li>
	<li>Preparation of BiCAP eluant for western blotting.
	<ol>
		<li>Centrifuge the beads at 300 x g and wash three times in Cell Lysis Buffer.</li>
		<li>Resuspend the washed beads in 50 &#181;L of appropriately diluted sample buffer (see Table of Materials) and heat the samples at 95 &#176;C for 2-3 min. 
		NOTE: Samples prepared in this manner can be stored at -20 &#176;C for several months.</li>
		<li>Perform SDS-PAGE and western blotting11 for both the V1 tag and V2 tags (see Table of Materials), as well as any other proteins of interest.</li>
	</ol>
	</li>
	<li>Preparation of BiCAP eluant for mass spectrometry. 
	NOTE: For analysis by label-free quantitative (LFQ) mass spectrometry it is recommended to prepare the samples in at least quadruplicate to ensure robust statistical power.
	<ol>
		<li>Centrifuge the beads at 300 x g and wash six times in Cell Lysis Buffer without detergent. This is necessary to remove both the excess protein and also the detergent that will interfere with mass spectrometry analysis. Proceed to the next step immediately, or store the beads at -80 &#176;C. </li>
		<li>Trypsinize beads in 60 &#181;L Buffer 1 [2 M urea, 50 mM Tris-HCl (pH 7.5), 5 &#181;g/mL trypsin]. Allow the beads to digest for 30 min at 27 &#176;C in a thermomixer shaking at 800 RPM.</li>
		<li>Briefly centrifuge beads, then collect supernatant and transfer to microcentrifuge tubes (see Table of Materials).</li>
		<li>Wash the beads in 25 &#181;L Buffer 2 [2 M urea, 50 mM Tris-HCl (pH 7.5), 1 mM dithiothreitol], in order to reduce the bound proteins.</li>
		<li>Pool the beads and supernatant into one microcentrifuge tube. Allow the digestion to occur overnight at room temperature.</li>
		<li>Alkylate the samples by adding 20 &#181;L of iodoacetamide (5 mg/mL in ultrapure water) to each sample and incubate in the dark for 30 min.</li>
		<li>Inhibit digestion by treating each sample with 1 &#181;L trifluoroacetic acid (TFA). This step will also acidify the samples in preparation for stage tipping.</li>
		<li>Construct C18 stage tips by stacking 6 layers of 1 mm solid phase extraction C18 (Octadecyl) membrane disks (see Table of Materials) into a 200 &#181;L micropipette tip. Prepare a single tip for each sample.</li>
		<li>Wet the stage tips with methanol, and equilibrate with 50 &#181;L 0.1% (v/v) trifluoroacetic acid (TFA), 80% (v/v) acetonitrile.</li>
		<li>Wash the tips with 50 &#181;L 0.1% (v/v) TFA.</li>
		<li>Load the acidified peptides onto the stage tips and then elute using 0.1% (v/v) TFA, 80% (v/v) acetonitrile in two steps.</li>
		<li>Evaporate the samples using a vacuum concentrator.</li>
		<li>If needed, store dried peptides at -80 &#176;C.</li>
	</ol>
	</li>
</ol>
4. Mass Spectrometry.
<ol>
	<li>Resuspend samples in 15 &#181;L of 5% formic acid, 2% acetonitrile (in ultrapure water).</li>
	<li>Carefully load 6 &#181;L onto an LC plate and place into a nanoLC HPLC system (see Table of Materials).</li>
	<li>Pack a 20 cm, 75 &#181;M inner diameter column with 1.9 &#181;m C18 stationary phase particles (see Table of Materials). Load 5 &#181;L peptide onto each column.</li>
	<li>Elute peptides using a linear gradient of acetonitrile at 250 nL/min over 140 min and introduce by nanoelectrospray into a Linear Trap Quadropole (LTQ) hybrid mass spectrometer coupled to the nanoLC HPLC system.</li>
	<li>Collect tandem MS data for the top 10 most abundant ions per scan over a 140-minute time gradient. Randomize the order of data collection and interchange with BSA run between each sample to minimize temporal bias.</li>
</ol>
5. Analysis
<ol>
	<li>Process raw MS data using default setting within MaxQuant software version 1.2.7.4 and analyze MaxQuant output using modified version of the Perseus statistical analysis workflow in the R software environment3. 
	NOTE: The statistical analysis workflow from this point is summarized in Figure 2. Briefly, LFQ intensities of proteins identified using MaxQuant are transformed and filtered followed by normalization and imputation. Interacting proteins of dimer pairs are identified by comparison with a Venus control, to exclude nonspecific background binders, with Student&#39;s t test and Benjamini-Hochberg correction for multiple comparisons. Data quality is confirmed by comparisons of individual histograms, multiple regression, and hierarchical clustering.</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57109/57109fig2.jpg" /> 
Figure 2: Outline of the statistical analysis workflow. Flow diagram of statistical analysis pipeline used to analyze the LFQ intensities of proteins identified from raw mass spectrometry data processed using MaxQuant. Green boxes: filtering, blue boxes: transformation/normalising/scaling, brown boxes: data quality control, yellow boxes: semi-quantitative exclusion/comparative analysis, grey boxes: statistical analysis. This figure is republished fromCroucher et al.3 Reprinted with permission from AAAS. <a href="//cloudfront.jove.com/files/ftp_upload/57109/57109fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<ol><li>Kolch, W., Pitt, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Functional+proteomics+to+dissect+tyrosine+kinase+signalling+pathways+in+cancer%5BTitle%5D)+AND+%22Nat+Rev+Cancer%22%5BJournal%5D)">Functional proteomics to dissect tyrosine kinase signalling pathways in cancer.</a> Nat Rev Cancer. 10 (9), 618-629 (2010).</li>
