We thank Dr. Ben Nacev for advice on design of the photoaffinity labeling protocol, Dr. Wei Shi for synthesizing the itraconazole photoaffinity probe, Dr. Yongjun Dang for advice on affinity pull-down experiments, and other members of the J.O.L. laboratory for helpful comments and support. This work was supported in part by a PhRMA Foundation Fellowship in Pharmacology/Toxicology (to S.A.H.); National Cancer Institute Grant R01CA184103; the Flight Attendant Medical Research Institute; Prostate Cancer Foundation (J.O.L.); and the Johns Hopkins Institute for Clinical and Translational Research, which is funded in part by Grant UL1 TR 001079 from the National Center for Advancing Translational Sciences (NCATS).

NOTE: This protocol was adapted from MacKinnon and Taunton10 for use in live cells.
1. Preparation of Cultured Cells
<ol>
	<li>Prepare sterile 6-cm cell culture dishes for the number of samples desired (see below). 
	NOTE: One dish of cells is used per treatment condition, but if more protein is required 2 or 3 dishes can be prepared per condition and combined after UV irradiation.
	<ol>
		<li>Prepare at least 3 dishes of cells; Negative control with DMSO only (D), Probe treatment only (P), and probe + competitor (C).</li>
		<li>Optionally, prepare a 4th dish as a no UV control, to be treated with probe but not subjected to UV light. 
		NOTE: If multiple competitor molecules are to be tested, such as different analogs of the small molecule of interest, prepare additional competitor plates as necessary.</li>
	</ol>
	</li>
	<li>Add HEK293T cells to culture dishes in 4 ml culture media. Use 3.5 million cells per plate. 
	NOTE: The number of cells should be determined for each cell type so that cells are almost 100% confluent at the beginning of the experiment, in order to achieve the maximum protein yield. For HUVEC, use 0.3 million cells per plate. A good approximation is 1/10 of the cells in a confluent 15 cm dish. HEK293T cells should be cultured in Dulbecco&#39;s Modified Eagles Medium supplemented with 10% Fetal Bovine Serum (FBS) and 1% Penicillin/Streptomycin.</li>
	<li>Return the cells to the culture incubator (37 &#176;C, 5% CO2) overnight.</li>
</ol>
2. Treatment of Cells and Photocrosslinking
<ol>
	<li>The next day, pre-aliquot the following drugs into 1.5 ml microcentrifuge tubes:
	<ol>
		<li>For the first treatment, add 4 &#181;l of DMSO to 2 tubes and 4 &#181;l of 10 mM competitor (i.e., the unmodified parent compound) to 1 tube.</li>
		<li>For the second treatment, add 16 &#181;l of DMSO to 1 tube and 16 &#181;l of 50 &#181;M photoaffinity probe to 2 tubes. 
		NOTE: The optimal concentration may be different for different probes (see discussion), but between 200-500 nM is a good starting point (here we use 200 nM). The concentration of competitor should exceed that of the probe at least 20 fold (here we use 10 &#181;M). The final concentration of DMSO should be equal in all plates and should not exceed 0.5% (20 &#181;l volume).</li>
	</ol>
	</li>
	<li>Bring the cell plates into the cell culture hood and add the pre-aliquoted drugs from step 2.1.1 as follows: Aspirate 1 ml of culture media, resuspending the pre-aliquoted drug in the media, and gently adding it back to the plate drop wise. Add the DMSO to the DMSO (D) plate and the probe (P) plate, and add the competitor to the competition (C) plate. Return the plates to the culture incubator for 30 min.</li>
	<li>With lights in the culture hood dimmed, add the pre-aliquoted probe from step 2.1.2 to probe (P) and competition (C) plates and add the DMSO to the DMSO (D) plate, as above. Return the plates to the culture incubator for 1 hr.</li>
	<li>During the incubation, prepare the following items; A tray of ice that can fit all of the plates; ice-cold lysis buffer (PBS [pH 8.5] + 1x protease inhibitor cocktail [complete EDTA-free, diluted from 50x stock solution made in water]; 200 &#181;l/plate); microcentrifuge tubes labeled D, P, or C for each of the different treatment conditions, on ice.</li>
	<li>15 min before the end of the 1 hr incubation, set up the UV lamp (365 nm) in the cold room and turn it on to warm up the bulb.</li>
	<li>After 1 hr incubation with the probe, place dishes on ice. Wash the cells gently with 5 ml ice-cold PBS (pH 7.4) to remove excess probe. Re-cover the cells with 4 ml ice-cold PBS.</li>
	<li>Place the dish of cells centered 3 cm under the UV lamp on top of an ice pack to minimize heating from the lamp. Irradiate for 3 min. Remove the dish to the tray of ice and repeat for all samples.</li>
	<li>After irradiation, aspirate the PBS from the cells and add 200 &#181;l of the ice-cold PBS (pH 8.5) with protease inhibitors to each plate. Detach the cells from the plate using a rubber scraper and transfer to the pre-labeled microcentrifuge tubes on ice.</li>
	<li>Add SDS to a final concentration of 0.4% (10 &#181;l of 10% SDS into 250 &#181;l of sample). 
	CAUTION: SDS powder is hazardous. Avoid inhaling SDS powder by wearing a mask over nose and mouth.</li>
	<li>Lyse the cells by sonicating the suspension for 10 pulses (output 1, duty cycle 30%) and incubate on ice for 1 min before a second round of 10 pulses.</li>
	<li>If needed, remove 2 &#181;l of suspension and check under a light microscope to ensure complete cell lysis.</li>
	<li>Boil the samples on a hot plate set to 95 &#176;C for 5 min to complete cell lysis and denature all the proteins.</li>
	<li>Measure the protein concentration in each sample using a spectrophotometric detergent-compatible protein assay, such as the Dc protein assay19, according to the manufacturer&#39;s instructions, with a spectrophotometer set to measure absorbance at 750 nm.</li>
