This work was supported by the National Science Foundation Graduate Research Fellowship under Grant No. DGE-1745301 (S.Y.), Pew-Stewart Scholar Award (S.C.), Searle Scholar Award (S.C.), the Shurl and Kay Curci Foundation Research Grant (S.C.), Merkin Innovation Seed Grant (S.C.), the Mallinckrodt Research Grant (S.C.), and the Margaret E. Early Medical Research Trust 2024 Grant (S.C.). S.C. is also supported by the NIH/NCI under Award Number P30CA016042.

1. Labeling of proteins in cells
<ol>
	<li>Express the protein of interest fused to HaloTag in the desired cell line.</li>
	<li>Stably express Halo-tagged H2B in the same type of cells as in 1.1 using transposons or viral transduction.</li>
</ol>
2. Preparation of coverslips
<ol>
	<li>Before using coverslips for cell culture, clean coverslips to remove autofluorescent contaminants.
	<ol>
		<li>Mount 25 mm diameter, #1.5 coverslips on a ceramic staining rack and place into a polypropylene container.</li>
		<li>Completely immerse coverslips in a 1 M solution of KOH and sonicate for 1 h.</li>
		<li>Rinse coverslips three times with double-distilled water (ddH2O).</li>
		<li>Completely immerse coverslips in 100% ethanol and sonicate for 1 h.</li>
		<li>Completely immerse coverslips in 20% ethanol diluted with ddH2O and store at 4 &#176;C until use.</li>
	</ol>
	</li>
</ol>
3. Preparation of cells for microscopy
NOTE: Perform steps from this section in a biosafety cabinet to prevent contamination of the cells.
<ol>
	<li>Plate cells on cleaned coverslips. Perform 2 days before imaging if the Halo-tagged protein is to be transiently expressed. Perform 1 day before imaging if the Halo-tagged protein is stably or endogenously expressed.
	<ol>
		<li>Use sterile forceps to place coverslips in a 6-well plate (one coverslip/well per cell sample) and allow residual ethanol to evaporate.</li>
		<li>Plate each well with an appropriate number of cells to achieve 70% confluency by the time of imaging. Plate an additional well with cells stably expressing H2B-Halo with the same target confluency.</li>
		<li>Follow this step only if transiently expressing the Halo-tagged protein of interest. Incubate cells for 24 h at 37 &#176;C and 5% CO2. Then, transfect the cells with the plasmid encoding the tagged protein of interest using appropriate transfection reagents and following the manufacturer&#39;s guidelines.</li>
		<li>Incubate the cells for 24 h at 37 &#176;C and 5% CO2.</li>
	</ol>
	</li>
	<li>Stain the cells expressing the Halo-tagged protein of interest with both a non-photoactivatable Halo ligand (e.g., JFX549) and a photoactivatable Halo ligand (e.g., PA-JF646). Stain cells expressing H2B-Halo with only the photoactivatable Halo ligand. 
	NOTE: Concentrations of Halo ligands listed here should be used as a starting point; the optimal concentration will depend on the expression level and in-condensate local concentration of the protein of interest.
	<ol>
		<li>For each cell sample expressing the Halo-tagged protein of interest, dilute 100 nM JFX549 Halo ligand35 and 20 nM PA-JF646 Halo ligand36 in 500 &#181;L of appropriate cell media. For each sample expressing H2B-Halo, dilute only 20 nM PA-JF646 Halo ligand in 500 &#181;L of appropriate cell media.</li>
		<li>For each well, aspirate existing media, add 2 mL of 1x PBS, then aspirate PBS (this is referred to as a &#34;rinse&#34; for the remainder of the protocol), and replace with media containing the appropriate Halo ligands.</li>
		<li>Incubate for 1 h at 37 &#176;C and 5% CO2.</li>
		<li>Rinse with PBS four times and replace with fresh media containing no Halo ligands.</li>
		<li>Incubate for 15 min at 37 &#176;C and 5% CO2.</li>
		<li>Repeat steps 3.2.4 and 3.2.5 three times (four rinse-incubation cycles total).</li>
		<li>Use sterile forceps to transfer the coverslip with cells to a cell-culture coverslip chamber compatible with the microscope stage.</li>
		<li>Add phenol red-free media to the chamber and proceed to imaging. Incubate the samples not immediately being imaged at 37 &#176;C and 5% CO2 until needed.</li>
	</ol>
	</li>
</ol>
4. Single-molecule imaging