<li>Pawson, T., Kofler, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Kinome+signaling+through+regulated+protein-protein+interactions+in+normal+and+cancer+cells%5BTitle%5D)+AND+%22Curr+Opin+Cell+Biol%22%5BJournal%5D)">Kinome signaling through regulated protein-protein interactions in normal and cancer cells.</a> Curr Opin Cell Biol. 21 (2), 147-153 (2009).</li>
<li>Croucher, D. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Bimolecular+complementation+affinity+purification+%28BiCAP%29+reveals+dimer-specific+protein+interactions+for+ERBB2+dimers%5BTitle%5D)+AND+%22Sci+Signal%22%5BJournal%5D)">Bimolecular complementation affinity purification (BiCAP) reveals dimer-specific protein interactions for ERBB2 dimers.</a> Sci Signal. 9 (436), ra69(2016).</li>
<li>Cassonnet, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Benchmarking+a+luciferase+complementation+assay+for+detecting+protein+complexes%5BTitle%5D)+AND+%22Nat+Methods%22%5BJournal%5D)">Benchmarking a luciferase complementation assay for detecting protein complexes.</a> Nat Methods. 8 (12), 990-992 (2011).</li>
<li>Rossi, F., Charlton, C. A., Blau, H. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Monitoring+protein-protein+interactions+in+intact+eukaryotic+cells+by+beta-galactosidase+complementation%5BTitle%5D)+AND+%22Proc+Natl+Acad+Sci+U+S+A%22%5BJournal%5D)">Monitoring protein-protein interactions in intact eukaryotic cells by beta-galactosidase complementation.</a> Proc Natl Acad Sci U S A. 94 (16), 8405-8410 (1997).</li>
<li>Magliery, T. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Detecting+protein-protein+interactions+with+a+green+fluorescent+protein+fragment+reassembly+trap%3A+scope+and+mechanism%5BTitle%5D)+AND+%22J+Am+Chem+Soc%22%5BJournal%5D)">Detecting protein-protein interactions with a green fluorescent protein fragment reassembly trap: scope and mechanism.</a> J Am Chem Soc. 127 (1), 146-157 (2005).</li>
<li>Hu, C. D., Kerppola, T. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Simultaneous+visualization+of+multiple+protein+interactions+in+living+cells+using+multicolor+fluorescence+complementation+analysis%5BTitle%5D)+AND+%22Nat+Biotechnol%22%5BJournal%5D)">Simultaneous visualization of multiple protein interactions in living cells using multicolor fluorescence complementation analysis.</a> Nat Biotechnol. 21 (5), 539-545 (2003).</li>
<li>Kubala, M. H., Kovtun, O., Alexandrov, K., Collins, B. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Structural+and+thermodynamic+analysis+of+the+GFP%3AGFP-nanobody+complex%5BTitle%5D)+AND+%22Protein+Sci%22%5BJournal%5D)">Structural and thermodynamic analysis of the GFP:GFP-nanobody complex.</a> Protein Sci. 19 (12), 2389-2401 (2010).</li>
<li>Fegan, A., White, B., Carlson, J. C., Wagner, C. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Chemically+controlled+protein+assembly%3A+techniques+and+applications%5BTitle%5D)+AND+%22Chem+Rev%22%5BJournal%5D)">Chemically controlled protein assembly: techniques and applications.</a> Chem Rev. 110 (6), 3315-3336 (2010).</li>
<li>JoVE Science Education Database. <a target="_blank" href="https://www.jove.com/science-education/5062/plasmid-purification">Basic methods in cellular and molecular biology. plasmid purification.</a> J Vis Exp. , (2017).</li>
<li>Eslami, A., Lujan, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Western+blotting%3A+sample+preparation+to+detection%5BTitle%5D)+AND+%22J+Vis+Exp%22%5BJournal%5D)">Western blotting: sample preparation to detection.</a> J Vis Exp. (44), (2010).</li>
<li>Shearer, R. F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+E3+ubiquitin+ligase+UBR5+regulates+centriolar+satellite+stability+and+primary+cilia+formation+via+ubiquitylation+of+CSPP-L%5BTitle%5D)+AND+%22Mol+Biol+Cell%22%5BJournal%5D)">The E3 ubiquitin ligase UBR5 regulates centriolar satellite stability and primary cilia formation via ubiquitylation of CSPP-L.</a> Mol Biol Cell. , (2018).</li>
<li>Shannon, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Cytoscape%3A+A+software+environment+for+integrated+models+of+biomolecular+interaction+networks%5BTitle%5D)+AND+%22Genome+Res%22%5BJournal%5D)">Cytoscape: A software environment for integrated models of biomolecular interaction networks.</a> Genome Res. 13 (11), 2498-2504 (2003).</li>
<li>Schopp, I. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Split-BioID+a+conditional+proteomics+approach+to+monitor+the+composition+of+spatiotemporally+defined+protein+complexes%5BTitle%5D)+AND+%22Nat+Commun%22%5BJournal%5D)">Split-BioID a conditional proteomics approach to monitor the composition of spatiotemporally defined protein complexes.</a> Nat Commun. 8, 15690(2017).</li>
</ol>