	<li>Normalize the protein concentration to 2.5 mg/ml (or if concentrations are lower than 2.5 mg/ml, normalize to the sample with the lowest protein concentration) by adding PBS pH 8.5 + 0.4% SDS as needed (for example, if samples are 5 mg/ml in 250 &#181;l, add 250 &#181;l of PBS pH 8.5 + 0.4% SDS to obtain a final concentration of 2.5 mg/ml).</li>
</ol>
3. Attachment of Fluorescent Tag by Click Chemistry for Visualization of Labeled Proteins
<ol>
	<li>Remove 40 &#181;l of the cell lysate and transfer to a new microcentrifuge tube. Keep the remaining lysates for reaction with biotin-azide (step 4).</li>
	<li>Add the following reagents in order: 0.2 &#181;l fluor-azide (1 mM stock solution in DMSO), 0.58 &#181;l TCEP (100 mM stock in water with 4 equivalents NaOH added, prepared fresh), and 3.38 &#181;l TBTA (1.7 mM stock in a 4:1 ratio of t-butanol to DMSO). Vortex to mix.</li>
	<li>Add 1.14 &#181;l CuSO4-5H2O (50 mM in water) to start the reaction. Vortex briefly and incubate at room temperature for 30 min in the dark.</li>
	<li>Add 50 &#181;l of 2x SDS sample buffer to quench the reaction. At this point, run the samples directly on an SDS-PAGE gel20 for best results, or store at -20 &#176;C overnight if necessary.</li>
	<li>If freezing samples, reheat for 5 min at 95 &#176;C before loading onto the gel. 
	NOTE: Since the molecular weight of the target protein is typically not known at this stage, it is advisable to run 2 gels with different acrylamide concentrations (i.e., 8% and 15%), or a wide molecular weight range gradient gel (i.e., 4-15%), to ensure complete molecular weight coverage.</li>
	<li>When the dye front has reached the end of the gel, continue running the gel for an additional 5 min to ensure all of the excess unreacted fluor-azide has completely exited the gel.</li>
	<li>Remove the gel to a container with ddH2O and incubate for 10 min with gentle agitation, to wash away all excess fluor-azide.</li>
	<li>Place the gel onto a glass plate and scan the gel using a Typhoon fluorescent gel scanner according to the manufacturer&#39;s instructions.</li>
</ol>
4. Attachment of Biotin Tag by Click Chemistry for Affinity Purification of Labeled Proteins
<ol>
	<li>Of the remaining lysates from step 3.1, use the maximum amount after protein normalization such that all samples are the same volume (for example, if after normalization to 2.5 mg/ml the resulting sample volumes are 500, 550, and 600 &#181;l, use 500 &#181;l of each).</li>
	<li>Pre-clear the lysates by adding to 50 &#181;l high-capacity streptavidin agarose beads, pre-washed 2x with PBS (pH 7.4). Incubate 1 hr at 4 &#176;C with rotation.</li>
	<li>Pellet the beads by centrifugation at 1,000 x g for 3 min. Remove the supernatant to a new microcentrifuge tube on ice and discard the beads.</li>
	<li>Remove 1-2% of the DMSO sample to a new tube labeled &#34;input&#34;. Add an equivalent volume of 2x SDS sample buffer and store at -20 &#176;C.</li>
	<li>Per 500 &#181;l of lysate, add the following reagents: 1.38 &#181;l biotin-azide (10 mM stock in DMSO), 5.5 &#181;l TCEP, and 32.5 &#181;l TBTA. Vortex to mix.</li>
	<li>Add 11 &#181;l CuSO4-5H2O per 500 &#181;l of lysate and vortex briefly. Incubate at room temperature for 30 min.</li>
	<li>Add 4 sample volumes of acetone chilled to &#8722;20 &#176;C. Vortex the samples and incubate overnight at &#8722;80 &#176;C to completely precipitate the proteins and remove unreacted biotin-azide.</li>
	<li>Centrifuge the samples at 17,000 x g for 15 min at 4 &#176;C to pellet the precipitated proteins.</li>
	<li>Aspirate the supernatant completely, and resolubilize the proteins by sonication in 150 &#181;l PBS (pH 7.4) + 1% SDS.</li>
	<li>Add 600 &#181;l of PBS (pH 7.4) to dilute the concentration of SDS to 0.2%.</li>
	<li>Add the samples to 30 &#181;l pre-washed high-capacity streptavidin agarose beads and incubate 1 hr at 4 &#176;C with rotation.</li>
	<li>Pellet the beads by centrifugation at 1,000 x g for 3 min. Aspirate the supernatant containing unbound proteins and discard.</li>
	<li>Add 1 ml of wash buffer (400 mM NaCl, 50 mM Tris, 0.2% SDS, pH 7.4) to the beads and incubate 5 min at room temperature with rotation.</li>
	<li>Repeat steps 4.12 and 4.13 3 times.</li>
	<li>Aspirate the wash buffer completely from the beads, and add 30 &#181;l of 2x SDS sample buffer.</li>
	<li>Incubate for 5 min on a 95 &#176;C heat block to release the proteins from the beads.</li>
	<li>Centrifuge the beads for 1 min at 13,000 x g at room temperature. 
	NOTE: At this point, the samples can be stored at -20 &#176;C until ready to run SDS-PAGE. If freezing samples, reheat for 5 min at 95 &#176;C before use.</li>
	<li>Carefully pipette the sample buffer containing proteins off of the beads, and load onto SDS-PAGE20 (see important considerations below). The beads can be saved for reboiling later if necessary. 
	NOTE: For protein detection by silver stain, better results are obtained using a 1 mm thick gel. If a band is to be cut out of the silver-stained gel for mass spectrometry (MS) analysis, prepare all reagents fresh from sterile solutions and leave an empty well in between samples on the gel. For every band cut from the probe lane, a parallel slice should be taken from the DMSO lane so that background proteins can be subtracted. For validation of protein ID by western blot, it is critical to also load the input sample from step 4.4 when running the SDS-PAGE gel.</li>
	<li>Follow a standard protocol for either silver staining21 or Western blot22 to detect the target protein(s).</li>
</ol>