NOTE: Independent experiments measuring the residence times of both H2B and the protein of interest should be conducted across multiple (&#8805;3) days to generate statistically significant results. Before imaging cell samples on the microscope, align the two cameras using 0.1 &#181;m stained microspheres (Table of Materials) or a similar calibration standard.
<ol>
	<li>Prepare the microscope.
	<ol>
		<li>Turn on the microscope and the computer that controls the microscope.</li>
		<li>Turn on live-cell incubation components (heater and CO2) on the microscope and set it to 37 &#176;C and 5% CO2. Allow the system to equilibrate.</li>
		<li>Put a drop of the appropriate oil onto a 100x TIRF oil immersion objective on the microscope and load the cell sample onto the microscope stage.</li>
	</ol>
	</li>
	<li>Identify a cell to image.
	<ol>
		<li>Image the cell sample under brightfield illumination and adjust the z-position until the cells are in focus.</li>
		<li>Identify a cell that exhibits morphology characteristics of healthy cells.</li>
		<li>Crop the field of view (FOV) so that it only captures the target cell nucleus.</li>
		<li>Use laser illumination and adjust the TIRF angle until HILO illumination41 with optimal signal-to-noise ratio of single molecules is achieved.</li>
	</ol>
	</li>
	<li>Image H2B-Halo in live cells. 
	NOTE: The laser powers used in this section are specific to this experiment and included as an example. Appropriate laser powers should be chosen according to the criteria listed in the discussion.
	<ol>
		<li>Set up an acquisition configuration as follows. Set the exposure time as 500 ms. During each 500 ms exposure, excite PA-JF646 labeled H2B-Halo molecules with a 9.1 mW, 640 nm beam. During the dead time between frames (158.01 &#181;s on the imaging system used here), photoactivate the molecules with a 111 &#181;W, 405 nm beam.</li>
		<li>Capture 2000 frames continuously under these settings. Gradually increase the 405 nm beam&#39;s power over the course of the acquisition when the number of photoactivated molecules becomes insufficient.</li>
	</ol>
	</li>
	<li>Image the Halo-tagged protein of interest in live cells.
	<ol>
		<li>Use a long-pass dichroic mirror to split emission wavelengths between two cameras, one to detect PA-JF646 and the other to detect JFX549. Use appropriate emission filters in front of the cameras.</li>
		<li>Use the same acquisition configuration described in 4.3.1 with the following modification. Add an additional JFX549 channel to track locations of the Halo-tagged protein condensates over time. Every 10 s, acquire one frame (500 ms, 2.1 mW, 561 nm excitation) in the JFX549 channel while keeping the acquisition of the PA-JF646 channel. This will cause bleedthrough into the PA-JF646 channel, which will be taken care of in the post-acquisition data analysis.</li>
		<li>Capture 2000 frames continuously under these settings. Gradually increase the 405 nm beam&#39;s power over the course of the acquisition when the number of photoactivated molecules becomes insufficient.</li>
	</ol>
	</li>
</ol>
5. Analysis of single-molecule imaging data
NOTE: The parameters used throughout section 5 are specific to this experiment and included as an example. Appropriate parameters should be chosen according to the criteria listed in the discussion.
<ol>
	<li>Prepare raw imaging data for processing.
	<ol>
		<li>For the data of the protein of interest, convert each channel from the raw imaging data file into an independent .tif file using ImageJ or similar image processing software. For the data of H2B, convert the single channel from the raw imaging data file into a .tif file in the same way. 
		NOTE: Depending on the acquisition software being used, there can be empty frames in the condensate movie, as only one condensate-tracking frame is taken every 10 s. The empty frames should be populated with the most recent condensate frame (some acquisition software automatically does this). Additionally, if there is significant bleedthrough from the condensate (JFX549) channel into the single-molecule (PA-JF646) channel during those condensate-tracking frames, the corresponding single-molecule frames should be replaced with the most recent single-molecule frame.</li>
		<li>If there are any frames that need to be replaced in either movie due to the above reasons, replace them with the most recent frame from that movie by modifying (if necessary) and running pretracking_comb.txt, an ImageJ macro available in Chong et al.1, and following the prompts.</li>