The authors have nothing to disclose

BiCAP is a powerful method for isolating specific protein dimers while excluding the individual components and their competing binding partners3. BiCAP is based on adaptation of a fluorescence protein complementation assay called BiFC6. Existing methods, including BiFC and proximity ligation assays, have been used extensively to visualize and quantify protein interactions in live cells7, but did not provide an effective means of isolating and charact...

Protein complex assembly is a key process in maintaining the spatiotemporal specificity of many signalling pathways1,2. While the critical nature of this regulatory role is widely recognized, there is a lack of experimental techniques available to scrutinize these complexes. Most interactomics studies focus upon interactions with individual proteins, or the sequential enrichment of individual complex components. Here we present a technique for the isolation of a specific protein dimer while excluding the individual moieties of the component proteins as well as complexes formed with competing binding partners3. We have called this technique Bimolecular Complementation Affinity Purification (BiCAP), as it is a combination of a previously existing protein fragment complementation assay, Bimolecular Fluorescence Complementation (BiFC), with the novel use of a conformation-specific recombinant nanobody towards GFP and its derivatives (see Table of Materials).
A typical protein-fragment complementation assay relies on the expression of &#34;bait&#34; and &#34;prey&#34; proteins fused to split fragments of reporters such as luciferase4, &#946;-galactosidase5, or green fluorescent protein (GFP)6 (Figure 1A). Through the interaction of the bait and prey proteins, the split reporter domains are encouraged to refold into a functional structure, allowing the interaction of the bait and prey proteins to be visualized or quantified. BiCAP was adapted from a version of this technique that made use of fragments of the GFP variant Venus. Fluorescent protein complementation assays are a popular method for visualizing protein-protein interactions in a live cell, but until now have been limited to this one function7. BiCAP represents a significant advance in this regard, as this technique not only allows for visualization, but also the isolation and interrogation of the resulting protein-protein interaction.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57109/57109fig1.jpg" /> 
Figure 1: The structural principal behind the BiCAP technique. (A) A schematic outlining the principal behind bimolecular fluorescence complementation showing the &#39;bait&#39; and &#39;prey&#39; proteins tagged with the N-terminal V1 or C-terminal V2 fragments of the full-length Venus protein. (B) Structural analysis of the interaction interface (cyan) between the GFP nanobody (green) and recombined Venus, showing the position of the V1 (grey) and V2 (orange) fragments (PDB accession 3OGO). This figure is republished fromCroucher et al.3 Reprinted with permission from AAAS. <a href="//cloudflare.jove.com/files/ftp_upload/57109/57109fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The BiCAP technique makes use of two non-fluorescent fragments of Venus (named V1 and V2) which associate with a low degree of affinity unless an interaction occurs between their fusion partners. In this instance, the two split domains refold into the functional &#946;-barrel structure of the fluorophore (Figure 1B)6. The key innovation of BiCAP comes from the introduction of the recombinant GFP nanobody, which recognizes a three-dimensional epitope on the &#946;-barrel of GFP (and variants such as Venus) which is only present on the correctly recombined and folded fluorophore (Figure 1B)8. Crucially, the GFP nanobody does not bind to either of the individual Venus fragments. This facilitates the isolation of protein dimers only after the two proteins have formed a complex of their own volition, leading to more representative results than those acquired from methods that make use of chemically-induced, forced interactions9.
BiCAP is a powerful technique that specifically focuses on multi-protein complexes, which can potentially be combined with a number of downstream applications to improve the granularity of our understanding of the role these complexes play in signal transduction. It also encompasses the important feature of allowing visualization of protein interactions in situ. To date, BiCAP has been demonstrated as an effective method of analyzing the interactome of receptor tyrosine kinase (RTK) dimers3, but the adaptability of this method means that it can be adopted in almost any protein interaction context.