<ol><li>Schenone, M., Dančìk, V., Wagner, B. K., Clemons, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Target+identification+and+mechanism+of+action+in+chemical+biology+and+drug+discovery%5BTitle%5D)+AND+%22Nat+Chem+Biol%22%5BJournal%5D)">Target identification and mechanism of action in chemical biology and drug discovery.</a> Nat Chem Biol. 9 (4), 232-240 (2013).</li>
<li>Cook, D., Brown, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Lessons+learned+from+the+fate+of+AstraZeneca%27s+drug+pipeline%3A+a+five-dimensional+framework%5BTitle%5D)+AND+%22Nat+Rev+Drug+Discov%22%5BJournal%5D)">Lessons learned from the fate of AstraZeneca's drug pipeline: a five-dimensional framework.</a> Nat Rev Drug Discov. 13 (6), 419-431 (2014).</li>
<li>Titov, D. V., Liu, J. O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Identification+and+validation+of+protein+targets+of+bioactive+small+molecules%5BTitle%5D)+AND+%22Bioorg+Med+Chem%22%5BJournal%5D)">Identification and validation of protein targets of bioactive small molecules.</a> Bioorg Med Chem. 20 (6), 1902-1909 (2012).</li>
<li>Jung, H. J., Kwon, H. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Target+deconvolution+of+bioactive+small+molecules%3A+the+heart+of+chemical+biology+and+drug+discovery%5BTitle%5D)+AND+%22Arch+Pharm+Res%22%5BJournal%5D)">Target deconvolution of bioactive small molecules: the heart of chemical biology and drug discovery.</a> Arch Pharm Res. 38 (9), 1627-1641 (2015).</li>
<li>Sumranjit, J., Chung, S. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Recent+Advances+in+Target+Characterization+and+Identification+by+Photoaffinity+Probes%5BTitle%5D)+AND+%22Molecules%22%5BJournal%5D)">Recent Advances in Target Characterization and Identification by Photoaffinity Probes.</a> Molecules. 18 (9), 10425-10451 (2013).</li>
<li>Robles, O., Romo, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Chemo-+and+site-selective+derivatizations+of+natural+products+enabling+biological+studies%5BTitle%5D)+AND+%22Nat+Prod+Rep%22%5BJournal%5D)">Chemo- and site-selective derivatizations of natural products enabling biological studies.</a> Nat Prod Rep. 31 (3), 318-334 (2014).</li>
<li>Zhou, Q., Gui, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Bioconjugation+by+Native+Chemical+Tagging+of+C-H+Bonds%5BTitle%5D)+AND+%22J+Am+Chem+Soc%22%5BJournal%5D)">Bioconjugation by Native Chemical Tagging of C-H Bonds.</a> J Am Chem Soc. 135 (35), (2013).</li>
<li>Li, Z., Hao, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Design+and+Synthesis+of+Minimalist+Terminal+Alkyne-Containing+Diazirine+Photo-Crosslinkers+and+Their+Incorporation+into+Kinase+Inhibitors+for+Cell-+and+Tissue-Based+Proteome+Profiling%5BTitle%5D)+AND+%22Angew+Chem+Int+Edit%22%5BJournal%5D)">Design and Synthesis of Minimalist Terminal Alkyne-Containing Diazirine Photo-Crosslinkers and Their Incorporation into Kinase Inhibitors for Cell- and Tissue-Based Proteome Profiling.</a> Angew Chem Int Edit. 52 (33), 8551-8556 (2013).</li>
<li>Chamni, S., He, Q. L., Dang, Y., Bhat, S., Liu, J. O., Romo, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Diazo+reagents+with+small+steric+footprints+for+simultaneous+arming%2FSAR+studies+of+alcohol-containing+natural+products+via+O-H+insertion%5BTitle%5D)+AND+%22ACS+chem+biol%22%5BJournal%5D)">Diazo reagents with small steric footprints for simultaneous arming/SAR studies of alcohol-containing natural products via O-H insertion.</a> ACS chem biol. 6 (11), 1175-1181 (2011).</li>
<li>Mackinnon, A. L., Taunton, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Target+Identification+by+Diazirine+Photo-Cross-linking+and+Click+Chemistry%5BTitle%5D)+AND+%22Curr+Protoc+Chem+Biol%22%5BJournal%5D)">Target Identification by Diazirine Photo-Cross-linking and Click Chemistry.</a> Curr Protoc Chem Biol. 1, 55-73 (2009).</li>
<li>Smith, E., Collins, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Photoaffinity+labeling+in+target-+and+binding-site+identification%5BTitle%5D)+AND+%22Future+med+chem%22%5BJournal%5D)">Photoaffinity labeling in target- and binding-site identification.</a> Future med chem. 7 (2), 159-183 (2015).</li>
<li>Huisgen, R., Szeimies, G., Möbius, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((1.3-Dipolare+Cycloadditionen%2C+XXXII.+Kinetik+der+Additionen+organischer+Azide+an+CC-Mehrfachbindungen%5BTitle%5D)+AND+%22Chem+Ber%22%5BJournal%5D)">1.3-Dipolare Cycloadditionen, XXXII. Kinetik der Additionen organischer Azide an CC-Mehrfachbindungen.</a> Chem Ber. 100 (8), 2494-2507 (1967).</li>