	</ol>
	</li>
	<li>Generate single-particle trajectories using an SPT software, e.g., SLIMfast39, a GUI implementation of the MTT algorithm38 available in Teves et al.42.
	<ol>
		<li>Load the file from 5.1.2 containing the single-molecule movie in SLIMfast by clicking on Load &#62; Imagestack.</li>
		<li>Set the parameters (localization error rate: 10-6; deflation loops: 3) for localization by clicking OPT &#62; Sheet: Localization.</li>
		<li>Set the parameters (peak emission: 664 nm; lag time: 500 ms) for acquisition by clicking OPT &#62; Sheet: Acquisition.</li>
		<li>Visualize the localizations of all the molecules in a single frame to ensure that the parameters chosen are appropriate (TEST LOC). If inappropriate, modify parameters in steps 5.2.2 and 5.2.3 and repeat this step.</li>
		<li>Generate a file containing the localizations of all the molecules in every frame (LOC ALL).</li>
		<li>Load the file from 5.2.5&#160;in SLIMfast by clicking Load &#62; Particle Data &#62; SLIMfast.</li>
		<li>Set the parameters (maximum expected diffusion coefficient: 0.1 &#181;m2/s; maximum number of competitors: 5) for trajectory generation by clicking OPT &#62; Sheet: Tracking.</li>
		<li>Generate a file containing the trajectories of all the molecules (GEN TRAJ).</li>
	</ol>
	</li>
	<li>Filter out the trajectories that are shorter than tmin, 2.5 s, using evalSPT39, available in Drosopoulos et al.43, to account for tracking errors that result in artificially short trajectories.
	<ol>
		<li>Load the file from 5.2.8 into evalSPT (+&#160;&#62; file from 5.2.8 &#62;&#160;OK).</li>
		<li>Set the parameters (min: 5 frames; max: maximum trajectory length for file from 5.2.8) to filter out the trajectories shorter than 2.5 s.</li>
		<li>Generate a file with all the filtered trajectories (EXPORT DATA).</li>
	</ol>
	</li>
	<li>Follow this step for the trajectories of the protein of interest only. Sort the trajectories into those in the condensates and out of the condensates, then extract the mean in-condensate residence time. 
	NOTE: All the codes for this section are available in Chong et al.1.
	<ol>
		<li>Threshold all the frames of the movie acquired in the JFX549 channel to generate a time-lapse movie populated with the time-evolving binary mask highlighting condensate locations by running the ImageJ macro, nucleus, and cluster mask_v2.txt, and following the prompts.</li>
		<li>Reformat the single-molecule trajectories generated in 5.3.3. using the MATLAB script, ConvertASCII_SlowTracking_css3.m, and adjusting the filenames, paths, and parameters as necessary. (Parameters: Exposure, 0.5 s; Resolution, 0.16 &#181;m/pixel).</li>
		<li>Sort trajectories based on the fraction of the lifetime a given molecule spends in a condensate, F, using the MATLAB script, categorization_v4.m, and adjusting the filenames and paths as necessary.</li>
		<li>Proceed using only in-condensate trajectories.</li>
	</ol>
	</li>
	<li>Extract kobs,s and kpb and compute the corrected mean residence time, <img alt="Equation 6" src="/files/ftp_upload/66169/66169eq05.jpg" style="margin: auto;" />.
	<ol>
		<li>Extract kobs,s&#160;from the in-condensate protein of interest trajectories using the MATLAB script, PLOT_ResidenceHist_css.m, and adjusting filenames, paths, and parameters as necessary. 
		(Parameters: Exposure, 0.5 s; StartFrameForFit, 5 frames).</li>
		<li>Extract kpb from the H2B trajectories using the MATLAB script, PLOT_ResidenceHist_css.m, and adjusting filenames, paths, and parameters as necessary. 
		(Parameters: Exposure, 0.5 s; StartFrameForFit, 5 frames).</li>
		<li>Calculate the corrected mean residence time of the protein of interest specifically bound to its condensates, <img alt="Equation 6" src="/files/ftp_upload/66169/66169eq05.jpg" style="margin: auto;" />, as 
		<img alt="Equation 7" src="/files/ftp_upload/66169/66169eq07v3.jpg" style="margin: auto;" /> 
		NOTE: Controls should be performed to verify that different exposure times that are long enough to blur out mobile particles, for example, 300 ms and 800 ms, still result in the same mean residence time of the protein of interest.</li>
	</ol>
	</li>
</ol>