<table><tbody><tr><td>LR Clonase II Plus enzyme</td><td>Thermo Fisher Scientific</td><td>12538120</td><td>Recombinase enzyme required for Gateway cloning (Step 1) into pDEST BiFC destination vectors</td></tr><tr><td>Proteinase K, recombinant, PCR grade</td><td>Thermo Fisher Scientific</td><td>EO0491</td><td></td></tr><tr><td>14 mL round-bottomed polypropylene tube</td><td>Corning</td><td>352059</td><td></td></tr><tr><td>Ampicillin</td><td>Roche Diagnostics Australia</td><td>10835242001</td><td>Stock solution prepared at 100 &amp;mu;g/mL in distilled water.</td></tr><tr><td>Miniprep kit</td><td>Promega Corporation</td><td>A1330</td><td></td></tr><tr><td>Maxiprep kit</td><td>Life Technologies Australia</td><td>K2100-07</td><td></td></tr><tr><td>DMEM</td><td>Gibco</td><td>11995-073</td><td></td></tr><tr><td>FBS</td><td>Life Technologies Australia</td><td>10099-141</td><td></td></tr><tr><td>Penicillin/Streptomycin</td><td>Life Technologies Australia</td><td>15070-063</td><td></td></tr><tr><td>jetPRIME transfection buffer</td><td>Polyplus</td><td>114-15</td><td></td></tr><tr><td>jetPRIME transfection reagent</td><td>Polyplus</td><td>114-15</td><td></td></tr><tr><td>PhosSTOP (Phosphatase inhibitor)</td><td>Sigma-Aldrich</td><td>4906837001</td><td></td></tr><tr><td>cOmplete, Mini, EDTA-free Protease inhibitor cocktail</td><td>Roche</td><td>11873580001</td><td></td></tr><tr><td>Cell Scraper</td><td>Sarstedt</td><td>83.1832</td><td></td></tr><tr><td>GFP-Trap_A</td><td>Chromotek Gmbh</td><td>gta-100</td><td>GFP nanobody coupled to agarose beads</td></tr><tr><td>N-terminal GFP monoclonal antibody&amp;nbsp;</td><td>Covance</td><td>MMS-118P</td><td>Will detect the V1 tag within the BiFC vectors</td></tr><tr><td>C-terminal GFP monoclonal antibody</td><td>Roche</td><td>11814460001</td><td>Will detect the V2 tag within the BiFC vectors</td></tr><tr><td>Sample buffer</td><td>Invitrogen</td><td>NP0008</td><td>Supplemented with 1 mL &amp;beta;-mercaptoethanol.</td></tr><tr><td>Sequencing grade modified trypsin</td><td>Promega Corporation</td><td>V5117h</td><td></td></tr><tr><td>LoBind microcentrifuge tubes</td><td>Point of Care Diagnostics</td><td>0030 108 116</td><td></td></tr><tr><td>Iodoacetamide</td><td>Sigma-Aldrich</td><td>I1149-5G</td><td>Prepared at 5 mg/mL in ultrapure water</td></tr><tr><td>Trifluoroacetic Acid - Sequanal Grade</td><td>Thermo Fisher</td><td>10628494</td><td></td></tr><tr><td>3M Empore solid phase extraction C18 disks (octadecyl) - 4.7 cm</td><td>Thermo Fisher</td><td>14-386-2</td><td>To prepare stage tips, cut 1 mm disks using an appropriately sized hole punch. Alternatively, pre-prepared stage tips can also be purchased, see below.</td></tr><tr><td>C18 Stage Tips, 10 &amp;micro;L bed</td><td>Thermo Fisher</td><td>87782</td><td></td></tr><tr><td>Formic acid OPTIMA for LC/MS grade 50mL</td><td>Thermo Fisher</td><td>FSBA117-50</td><td></td></tr><tr><td>1.9 &amp;mu;m C18 ReproSil particles</td><td>Dr. Maisch GmbH</td><td>r119.aq.</td><td>Stationary phase particles</td></tr><tr><td>Acetonitrile OPTIMA LC/MS grade&amp;nbsp;</td><td>Thermo Fisher</td><td>FSBA955-4</td><td></td></tr><tr><td>Easy-nLC HPLC&amp;nbsp;</td><td>Thermo Fisher</td><td></td><td></td></tr><tr><td>LTQ Orbitrap Velos Pro</td><td>Thermo Fisher</td><td></td><td></td></tr><tr><td>Triton X-100</td><td>Sigma-Aldrich</td><td>T8787</td><td>Non-ionic detergent (100%)</td></tr><tr><td>DH5&amp;alpha; cells</td><td>Thermo Fisher</td><td></td><td>Heat-shock-competent cells</td></tr></tbody></table>

dissecting multi-protein signaling complexes by bimolecular complementation affinity purification (bicap)

The assembly of protein complexes is a central mechanism underlying the regulation of many cell signaling pathways. A major focus of biomedical research is deciphering how these dynamic protein complexes act to integrate signals from multiple sources in order to direct a specific biological response, and how this becomes deregulated in many disease settings. Despite the importance of this key biochemical mechanism, there is a lack of experimental techniques that can facilitate the specific and sensitive deconvolution of these multi-molecular signaling complexes.
Here this shortcoming is addressed through the combination of a protein complementation assay with a conformation-specific nanobody, which we have termed Bimolecular Complementation Affinity Purification (BiCAP). This novel technique facilitates the specific isolation and downstream proteomic characterization of any pair of interacting proteins, to the exclusion of un-complexed individual proteins and complexes formed with competing binding partners. 
The BiCAP technique is adaptable to a wide array of downstream experimental assays, and the high degree of specificity afforded by this technique allows more nuanced investigations into the mechanics of protein complex assembly than is currently possible using standard affinity purification techniques.

Following the use of recombination cloning to generate of V1 and V2 tagged genes of interest with the BiFC pDEST plasmids, co-transfection of two plasmids containing an interacting pair of proteins will result in the generation of a Venus fluorescent signal after approximately 8-24 hours. In the absence of a positive signal it is possible that the protein interaction may not be occurring due to the choice of cell line, a low transfection efficiency or that the orientation of the BiFC tags...

Watch this Scientific Journal Video about Dissecting Multi-protein Signaling Complexes by Bimolecular Complementation Affinity Purification BiCAP at JoVE.com

Dissecting Multi-protein Signaling Complexes by Bimolecular Complementation Affinity Purification BiCAP

This manuscript describes the protocol for Bimolecular Complementation Affinity Purification (BiCAP). This novel method facilitates the specific isolation and downstream proteomic characterization of any two interacting proteins, while excluding un-complexed individual proteins as well as complexes formed with competing binding partners.