<li>Kolb, H. C., Finn, M. G., Sharpless, K. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Click+Chemistry%3A+Diverse+Chemical+Function+from+a+Few+Good+Reactions%5BTitle%5D)+AND+%22Angew+Chem+Int+Ed+Engl%22%5BJournal%5D)">Click Chemistry: Diverse Chemical Function from a Few Good Reactions.</a> Angew Chem Int Ed Engl. 40 (11), 2004-2021 (2001).</li>
<li>Rostovtsev, V. V., Green, L. G., Fokin, V. V., Sharpless, K. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+stepwise+huisgen+cycloaddition+process%3A+copper%28I%29-catalyzed+regioselective+%26%2334%3Bligation%26%2334%3B+of+azides+and+terminal+alkynes%5BTitle%5D)+AND+%22Angew+Chem+Int+Ed+Engl%22%5BJournal%5D)">A stepwise huisgen cycloaddition process: copper(I)-catalyzed regioselective &#34;ligation&#34; of azides and terminal alkynes.</a> Angew Chem Int Ed Engl. 41 (14), 2596-2599 (2002).</li>
<li>Tornøe, C. W., Christensen, C., Meldal, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Peptidotriazoles+on+Solid+Phase%3A+%5B1%2C2%2C3%5D-Triazoles+by+Regiospecific+Copper%28I%29-Catalyzed+1%2C3-Dipolar+Cycloadditions+of+Terminal+Alkynes+to+Azides%5BTitle%5D)+AND+%22J+Org+Chem%22%5BJournal%5D)">Peptidotriazoles on Solid Phase: [1,2,3]-Triazoles by Regiospecific Copper(I)-Catalyzed 1,3-Dipolar Cycloadditions of Terminal Alkynes to Azides.</a> J Org Chem. 67 (9), 3057-3064 (2002).</li>
<li>Head, S. A., Shi, W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Antifungal+drug+itraconazole+targets+VDAC1+to+modulate+the+AMPK%2FmTOR+signaling+axis+in+endothelial+cells%5BTitle%5D)+AND+%22P+Natl+Acad+Sci+USA%22%5BJournal%5D)">Antifungal drug itraconazole targets VDAC1 to modulate the AMPK/mTOR signaling axis in endothelial cells.</a> P Natl Acad Sci USA. 112 (52), E7276-E7285 (2015).</li>
<li>Shi, W., Nacev, B. A., Aftab, B. T., Head, S., Rudin, C. M., Liu, J. O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Itraconazole+Side+Chain+Analogues%3A+Structure-Activity+Relationship+Studies+for+Inhibition+of+Endothelial+Cell+Proliferation%2C+Vascular+Endothelial+Growth+Factor+Receptor+2+%28VEGFR2%29+Glycosylation%2C+and+Hedgehog+Signaling%5BTitle%5D)+AND+%22J+Med+Chem%22%5BJournal%5D)">Itraconazole Side Chain Analogues: Structure-Activity Relationship Studies for Inhibition of Endothelial Cell Proliferation, Vascular Endothelial Growth Factor Receptor 2 (VEGFR2) Glycosylation, and Hedgehog Signaling.</a> J Med Chem. 54 (20), 7363-7374 (2011).</li>
<li>Shi, W., Nacev, B. A., Bhat, S., Liu, J. O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Impact+of+Absolute+Stereochemistry+on+the+Antiangiogenic+and+Antifungal+Activities+of+Itraconazole%5BTitle%5D)+AND+%22ACS+Med+Chem+Lett%22%5BJournal%5D)">Impact of Absolute Stereochemistry on the Antiangiogenic and Antifungal Activities of Itraconazole.</a> ACS Med Chem Lett. 1 (4), 155-159 (2010).</li>
<li>Lowry, O. H., Rosebrough, N. J., Farr, A. L., Randall, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Protein+measurement+with+the+Folin+phenol+reagent%5BTitle%5D)+AND+%22J+Biol+Chem%22%5BJournal%5D)">Protein measurement with the Folin phenol reagent.</a> J Biol Chem. 193 (1), 265-275 (1951).</li>
<li>Shapiro, A. L., Viñuela, E., Maizel, J. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Molecular+weight+estimation+of+polypeptide+chains+by+electrophoresis+in+SDS-polyacrylamide+gels%5BTitle%5D)+AND+%22Biochem+Bioph+Res+Co%22%5BJournal%5D)">Molecular weight estimation of polypeptide chains by electrophoresis in SDS-polyacrylamide gels.</a> Biochem Bioph Res Co. 28 (5), 815-820 (1967).</li>
<li>Switzer, R. C., Merril, C. R., Shifrin, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+highly+sensitive+silver+stain+for+detecting+proteins+and+peptides+in+polyacrylamide+gels%5BTitle%5D)+AND+%22Anal+Biochem%22%5BJournal%5D)">A highly sensitive silver stain for detecting proteins and peptides in polyacrylamide gels.</a> Anal Biochem. 98 (1), 231-237 (1979).</li>
<li>Towbin, H., Staehelin, T., Gordon, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Electrophoretic+transfer+of+proteins+from+polyacrylamide+gels+to+nitrocellulose+sheets%3A+procedure+and+some+applications%5BTitle%5D)+AND+%22P+Natl+Acad+Sci+USA%22%5BJournal%5D)">Electrophoretic transfer of proteins from polyacrylamide gels to nitrocellulose sheets: procedure and some applications.</a> P Natl Acad Sci USA. 76 (9), 4350-4354 (1979).</li>
</ol>