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

The authors have nothing to disclose.

The protocol as presented here is designed for systems like those investigated in Irgen-Gioro et al.40. Depending on the application, some components of the protocol can be modified, e.g., the method for generating fluorescently labeled cell lines, the fluorescent labeling system, and the style of coverslip used. Halo-tagging of a protein in a cell can be done using two strategies, depending on which is more suitable for a given experiment. 1) Exogenous expression: fusing the protein of interest t...

A growing body of work suggests that biomolecular condensates play an important role in cellular organization and numerous cellular functions, e.g., transcriptional regulation1,2,3,4,5, DNA damage repair6,7,8, chromatin organization9,10,11,12, X-chromosome inactivation13,14,15, and intracellular signaling16,17,18. In addition, the dysregulation of biomolecular condensates is implicated in many diseases, including cancers19,20,21 and neurodegenerative disorders22,23,24,25,26. Condensate formation is often driven by transient, selective, and multivalent protein-protein, protein-nucleic acid, or nucleic acid-nucleic acid interactions27. Under certain conditions, these interactions can lead to liquid-liquid phase separation (LLPS), a density transition that locally enriches specific biomolecules in membraneless droplets. Such multivalent interactions are often mediated by the intrinsically disordered regions (IDRs) of proteins1,28,29. Biophysical characterization of these interactions at the molecular level is critical to our understanding of numerous healthy and aberrant cellular functions, given the pervasiveness of condensates across them. Although techniques based on confocal fluorescence microscopy, e.g., fluorescence recovery after photobleaching (FRAP)30,31,32, have been widely used to qualitatively show that the molecular exchanges between condensates and the surrounding cellular environment are dynamic, quantifying the interaction dynamics of specific biomolecules within condensates is generally not possible using conventional confocal microscopy or single-molecule microscopy without specialized data analysis methods. The single-particle tracking (SPT) technique described in this protocol is based on live-cell single-molecule microscopy33 and provides a uniquely powerful tool to quantify the interaction dynamics between specific proteins within condensates. The readout of SPT for such measurement is the mean residence time of a protein of interest in the condensates.
The protocol can be broken down into two parts - data acquisition and data analysis. The first step of imaging data acquisition is to express in cells a protein of interest that is fused to a HaloTag34. This enables labeling of the protein of interest with two fluorophores, where a majority of the protein molecules are to be labeled with a non-photoactivatable fluorophore (e.g., JFX549 Halo ligand35) and a small fraction of them are to be labeled with a spectrally distinct, photoactivatable fluorophore (e.g., PA-JF646 Halo ligand36). This allows for the simultaneous acquisition of all condensate locations in the cell and the acquisition of single-molecule movies of the protein of interest binding and unbinding to the condensates. Meanwhile, the same type of cells are modified to stably express Halo-tagged H2B, a histone that is largely immobile on chromatin. The cells are then stained with the PA-JF646 Halo ligand to enable single-molecule imaging of H2B. As will be discussed in detail below, this experiment accounts for the contribution of photobleaching to enable precise quantification of the interaction dynamics of the protein of interest. Cells for imaging experiments must then be cultured on clean coverslips, stained with HaloTag ligand(s), and assembled into a live-cell imaging chamber. From there, the sample is imaged under highly inclined and laminated optical sheet (HILO) illumination on a total internal reflection fluorescence (TIRF) microscope capable of two-channel imaging and single-molecule detection. The emission is then split onto two cameras, one tracking condensate positions and one tracking single molecules. Acquisition is performed with a long integration time (on the order of hundreds of ms) to blur out freely-diffusing proteins and only capture proteins that are less mobile due to binding to stable structures in the cell37.
The first step of data analysis is using an established single-particle tracking (SPT) algorithm38,39 to localize individual protein molecules in each frame of the movie and assemble the localizations into a trajectory for each molecule over its detectable lifetime. The trajectories are then sorted into those representing molecules inside and those representing molecules outside the condensates by comparing the localizations of the molecules throughout their trajectories to the localizations of all the condensates at the corresponding times1.
Next, a survival curve (1 - CDF) is generated using the lengths of all the in-condensate trajectories. The apparent mean residence time of the molecules is then extracted by fitting the survival curve to the following two-component exponential model of protein binding,
<img align="center" alt="Equation 1" src="/files/ftp_upload/66169/66169eq01.jpg" style="margin: auto;" />,
with A as the fraction of molecules non-specifically bound and with kobs,ns and kobs,s as the observed dissociation rates of the non-specifically bound and specifically bound molecules, respectively. Only kobs,s is considered from here onward. The dynamics of both protein dissociation, ktrue,s, and photobleaching of the fluorophore, kpb, contribute to kobs,s as
<img alt="Equation 2" src="/files/ftp_upload/66169/66169eq02.jpg" style="margin: auto;" />;
thus, to isolate the effects of protein dissociation, the specific dissociation rate of H2B-Halo in the cell line mentioned prior is measured.
<img alt="Equation 3" src="/files/ftp_upload/66169/66169eq03.jpg" style="margin: auto;" />
H2B is a protein that is stably integrated into chromatin and that experiences minimal dissociation in the time scale of a single-molecule movie acquisition37. Its specific dissociation rate is then equal to the photobleaching rate of the PA-JF646 Halo ligand, or
<img alt="Equation 4" src="/files/ftp_upload/66169/66169eq04.jpg" style="margin: auto;" />.
The mean in-condensate residence time of the protein of interest, <img alt="Equation 5" src="/files/ftp_upload/66169/66169eq05.jpg" style="margin: auto;" />, is then
<img alt="Equation 6" src="/files/ftp_upload/66169/66169eq06.jpg" style="margin: auto;" />.
Representative results from Irgen-Gioro et al.40 are shown, where this protocol was applied to demonstrate that fusing an oligomerization domain to IDR results in longer residence times of the IDR in its condensates. This result suggests that the added oligomerization domain stabilizes the homotypic interactions of the IDR that drives LLPS. In principle, the same method with slightly modified protocols can be applied to characterize the homotypic or heterotypic interactions of any protein that participates in the formation of any types of condensates.