Dissecting Multi-protein Signaling Complexes by Bimolecular Complementation Affinity Purification (BiCAP)

The assembly of protein complexes is a central mechanism underlying the regulation of many cell signaling pathways. A major focus of biomedical research ...

dissecting-multi-protein-signaling-complexes-bimolecular

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JoVE Journal

Immunology and Infection

7.4K Views.  Garvan Institute of Medical Research. Dit manuscript beschrijft het protocol voor Bimolecular complementatie affiniteit zuivering (BiCAP). Deze nieuwe methode vergemakkelijkt de specifieke isolatie en downstream proteomic karakterisering van elk twee interacterende eiwitten, terwijl met uitzondering van VN-complexvorm individuele eiwitten evenals complexen gevormd met concurrerende bindende partners.

Video: Dissecting Multi-protein Signaling Complexes by Bimolecular Complementation Affinity Purification BiCAP

Tandem Affinity Purification of Protein Complexes from Eukaryotic Cells

The purification of active protein-protein and protein-nucleic acid complexes is crucial for the characterization of enzymatic activities and de novo identification of novel subunits and post-translational modifications. Bacterial systems allow for the expression and purification of a wide variety of single polypeptides and protein complexes. However, this system does not enable the purification of protein subunits that contain post-translational modifications (e.g., phosphorylation and acetylation), and the identification of novel regulatory subunits that are only present/expressed in the eukaryotic system. Here, we provide a detailed description of a novel, robust, and efficient tandem affinity purification (TAP) method using STREP- and FLAG-tagged proteins that facilitates the purification of protein complexes with transiently or stably expressed epitope-tagged proteins from eukaryotic cells. This protocol can be applied to characterize protein complex functionality, to discover post-translational modifications on complex subunits, and to identify novel regulatory complex components by mass spectrometry. Notably, this TAP method can be applied to study protein complexes formed by eukaryotic or pathogenic (viral and bacterial) components, thus yielding a wide array of downstream experimental opportunities. We propose that researchers working with protein complexes could utilize this approach in many different ways.

We describe here a novel, robust, and efficient tandem affinity purification (TAP) method for the expression, isolation, and characterization of protein complexes from eukaryotic cells. This protocol could be utilized for the biochemical characterization of discrete complexes as well as the identification of novel interactors and post-translational modifications that regulate their function.

The purification of active protein-protein and protein-nucleic acid complexes is crucial for the characterization of enzymatic activities and de novo ...

Biochemistry

Resolving Affinity Purified Protein Complexes by Blue Native PAGE and Protein Correlation Profiling

Most proteins act in association with others; hence, it is crucial to characterize these functional units in order to fully understand biological processes. Affinity purification coupled to mass spectrometry (AP-MS) has become the method of choice for identifying protein-protein interactions. However, conventional AP-MS studies provide information on protein interactions, but the organizational information is lost. To address this issue, we developed a strategy to unravel the distinct functional assemblies a protein might be involved in, by resolving affinity-purified protein complexes prior to their characterization by mass spectrometry. Protein complexes isolated through affinity purification of a bait protein using an epitope tag and competitive elution are separated through blue native electrophoresis. Comparison of protein migration profiles through correlation profiling using quantitative mass spectrometry allows assignment of interacting proteins to distinct molecular entities. This method is able to resolve protein complexes of close molecular weights that might not be resolved by traditional chromatographic techniques such as gel filtration. With little more work than conventional AP-geLC-MS/MS, we demonstrate this strategy may in many cases be adequate for obtaining protein complex topological information concomitantly to identifying protein interactions.

Here we present protocols for affinity purification of protein complexes and their separation by blue native PAGE, followed by protein correlation profiling using label free quantitative mass spectrometry. This method is useful to resolve interactomes into distinct protein complexes.

Most proteins act in association with others; hence, it is crucial to characterize these functional units in order to fully understand biological ...

Split-BioID &#8212; Proteomic Analysis of Context-specific Protein Complexes in Their Native Cellular Environment

To complement existing affinity purification (AP) approaches for the identification of protein-protein interactions (PPI), enzymes have been introduced that allow the proximity-dependent labeling of proteins in living cells. One such enzyme, BirA* (used in the BioID approach), mediates the biotinylation of proteins within a range of approximately 10 nm. Hence, when fused to a protein of interest and expressed in cells, it allows the labeling of proximal proteins in their native environment. As opposed to AP that relies on the purification of assembled protein complexes, BioID detects proteins that have been marked within cells no matter whether they are still interacting with the protein of interest when they are isolated. Since it biotinylates proximal proteins, one can moreover capitalize on the exceptional affinity of streptavidin for biotin to very efficiently isolate them. While BioID performs better than AP for identifying transient or weak interactions, both AP- and BioID-mass spectrometry approaches provide an overview of all possible interactions a given protein may have. However, they do not provide information on the context of each identified PPI. Indeed, most proteins are typically part of several complexes, corresponding to distinct maturation steps or different functional units. To address this common limitation of both methods, we have engineered a protein-fragments complementation assay based on the BirA* enzyme. In this assay, two inactive fragments of BirA* can reassemble into an active enzyme when brought in close proximity by two interacting proteins to which they are fused. The resulting split-BioID assay thus allows the labeling of proteins that assemble around a pair of interacting proteins. Provided these two only interact in a given context, split-BioID then allows the analysis of specific context-dependent functional units in their native cellular environment. Here, we provide a step-by-step protocol to test and apply split-BioID to a pair of interacting proteins.