The authors have nothing to disclose.

Different approaches to identifying the targets of small molecules can be broadly grouped into two categories: top-down, where the cellular phenotype of the drug is used to narrow down its potential targets based on their known functions, or bottom-up, where the target is identified directly by chemical or genetic means3. Top-down or phenotypic studies can identify certain cellular processes affected by the drug (e.g., transcription/translation/DNA synthesis, cell cycle block, signaling pathway activa...

Bioactive small molecules fundamentally work by interacting with and altering the function of one or more &#34;target&#34; molecules, most commonly proteins, in the cell. In drug discovery, when an active compound is discovered through phenotypic screening, identification of the molecular target(s) of that compound is crucial, not just for understanding the mechanism of action and potential side-effects of the compound, but also for potentially discovering new biology underlying the disease model and paving the way for development of new mechanistic classes of therapeutics1. Although target identification is not required for a drug to be used therapeutically, in recent years there has been an increasing recognition that novel drug candidates are more likely to succeed in clinical trials, and therefore yield better returns on investment, if a validated target is known2. Thus, there has been a growing interest in methods for identifying small molecule target proteins.
A classical target identification experiment typically relies on affinity purification, where the small molecule of interest is immobilized onto a resin and incubated with whole cell lysates, after which unbound proteins are washed away and the remaining proteins are eluted and identified3. While this technique has been used to identify the targets of many small molecules4, it is unsuitable as a universal target ID method for several reasons. First, the target protein must retain its native conformation upon cell lysis in order to retain its ability to bind to the small molecule. This can be particularly problematic for membrane proteins, which often undergo conformational changes after being removed from their native environment, or simply aggregate and precipitate out of solution. Second, the small molecule must be chemically modified in such a way that it can be immobilized onto the resin while maintaining its ability to bind the target protein. Deep binding pockets may therefore become inaccessible to a small molecule once it is fixed to the resin. Third, the binding affinity must be sufficiently high that the interaction is maintained during the washing steps, making identification of lower affinity interactions challenging. Fourth, environmental conditions such as pH, ion concentration, or the presence of other endogenous molecules can vary spatially within the cell and are sometimes prerequisites for drug-target interactions. Thus, finding the correct conditions to allow and maintain binding outside of the cell can require a significant amount of trial and error.
Photoaffinity labeling circumvents these issues by allowing the covalent binding of a small molecule and its target within the native context of a cell. Rather than immobilizing the small molecule to a large bulky resin, the molecule is instead chemically modified to install two small functional groups: a photoactivatable moiety which allows covalent crosslinking to the target protein when irradiated with a particular wavelength of light, and a reporter group that allows the target protein to be detected and subsequently isolated. Live cells are treated with the photoaffinity probe, the probe binds and covalently crosslinks to the target protein, and the probe-protein complex is then isolated intact. The specificity of probe binding to the target is demonstrated by performing a competition experiment in parallel, where an excess of the parent compound is used to compete away binding of the probe to the target protein.
The design and synthesis of photoaffinity probes varies greatly from one small molecule to another, and will not be covered in this protocol; however, several excellent discussions on the subject have been published5-9. The main consideration is that the probe retains the bioactivity of the parent compound, therefore presumably binding to the same target(s). Structure-activity relationship (SAR) studies must be performed to determine which parts of the molecule can be modified without loss of bioactivity. A variety of different chemical groups have been used as photoactivatable crosslinkers, including diazirine, benzophenone, and aryl azide, which each have advantages and disadvantages10. Likewise, there are multiple reporter tags that have been used to isolate probe-binding proteins. Reporter groups may be functional on their own, such as the commonly used biotin or fluorescent tags, or may be precursors that require further functionalization subsequent to the photocrosslinking step, which have the advantage of being smaller and thus less likely to compromise bioactivity11.
In this protocol, we have used a photoaffinity probe containing a diazirine photocrosslinking group, and a terminal alkyne for the attachment of a reporter group through a Cu(I)-catalyzed Azide-Alkyne Sharpless-H&#252;isgen cycloaddition (or click) reaction12-15. The SAR studies, probe design and synthesis, and results of these studies have been published elsewhere16-18.

<table><tbody><tr><td>Tris(2-carboxyethyl)phosphine (TCEP)</td><td>Life Technologies</td><td>20490</td><td>Make fresh day of use. Prepare 100 mM stock in water with 4 eq NaOH.</td></tr><tr><td>Tris[(1-benzyl-1H-1,2,3-triazol-4-yl)methyl] amine (TBTA)</td><td>AnaSpec</td><td>63360-50&amp;nbsp;</td><td>Prepare 1.7 mM stock in a 4:1 ratio of t-butanol to DMSO, store at -20&amp;nbsp;&amp;deg;C.</td></tr><tr><td>Copper Sulfate (CuSO&lt;sub&gt;4&lt;/sub&gt;&amp;#8226;5H&lt;sub&gt;2&lt;/sub&gt;O)</td><td>LabChem, Inc.</td><td>LC13440-1</td><td>Prepare 50 mM stock in water, store at room temperature.</td></tr><tr><td>Biotin-azide</td><td>Click Chemistry Tools</td><td>AZ104-100</td><td>Prepare 10 mM stock in DMSO, store at -20&amp;nbsp;&amp;deg;C.</td></tr><tr><td>Alexa Fluor 647-azide&amp;nbsp;</td><td>Life Technologies</td><td>A10277</td><td>Prepare 1 mM stock in DMSO, store at -20&amp;nbsp;&amp;deg;C.</td></tr><tr><td>365 nm UV lamp</td><td>Spectroline</td><td>FC100</td><td>UV-blocking glasses should be worn while operating.</td></tr><tr><td>Protease inhibitor tablets, EDTA-free</td><td>Roche Life Science</td><td>11873580001</td><td>Prepare 50x solution in water and store at -20&amp;nbsp;&amp;deg;C.</td></tr><tr><td>Sonicator</td><td>Branson</td><td>Sonifier 250</td><td>Set to output 1, duty 30%.&amp;nbsp;</td></tr><tr><td>Fluorescent gel scanner</td><td>GE Healthcare Life Sciences</td><td>FLA 9500</td><td>Use red laser to detect Alexa-fluor 647.</td></tr><tr><td>Detergent-compatible Dc protein assay kit</td><td>Bio-Rad</td><td>5000112</td><td></td></tr><tr><td>High Capacity Streptavidin Agarose beads&amp;nbsp;</td><td>Life Technologies</td><td>20359</td><td></td></tr><tr><td>Dulbecco&#039;s Modified Eagles Medium, low glucose</td><td>ThermoFisher Scientific</td><td>11885092</td><td></td></tr><tr><td>Fetal Bovine Serum, qualified</td><td>ThermoFisher Scientific</td><td>26140079</td><td></td></tr><tr><td>Penicillin/Streptomycin solution</td><td>ThermoFisher Scientific</td><td>15140122</td><td></td></tr><tr><td>SDS Sample Buffer (2x)</td><td>ThermoFisher Scientific</td><td>LC2676</td><td></td></tr></tbody></table>

identification of small molecule-binding proteins in a native cellular environment by live-cell photoaffinity labeling

Identifying the molecular target(s) of small molecules is a challenging but necessary step towards understanding their mechanism of action. While several target identification methods have been developed and used to successfully elucidate the binding proteins of a variety of small molecules, these techniques have drawbacks that make them unsuitable for detecting certain types of small molecule-target interactions. In particular, non-covalent interactions that depend on native cellular conditions, such as those of membrane proteins whose structures may be perturbed upon cell lysis, are often not amenable to affinity-based target identification methods. Here, we demonstrate a method wherein a probe containing a photolabile group is used to covalently crosslink to the small molecule binding protein within the environment of the live cell, allowing the detection and isolation of the target protein without the need for maintenance of the interaction after cell lysis. This technique is a valuable tool for studying biologically interesting small molecules with unknown mechanisms, both in the context of basic biology as well as drug discovery.