<table><tbody><tr><td>0.1 &amp;micro;m TetraSpeck microsphere</td><td>Invitrogen</td><td>T7279</td><td>Single-molecule imaging</td></tr><tr><td>25 mm Diameter, #1.5 Coverslips</td><td>Marienfeld Superior</td><td>111650</td><td>Preparation of coverslips</td></tr><tr><td>593/40 nm bandpass filter</td><td>Semrock</td><td>FF01-593/40-25</td><td>Single-molecule imaging</td></tr><tr><td>676/37 nm bandpass filter</td><td>Semrock</td><td>FF01-676/37-25</td><td>Single-molecule imaging</td></tr><tr><td>6-Well TC Plate</td><td>Genesee</td><td>25-105MP</td><td>Preparation of cells for microscopy</td></tr><tr><td>Cell Line: U-2 OS</td><td>ATCC</td><td>HTB-96</td><td>Labeling of proteins in cells</td></tr><tr><td>ConvertASCII_SlowTracking_css3&lt;br/&gt; .m</td><td></td><td></td><td>Analysis of single-molecule imaging data: Available in Chong et al., 2018</td></tr><tr><td>Coverglass Staining Rack</td><td>Thomas</td><td>24957</td><td>Preparation of coverslips</td></tr><tr><td>Deuterated Janelia Fluor 549 (JFX549)</td><td>Janelia Research Campus</td><td></td><td>Preparation of cells for microscopy</td></tr><tr><td>DMEM, Low Glucose</td><td>Gibco</td><td>10-567-022</td><td>Labeling of proteins in cells: Growth media used: DMEM with 5% fetal bovine serum, 1% penstrep</td></tr><tr><td>Eclipse Ti2-E Inverted Microscope</td><td>Nikon</td><td></td><td>Single-molecule imaging</td></tr><tr><td>Ethanol 200 Proof</td><td>Lab Alley</td><td>EAP200-1GAL</td><td>Preparation of coverslips</td></tr><tr><td>evalSPT</td><td></td><td></td><td>Analysis of single-molecule imaging data: Available in Drosopoulos et al., 2020</td></tr><tr><td>Fetal Bovine Serum</td><td>Cytiva</td><td>SH30396.03</td><td>Labeling of proteins in cells: Growth media used: DMEM with 5% fetal bovine serum, 1% penstrep</td></tr><tr><td>Fiji</td><td></td><td></td><td>Analysis of single-molecule imaging data</td></tr><tr><td>Ikon Ultra CCD Camera</td><td>Andor</td><td>X-13723</td><td>Single-molecule imaging</td></tr><tr><td>Longpass dichroic beamsplitter</td><td>Semrock</td><td>Di02-R635-25x36</td><td>Single-molecule imaging: Red/Far Red beamsplitter</td></tr><tr><td>LUN-F Laser Unit</td><td>Nikon</td><td></td><td>Single-molecule imaging: 405/488/561/640</td></tr><tr><td>MatTek glass-bottom dish</td><td>MatTek</td><td>P35G-1.5-20-C</td><td>Preparation of cells for microscopy: 35 mm, #1.5 coverslip dish for cell culture.</td></tr><tr><td>NIS-Elements</td><td>Nikon</td><td></td><td>Single-molecule imaging: Microscope acquisition software</td></tr><tr><td>nucleus and cluster mask_v2.txt</td><td></td><td></td><td>Analysis of single-molecule imaging data: Available in Chong et al., 2018</td></tr><tr><td>Penicillin-Streptomycin</td><td>Gibco</td><td>15-140-122</td><td>Labeling of proteins in cells: Growth media used: DMEM with 5% fetal bovine serum, 1% penstrep</td></tr><tr><td>Phosphate Buffered Saline</td><td>Thermo Fisher Scientific</td><td>18912014</td><td>Labeling of proteins in cells</td></tr><tr><td>Photoactivatable Janelia Fluor 646 (PA-JF646)</td><td>Janelia Research Campus</td><td></td><td>Preparation of cells for microscopy</td></tr><tr><td>PLOT_ResidenceHist_css.m</td><td></td><td></td><td>Analysis of single-molecule imaging data: Available in Chong et al., 2018</td></tr><tr><td>Potassium Hydroxide</td><td>Mallinckrodt Chemicals</td><td>6984-06</td><td>Preparation of coverslips</td></tr><tr><td>pretracking_comb.txt</td><td></td><td></td><td>Analysis of single-molecule imaging data: Available in Chong et al., 2018</td></tr><tr><td>SLIMfast</td><td></td><td></td><td>Analysis of single-molecule imaging data: Available in Teves et al., 2016</td></tr><tr><td>Stage-top incubation system</td><td>Tokai Hit</td><td></td><td>Single-molecule imaging: For live-cell imaging</td></tr><tr><td>TwinCam dual emission image splitter</td><td>Cairn Research</td><td></td><td>Single-molecule imaging</td></tr><tr><td>Ultrasonic Cleaner</td><td>Branson</td><td>5800</td><td>Preparation of coverslips</td></tr></tbody></table>