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.

To complement existing affinity purification (AP) approaches for the identification of protein-protein interactions (PPI), enzymes have been introduced ...

Analyzing Dynamic Protein Complexes Assembled On and Released From Biolayer Interferometry Biosensor Using Mass Spectrometry and Electron Microscopy

In vivo, proteins are often part of large macromolecular complexes where binding specificity and dynamics ultimately dictate functional outputs. In this work, the pre-endosomal anthrax toxin is assembled and transitioned into the endosomal complex. First, the N-terminal domain of a cysteine mutant lethal factor (LFN) is attached to a biolayer interferometry (BLI) biosensor through disulfide coupling in an optimal orientation, allowing protective antigen (PA) prepore to bind (Kd 1 nM). The optimally oriented LFN-PAprepore complex then binds to soluble capillary morphogenic gene-2 (CMG2) cell surface receptor (Kd 170 pM), resulting in a representative anthrax pre-endosomal complex, stable at pH 7.5. This assembled complex is then subjected to acidification (pH 5.0) representative of the late endosome environment to transition the PAprepore into the membrane inserted pore state. This PApore state results in a weakened binding between the CMG2 receptor and the LFN-PApore and a substantial dissociation of CMG2 from the transition pore. The thio-attachment of LFN to the biosensor surface is easily reversed by dithiothreitol. Reduction on the BLI biosensor surface releases the LFN-PAprepore-CMG2 ternary complex or the acid transitioned LFN-PApore complexes into microliter volumes. Released complexes are then visualized and identified using electron microscopy and mass spectrometry. These experiments demonstrate how to monitor the kinetic assembly/disassembly of specific protein complexes using label-free BLI methodologies and evaluate the structure and identity of these BLI assembled complexes by electron microscopy and mass spectrometry, respectively, using easy-to-replicate sequential procedures.

Here we present a protocol to monitor the assembly and disassembly of the anthrax toxin using biolayer interferometry (BLI). Following assembly/disassembly on the biosensor surface, the large protein complexes are released from the surface for visualization and identification of components of the complexes using electron microscopy and mass spectrometry, respectively.

In vivo, proteins are often part of large macromolecular complexes where binding specificity and dynamics ultimately dictate functional outputs. In this ...

Quantification of Protein Interaction Network Dynamics using Multiplexed Co-Immunoprecipitation

Dynamic protein-protein interactions control cellular behavior, from motility to DNA replication to signal transduction. However, monitoring dynamic interactions among multiple proteins in a protein interaction network is technically difficult. Here, we present a protocol for Quantitative Multiplex Immunoprecipitation (QMI), which allows quantitative assessment of fold changes in protein interactions based on relative fluorescence measurements of Proteins in Shared Complexes detected by Exposed Surface epitopes (PiSCES). In QMI, protein complexes from cell lysates are immunoprecipitated onto microspheres, and then probed with a labeled antibody for a different protein in order to quantify the abundance of PiSCES. Immunoprecipitation antibodies are conjugated to different MagBead spectral regions, which allows a flow cytometer to differentiate multiple parallel immunoprecipitations and simultaneously quantify the amount of probe antibody associated with each. QMI does not require genetic tagging and can be performed using minimal biomaterial compared to other immunoprecipitation methods. QMI can be adapted for any defined group of interacting proteins, and has thus far been used to characterize signaling networks in T cells and neuronal glutamate synapses. Results have led to new hypothesis generation with potential diagnostic and therapeutic applications. This protocol includes instructions to perform QMI, from the initial antibody panel selection through to running assays and analyzing data. The initial assembly of a QMI assay involves screening antibodies to generate a panel, and empirically determining an appropriate lysis buffer. The subsequent reagent preparation includes covalently coupling immunoprecipitation antibodies to MagBeads, and biotinylating probe antibodies so they can be labeled by a streptavidin-conjugated fluorophore. To run the assay, lysate is mixed with MagBeads overnight, and then beads are divided and incubated with different probe antibodies, and then a fluorophore label, and read by flow cytometry. Two statistical tests are performed to identify PiSCES that differ significantly between experimental conditions, and results are visualized using heatmaps or node-edge diagrams.

Quantitative Multiplex Immunoprecipitation (QMI) uses flow cytometry for sensitive detection of differences in the abundance of targeted protein-protein interactions between two samples. QMI can be performed using a small amount of biomaterial, does not require genetically engineered tags, and can be adapted for any previously defined protein interaction network.

Dynamic protein-protein interactions control cellular behavior, from motility to DNA replication to signal transduction. However, monitoring dynamic ...

Isolation of Labile Multi-protein Complexes by in vivo Controlled Cellular Cross-Linking and Immuno-magnetic Affinity Chromatography

The dynamic nature of cellular machineries is frequently built on transient and/or weak protein associations. These low affinity interactions preclude stringent methods for the isolation and identification of protein networks around a protein of interest. The use of chemical crosslinkers allows the selective stabilization of labile interactions, thus bypassing biochemical limitations for purification. Here we present a protocol amenable for cells in culture that uses a homobifunctional crosslinker with a spacer arm of 12 &Aring;, dithiobis-(succinimidyl proprionate) (DSP). DSP is cleaved by reduction of a disulphide bond present in the molecule. Cross-linking combined with immunoaffinity chromatography of proteins of interest with magnetic beads allows the isolation of protein complexes that otherwise would not withstand purification. This protocol is compatible with regular western blot techniques and it can be scaled up for protein identification by mass spectrometry1.
Stephanie A. Zlatic and Pearl V. Ryder contributed equally to this work.