The results shown here were obtained with a photo-affinity probe of the antifungal drug itraconazole, the use of which has been previously published16. These results demonstrate the use of the live-cell photoaffinity labeling technique to successfully identify a major itraconazole-binding protein as the 35 kDa membrane protein Voltage-Dependent Anion Channel 1 (VDAC1).
The above protocol was performed in HEK293T cells...

Watch this Scientific Journal Video about Identification of Small Molecule-binding Proteins in a Native Cellular Environment by Live-cell Photoaffinity Labeling at JoVE.com

Identification of Small Molecule-binding Proteins in a Native Cellular Environment by Live-cell Photoaffinity Labeling

We describe here a method for identification of small molecule-binding proteins using photoaffinity labeling. The advantage of this technique is that binding and covalent labeling of the target proteins occurs within the live cellular environment, removing the risk of disrupting native protein structure and binding conditions upon cell lysis.

Identifying the molecular target(s) of small molecules is a challenging but necessary step towards understanding their mechanism of action. While several ...

identification-small-molecule-binding-proteins-native-cellular

Research

JoVE Journal

Biochemistry

12.6K Views.  Johns Hopkins University School of Medicine. We describe here a method for identification of small molecule-binding proteins using photoaffinity labeling. The advantage of this technique is that binding and covalent labeling of the target proteins occurs within the live cellular environment, removing the risk of disrupting native protein structure and binding conditions upon cell lysis. 

Video: Identification of Small Molecule-binding Proteins in a Native Cellular Environment by Live-cell Photoaffinity Labeling

Detection and Visualization of DNA Damage-induced Protein Complexes in Suspension Cell Cultures Using the Proximity Ligation Assay

The DNA damage response orchestrates the repair of DNA lesions that occur spontaneously, are caused by genotoxic stress, or appear in the context of programmed DNA breaks in lymphocytes. The Ataxia-Telangiectasia Mutated kinase (ATM), ATM- and Rad3-Related kinase (ATR) and the catalytic subunit of DNA-dependent Protein Kinase (DNA-PKcs) are among the first that are activated upon induction of DNA damage, and are central regulators of a network that controls DNA repair, apoptosis and cell survival. As part of a tumor-suppressive pathway, ATM and ATR activate p53 through phosphorylation, thereby regulating the transcriptional activity of p53. DNA damage also results in the formation of so-called ionizing radiation-induced foci (IRIF) that represent complexes of DNA damage sensor and repair proteins that accumulate at the sites of DNA damage, which are visualized by fluorescence microscopy. Co-localization of proteins in IRIFs, however, does not necessarily imply direct protein-protein interactions, as the resolution of fluorescence microscopy is limited. 
In situ Proximity Ligation Assay (PLA) is a novel technique that allows the direct visualization of protein-protein interactions in cells and tissues with unprecedented specificity and sensitivity. This technique is based on the spatial proximity of specific antibodies binding to the proteins of interest. When the interrogated proteins are within ~40 nm an amplification reaction is triggered by oligonucleotides that are conjugated to the antibodies, and the amplification product is visualized by fluorescent labeling, yielding a signal that corresponds to the subcellular location of the interacting proteins. Using the established functional interaction between ATM and p53 as an example, it is demonstrated here how PLA can be used in suspension cell cultures to study the direct interactions between proteins that are integral parts of the DNA damage response.

Here, it is demonstrated how the in situ Proximity Ligation Assay (PLA) can be used to detect and visualize the direct protein-protein interactions between ATM and p53 in suspension cell cultures exposed to genotoxic stress.

The DNA damage response orchestrates the repair of DNA lesions that occur spontaneously, are caused by genotoxic stress, or appear in the context of ...

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 ...

Optimizing the Genetic Incorporation of Chemical Probes into GPCRs for Photo-crosslinking Mapping and Bioorthogonal Chemistry in Live Mammalian Cells

The genetic incorporation of non-canonical amino acids (ncAAs) via amber stop codon suppression is a powerful technique to install artificial probes and reactive moieties onto proteins directly in the live cell. Each ncAA is incorporated by a dedicated orthogonal suppressor-tRNA/amino-acyl-tRNA-synthetase (AARS) pair that is imported into the host organism. The incorporation efficiency of different ncAAs can greatly differ, and be unsatisfactory in some cases. Orthogonal pairs can be improved by manipulating either the AARS or the tRNA. However, directed evolution of tRNA or AARS using large libraries and dead/alive selection methods are not feasible in mammalian cells. Here, a facile and robust fluorescence-based assay to evaluate the efficiency of orthogonal pairs in mammalian cells is presented. The assay allows screening tens to hundreds of AARS/tRNA variants with a moderate effort and within a reasonable time. Use of this assay to generate new tRNAs that significantly improve the efficiency of the pyrrolysine orthogonal system is described, along with the application of ncAAs to the study of G-protein coupled receptors (GPCRs), which are challenging objects for ncAA mutagenesis. First, by systematically incorporating a photo-crosslinking ncAA throughout the extracellular surface of a receptor, binding sites of different ligands on the intact receptor are mapped directly in the live cell. Second, by incorporating last-generation ncAAs into a GPCR, ultrafast catalyst-free receptor labeling with a fluorescent dye is demonstrated, which exploits bioorthogonal strain-promoted inverse Diels Alder cycloaddition (SPIEDAC) on the live cell. As ncAAs can be generally applied to any protein independently on its size, the method is of general interest for a number of applications. In addition, ncAA incorporation does not require any special equipment and is easily performed in standard biochemistry labs.

A facile fluorescence assay is presented to evaluate the efficiency of amino-acyl-tRNA-synthetase/tRNA pairs incorporating non-canonical amino-acids (ncAAs) into proteins expressed in mammalian cells. The application of ncAAs to study G-protein coupled receptors (GPCRs) is described, including photo-crosslinking mapping of binding sites and bioorthogonal GPCR labeling on live cells.