Isolation of Labile Multi-protein Complexes by in vivo Controlled Cellular Cross-Linking and Immuno-magnetic Affinity Chromatography

The cell permeable crosslinker DSP [dithiobis-(succinimidyl propionate)] stabilizes transient and labile interactions in vivo, which allows their isolation using stringent protein complex purification techniques. Here we present a technique for crosslinking cells grown in culture followed by isolation of protein complexes by immunoprecipitation.

The dynamic nature of cellular machineries is frequently built on transient and/or weak protein associations. These low affinity interactions preclude ...

Biology

Bimolecular Fluorescence Complementation

Defining the subcellular distribution of signaling complexes is imperative to understanding the output from that complex. Conventional methods such as immunoprecipitation do not provide information on the spatial localization of complexes. In contrast, BiFC monitors the interaction and subcellular compartmentalization of protein complexes. In this method, a fluororescent protein is split into amino- and carboxy-terminal non-fluorescent fragments which are then fused to two proteins of interest. Interaction of the proteins results in reconstitution of the fluorophore (Figure 1)1,2. A limitation of BiFC is that once the fragmented fluorophore is reconstituted the complex is irreversible3. This limitation is advantageous in detecting transient or weak interactions, but precludes a kinetic analysis of complex dynamics. An additional caveat is that the reconstituted flourophore requires 30min to mature and fluoresce, again precluding the observation of real time interactions4. BiFC is a specific example of the protein fragment complementation assay (PCA) which employs reporter proteins such as green fluorescent protein variants (BiFC), dihydrofolate reductase, b-lactamase, and luciferase to measure protein:protein interactions5,6. Alternative methods to study protein:protein interactions in cells include fluorescence co-localization and F&ouml;rster resonance energy transfer (FRET)7. For co-localization, two proteins are individually tagged either directly with a fluorophore or by indirect immunofluorescence. However, this approach leads to high background of non-interacting proteins making it difficult to interpret co-localization data. In addition, due to the limits of resolution of confocal microscopy, two proteins may appear co-localized without necessarily interacting. With BiFC, fluorescence is only observed when the two proteins of interest interact. FRET is another excellent method for studying protein:protein interactions, but can be technically challenging. FRET experiments require the donor and acceptor to be of similar brightness and stoichiometry in the cell. In addition, one must account for bleed through of the donor into the acceptor channel and vice versa. Unlike FRET, BiFC has little background fluorescence, little post processing of image data, does not require high overexpression, and can detect weak or transient interactions. Bioluminescence resonance energy transfer (BRET) is a method similar to FRET except the donor is an enzyme (e.g. luciferase) that catalyzes a substrate to become bioluminescent thereby exciting an acceptor. BRET lacks the technical problems of bleed through and high background fluorescence but lacks the ability to provide spatial information due to the lack of substrate localization to specific compartments8. Overall, BiFC is an excellent method for visualizing subcellular localization of protein complexes to gain insight into compartmentalized signaling.

The subcellular localization of proteins is important in determining the spatio-temporal regulation of cell signaling. Here, we describe bimolecular fluorescence complementation (BiFC) as a straightforward method for monitoring the spatial interactions of proteins in the cell.

Defining the subcellular distribution of signaling complexes is imperative to understanding the output from that complex. Conventional methods such as ...

Identification of Protein Complexes in Escherichia coli using Sequential Peptide Affinity Purification in Combination with Tandem Mass Spectrometry

Since most cellular processes are mediated by macromolecular assemblies, the systematic identification of protein-protein interactions (PPI) and the identification of the subunit composition of multi-protein complexes can provide insight into gene function and enhance understanding of biological systems1, 2. Physical interactions can be mapped with high confidence vialarge-scale isolation and characterization of endogenous protein complexes under near-physiological conditions based on affinity purification of chromosomally-tagged proteins in combination with mass spectrometry (APMS). This approach has been successfully applied in evolutionarily diverse organisms, including yeast, flies, worms, mammalian cells, and bacteria1-6. In particular, we have generated a carboxy-terminal Sequential Peptide Affinity (SPA) dual tagging system for affinity-purifying native protein complexes from cultured gram-negative Escherichia coli, using genetically-tractable host laboratory strains that are well-suited for genome-wide investigations of the fundamental biology and conserved processes of prokaryotes1, 2, 7. Our SPA-tagging system is analogous to the tandem affinity purification method developed originally for yeast8, 9, and consists of a calmodulin binding peptide (CBP) followed by the cleavage site for the highly specific tobacco etch virus (TEV) protease and three copies of the FLAG epitope (3X FLAG), allowing for two consecutive rounds of affinity enrichment. After cassette amplification, sequence-specific linear PCR products encoding the SPA-tag and a selectable marker are integrated and expressed in frame as carboxy-terminal fusions in a DY330 background that is induced to transiently express a highly efficient heterologous bacteriophage lambda recombination system10. Subsequent dual-step purification using calmodulin and anti-FLAG affinity beads enables the highly selective and efficient recovery of even low abundance protein complexes from large-scale cultures. Tandem mass spectrometry is then used to identify the stably co-purifying proteins with high sensitivity (low nanogram detection limits).
Here, we describe detailed step-by-step procedures we commonly use for systematic protein tagging, purification and mass spectrometry-based analysis of soluble protein complexes from E. coli, which can be scaled up and potentially tailored to other bacterial species, including certain opportunistic pathogens that are amenable to recombineering. The resulting physical interactions can often reveal interesting unexpected components and connections suggesting novel mechanistic links. Integration of the PPI data with alternate molecular association data such as genetic (gene-gene) interactions and genomic-context (GC) predictions can facilitate elucidation of the global molecular organization of multi-protein complexes within biological pathways. The networks generated for E. coli can be used to gain insight into the functional architecture of orthologous gene products in other microbes for which functional annotations are currently lacking.