The genetic incorporation of non-canonical amino acids (ncAAs) via amber stop codon suppression is a powerful technique to install artificial probes and ...

Chemistry

High-resolution Spatiotemporal Analysis of Receptor Dynamics by Single-molecule Fluorescence Microscopy

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., kinetics, coexistence of different states and populations, transient interactions), which are typically hidden in ensemble measurements, such as those obtained with standard biochemical or microscopy methods. Thus, dynamic events, such as receptor-receptor interactions, can be followed in real time in a living cell with high spatiotemporal resolution. This protocol describes a method based on labeling with small and bright organic fluorophores and total internal reflection fluorescence (TIRF) microscopy to directly visualize single receptors on the surface of living cells. This approach allows one to precisely localize receptors, measure the size of receptor complexes, and capture dynamic events such as transient receptor-receptor interactions. The protocol provides a detailed description of how to perform a single-molecule experiment, including sample preparation, image acquisition and image analysis. As an example, the application of this method to analyze two G-protein-coupled receptors, i.e., &#946;2-adrenergic and &#947;-aminobutyric acid type B (GABAB) receptor, is reported. The protocol can be adapted to other membrane proteins and different cell models, transfection methods and labeling strategies.

This protocol describes how to use total internal reflection fluorescence microscopy to visualize and track single receptors on the surface of living cells and thereby analyze receptor lateral mobility, size of receptor complexes as well as to visualize transient receptor-receptor interactions. This protocol can be extended to other membrane proteins.

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., ...

Bioengineering

Visualizing Protein-DNA Interactions in Live Bacterial Cells Using Photoactivated Single-molecule Tracking

Protein-DNA interactions are at the heart of many fundamental cellular processes. For example, DNA replication, transcription, repair, and chromosome organization are governed by DNA-binding proteins that recognize specific DNA structures or sequences. In vitro experiments have helped to generate detailed models for the function of many types of DNA-binding proteins, yet, the exact mechanisms of these processes and their organization in the complex environment of the living cell remain far less understood. We recently introduced a method for quantifying DNA-repair activities in live Escherichia coli cells using Photoactivated Localization Microscopy (PALM) combined with single-molecule tracking. Our general approach identifies individual DNA-binding events by the change in the mobility of a single protein upon association with the chromosome. The fraction of bound molecules provides a direct quantitative measure for the protein activity and abundance of substrates or binding sites at the single-cell level. Here, we describe the concept of the method and demonstrate sample preparation, data acquisition, and data analysis procedures.

Photoactivated localization microscopy (PALM) combined with single-molecule tracking allows direct observation and quantification of protein-DNA interactions in live Escherichia coli cells.

Protein-DNA interactions are at the heart of many fundamental cellular processes. For example, DNA replication, transcription, repair, and chromosome ...

Immunology and Infection

Photoactivated Localization Microscopy with Bimolecular Fluorescence Complementation (BiFC-PALM)

Protein-protein interactions (PPIs) are key molecular events to biology. However, it remains a challenge to visualize PPIs with sufficient resolution and sensitivity in cells because the resolution of conventional light microscopy is diffraction-limited to ~250 nm. By combining bimolecular fluorescence complementation (BiFC) with photoactivated localization microscopy (PALM), PPIs can be visualized in cells with single molecule sensitivity and nanometer spatial resolution. BiFC is a commonly used technique for visualizing PPIs with fluorescence contrast, which involves splitting of a fluorescent protein into two non-fluorescent fragments. PALM is a recent superresolution microscopy technique for imaging biological samples at the nanometer and single molecule scales, which uses phototransformable fluorescent probes such as photoactivatable fluorescent proteins (PA-FPs). BiFC-PALM was demonstrated by splitting PAmCherry1, a PA-FP compatible with PALM, for its monomeric nature, good single molecule brightness, high contrast ratio, and utility for stoichiometry measurements. When split between amino acids 159 and 160, PAmCherry1 can be made into a BiFC probe that reconstitutes efficiently at 37 &#176;C with high specificity to PPIs and low non-specific reconstitution. Ras-Raf interaction is used as an example to show how BiFC-PALM helps to probe interactions at the nanometer scale and with single molecule resolution. Their diffusion can also be tracked in live cells using single molecule tracking (smt-) PALM. In this protocol, factors to consider when designing the fusion proteins for BiFC-PALM are discussed, sample preparation, image acquisition, and data analysis steps are explained, and a few exemplary results are showcased. Providing high spatial resolution, specificity, and sensitivity, BiFC-PALM is a useful tool for studying PPIs in intact biological samples.

Photoactivated Localization Microscopy with Bimolecular Fluorescence Complementation BiFC-PALM

Protein-protein interactions are visualized in cells with nanometer spatial resolution by combining bimolecular fluorescence complementation (BiFC) with photoactivated localization microscopy (PALM). Described here is the use of BiFC-PALM for imaging Ras-Raf interactions in U2OS cells for visualizing the nanoscale clustering and diffusion of individual Ras-Raf complexes.

Protein-protein interactions (PPIs) are key molecular events to biology. However, it remains a challenge to visualize PPIs with sufficient resolution and ...

Measuring TCR-pMHC Binding In Situ using a FRET-based Microscopy Assay

T-cells are remarkably specific and effective when recognizing antigens in the form of peptides embedded in MHC molecules (pMHC) on the surface of Antigen Presenting Cells (APCs). This is despite T-cell antigen receptors (TCRs) exerting usually a moderate affinity (&#181;M range) to antigen when binding is measured in vitro1. In view of the molecular and cellular parameters contributing to T-cell antigen sensitivity, a microscopy-based methodology has been developed as a means to monitor TCR-pMHC binding in situ, as it occurs within the synapse of a live T-cell and an artificial and functionalized glass-supported planar lipid bilayer (SLB), which mimics the cell membrane of an Antigen presenting Cell (APC) 2. Measurements are based on F&#246;rster Resonance Energy Transfer (FRET) between a blue- and red-shifted fluorescent dye attached to the TCR and the pMHC. Because the efficiency of FRET is inversely proportional to the sixth power of the inter-dye distance, one can employ FRET signals to visualize synaptic TCR-pMHC binding. The sensitive of the microscopy approach supports detection of single molecule FRET events. This allows to determine the affinity and off-rate of synaptic TCR-pMHC interactions and in turn to interpolate the on-rate of binding. Analogous assays could be applied to measure other receptor-ligand interactions in their native environment.

Measuring TCR-pMHC Binding In Situ using a FRET-based Microscopy Assay

This manuscript describes how to conduct (single molecule) F&#246;rster Resonance Energy Transfer (FRET)- based assays to measure the binding dynamics between T-cell antigen receptor (TCR) and antigenic peptide-loaded MHC molecules as they occur within the immunological synapse of a T-cell in contact with a functionalized planar supported lipid bilayer.

T-cells are remarkably specific and effective when recognizing antigens in the form of peptides embedded in MHC molecules (pMHC) on the surface of Antigen ...

Fluorescent Labeling of COS-7 Expressing SNAP-tag Fusion Proteins for Live Cell Imaging

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest in live cells. These systems offer a broad selection of fluorescent substrates optimized for a range of imaging instrumentation. Once cloned and expressed, the tagged protein can be used with a variety of substrates for numerous downstream applications without having to clone again. There are two steps to using this system: cloning and expression of the protein of interest as a SNAP-tag fusion, and labeling of the fusion with the SNAP-tag substrate of choice. The SNAP-tag is a small protein based on human O6-alkylguanine-DNA-alkyltransferase (hAGT), a DNA repair protein. SNAP-tag labels are dyes conjugated to guanine or chloropyrimidine leaving groups via a benzyl linker. In the labeling reaction, the substituted benzyl group of the substrate is covalently attached to the SNAP-tag. CLIP-tag is a modified version of SNAP-tag, engineered to react with benzylcytosine rather than benzylguanine derivatives. When used in conjunction with SNAP-tag, CLIP-tag enables the orthogonal and complementary labeling of two proteins simultaneously in the same cells.

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest in live cells. Once cloned and expressed, the tagged protein can be used with a variety of substrates for numerous downstream applications without having to clone again.

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest ...

Biology

Multi-color Localization Microscopy of Single Membrane Proteins in Organelles of Live Mammalian Cells

Knowledge about the localization of proteins in cellular subcompartments is crucial to understand their specific function. Here, we present a super-resolution technique that allows for the determination of the microcompartments that are accessible for proteins by generating localization and tracking maps of these proteins. Moreover, by multi-color localization microscopy, the localization and tracking profiles of proteins in different subcompartments are obtained simultaneously. The technique is specific for live cells and is based on the repetitive imaging of single mobile membrane proteins. Proteins of interest are genetically fused with specific, so-called self-labeling tags. These tags are enzymes that react with a substrate in a covalent manner. Conjugated to these substrates are fluorescent dyes. Reaction of the enzyme-tagged proteins with the fluorescence labeled substrates results in labeled proteins. Here, Tetramethylrhodamine (TMR) and Silicon Rhodamine (SiR) are used as fluorescent dyes attached to the substrates of the enzymes. By using substrate concentrations in the pM to nM range, sub-stoichiometric labeling is achieved that results in distinct signals. These signals are localized with ~15&#8211;27 nm precision. The technique allows for multi-color imaging of single molecules, whereby the number of colors is limited by the available membrane-permeable dyes and the repertoire of self-labeling enzymes. We show the feasibility of the technique by determining the localization of the quality control enzyme (Pten)-induced kinase 1 (PINK1) in different mitochondrial compartments during its processing in relation to other membrane proteins. The test for true physical interactions between differently labeled single proteins by single molecule FRET or co-tracking is restricted, though, because the low labeling degrees decrease the probability for having two adjacent proteins labeled at the same time. While the technique is strong for imaging proteins in membrane compartments, in most cases it is not appropriate to determine the localization of highly mobile soluble proteins.

Here, we present a protocol for multi-color localization of single membrane proteins in organelles of live cells. To attach fluorophores, self-labeling proteins are used. Proteins, located in different membranes compartments of the same organelle, can be localized with a precision of ~18 nm.

Knowledge about the localization of proteins in cellular subcompartments is crucial to understand their specific function. Here, we present a ...

Live Cell Single-Molecule Imaging-Based Quantification of Protein Interactions

Single-Molecule Measurement of Protein Interaction Dynamics Within Biomolecular Condensates

Biomolecular condensates formed via liquid-liquid phase separation (LLPS) have been considered critical in cellular organization and an increasing number of cellular functions. Characterizing LLPS in live cells is also important because aberrant condensation has been linked to numerous diseases, including cancers and neurodegenerative disorders. LLPS is often driven by selective, transient, and multivalent interactions between intrinsically disordered proteins. Of great interest are the interaction dynamics of proteins participating in LLPS, which are well-summarized by measurements of their binding residence time (RT), that is, the amount of time they spend bound within condensates. Here, we present a method based on live-cell single-molecule imaging that allows us to measure the mean RT of a specific protein within condensates. We simultaneously visualize individual protein molecules and the condensates with which they associate, use single-particle tracking (SPT) to plot single-molecule trajectories, and then fit the trajectories to a model of protein-droplet binding to extract the mean RT of the protein. Finally, we show representative results where this single-molecule imaging method was applied to compare the mean RTs of a protein at its LLPS condensates when fused and unfused to an oligomerizing domain. This protocol is broadly applicable to measuring the interaction dynamics of any protein that participates in LLPS.

Author Spotlight: Evaluation of Protein-Condensate Dynamics in Live Human Cells

Many intrinsically disordered proteins have been shown to participate in the formation of highly dynamic biomolecular condensates, a behavior important for numerous cellular processes. Here, we present a single-molecule imaging-based method for quantifying the dynamics by which proteins interact with each other in biomolecular condensates in live cells.

Biomolecular condensates formed via liquid-liquid phase separation (LLPS) have been considered critical in cellular organization and an increasing number ...

Identification of Small Molecule-binding Proteins in a Native Cellular Environment by Live-cell Photoaffinity Labeling

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