Identification of Protein Complexes in Escherichia coli using Sequential Peptide Affinity Purification in Combination with Tandem Mass Spectrometry

Affinity purification of tagged proteins in combination with mass spectrometry (APMS) is a powerful method for the systematic mapping of protein interaction networks and for investigating the mechanistic basis of biological processes. Here, we describe an optimized sequential peptide affinity (SPA) APMS procedure developed for the bacterium Escherichia coli that can be used to isolate and characterize stable multi-protein complexes to near homogeneity even starting from low copy numbers per cell.

Since most cellular processes are mediated by macromolecular assemblies, the systematic identification of protein-protein interactions (PPI) and the ...

The MultiBac Protein Complex Production Platform at the EMBL

Proteomics research revealed the impressive complexity of eukaryotic proteomes in unprecedented detail. It is now a commonly accepted notion that proteins in cells mostly exist not as isolated entities but exert their biological activity in association with many other proteins, in humans ten or more, forming assembly lines in the cell for most if not all vital functions.1,2 Knowledge of the function and architecture of these multiprotein assemblies requires their provision in superior quality and sufficient quantity for detailed analysis. The paucity of many protein complexes in cells, in particular in eukaryotes, prohibits their extraction from native sources, and necessitates recombinant production. The baculovirus expression vector system (BEVS) has proven to be particularly useful for producing eukaryotic proteins, the activity of which often relies on post-translational processing that other commonly used expression systems often cannot support.3 BEVS use a recombinant baculovirus into which the gene of interest was inserted to infect insect cell cultures which in turn produce the protein of choice. MultiBac is a BEVS that has been particularly tailored for the production of eukaryotic protein complexes that contain many subunits.4 A vital prerequisite for efficient production of proteins and their complexes are robust protocols for all steps involved in an expression experiment that ideally can be implemented as standard operating procedures (SOPs) and followed also by non-specialist users with comparative ease. The MultiBac platform at the European Molecular Biology Laboratory (EMBL) uses SOPs for all steps involved in a multiprotein complex expression experiment, starting from insertion of the genes into an engineered baculoviral genome optimized for heterologous protein production properties to small-scale analysis of the protein specimens produced.5-8 The platform is installed in an open-access mode at EMBL Grenoble and has supported many scientists from academia and industry to accelerate protein complex research projects.

Protein complexes catalyze key cellular functions. Detailed functional and structural characterization of many essential complexes requires recombinant production. MultiBac is a baculovirus/insect cell system particularly tailored for expressing eukaryotic proteins and their complexes. MultiBac was implemented as an open-access platform, and standard operating procedures developed to maximize its utility.

Proteomics  research revealed the impressive complexity of eukaryotic proteomes in  unprecedented detail. It is now a commonly accepted notion that ...

Flow Cytometric Analysis of Bimolecular Fluorescence Complementation: A High Throughput Quantitative Method to Study Protein-protein Interaction

Among methods to study protein-protein interaction inside cells, Bimolecular Fluorescence Complementation (BiFC) is relatively simple and sensitive. BiFC is based on the production of fluorescence using two non-fluorescent fragments of a fluorescent protein (Venus, a Yellow Fluorescent Protein variant, is used here). Non-fluorescent Venus fragments (VN and VC) are fused to two interacting proteins (in this case, AKAP-Lbc and PDE4D3), yielding fluorescence due to VN-AKAP-Lbc-VC-PDE4D3 interaction and the formation of a functional fluorescent protein inside cells.
BiFC provides information on the subcellular localization of protein complexes and the strength of protein interactions based on fluorescence intensity. However, BiFC analysis using microscopy to quantify the strength of protein-protein interaction is time-consuming and somewhat subjective due to heterogeneity in protein expression and interaction. By coupling flow cytometric analysis with BiFC methodology, the fluorescent BiFC protein-protein interaction signal can be accurately measured for a large quantity of cells in a short time. Here, we demonstrate an application of this methodology to map regions in PDE4D3 that are required for the interaction with AKAP-Lbc. This high throughput methodology can be applied to screening factors that regulate protein-protein interaction.

Flow cytometric analysis of Bimolecular Fluorescence Complementation provides a high throughput quantitative method to study protein-protein interaction. This methodology can be applied to mapping protein binding sites and for screening factors that regulate protein-protein interaction.

Among methods to study protein-protein interaction inside cells, Bimolecular Fluorescence Complementation (BiFC) is relatively simple and sensitive. BiFC ...

Dissecting Multi-protein Signaling Complexes by Bimolecular Complementation Affinity Purification (BiCAP)

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