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.

Here, we present representative results from Irgen-Gioro et al.40, where we used this SPT protocol to compare the interaction dynamics of two proteins in their respective self-assembled LLPS condensates. TAF15 (TATA-box binding protein associated factor 15) contains an IDR that can undergo LLPS upon overexpression in human cells. We hypothesized that fusing TAF15(IDR) to FTH1 (ferritin heavy chain 1), which forms a 24-subunit oligomer, would lead to more stable homotypic protein-protein interactio...

Watch this Scientific Journal Video about Author Spotlight: Evaluation of Protein-Condensate Dynamics in Live Human Cells at JoVE.com

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.

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 ...

single-molecule-measurement-protein-interaction-dynamics-within

Research

JoVE Journal

Biology

3.3K Views.  California Institute of Technology. 1 Ons lab heeft tot doel het interactiegedrag2 van intrinsiek ongeordende eiwitregio's 3 te begrijpen en hoe ze een rol spelen bij transcriptionele regulatie 4 in gezonde en zieke menselijke cellen. 5 In ons lab ontwikkelen en gebruiken we nieuwe 6 single molecule microscopie technieken 7 in combinatie met moleculaire biologie, 8 biochemische en proteomische benaderingen 9 om biomoleculaire condensaten te bestuderen. 10 Dit protocol maakt het mogelijk de d...

Video: Auteur in de schijnwerpers: evaluatie van de dynamiek van eiwitcondensaat in levende menselijke cellen

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 ...

Biochemistry

Calibration-free In Vitro Quantification of Protein Homo-oligomerization Using Commercial Instrumentation and Free, Open Source Brightness Analysis Software

Number and brightness is a calibration-free fluorescence fluctuation spectroscopy (FFS) technique for detecting protein homo-oligomerization. It can be employed using a conventional confocal microscope equipped with digital detectors. A protocol for the use of the technique in vitro is shown by means of a use case where number and brightness can be seen to accurately quantify the oligomeric state of mVenus-labelled FKBP12F36V before and after the addition of the dimerizing drug AP20187. The importance of using the correct microscope acquisition parameters and the correct data preprocessing and analysis methods are discussed. In particular, the importance of the choice of photobleaching correction is stressed. This inexpensive method can be employed to study protein-protein interactions in many biological contexts.

Calibration-free In Vitro Quantification of Protein Homo-oligomerization Using Commercial Instrumentation and Free, Open Source Brightness Analysis Software

This protocol describes a calibration-free approach for quantifying protein homo-oligomerization in vitro based on fluorescence fluctuation spectroscopy using commercial light scanning microscopy. The correct acquisition settings and analysis methods are shown.

Number and brightness is a calibration-free fluorescence fluctuation spectroscopy (FFS) technique for detecting protein homo-oligomerization. It can be ...

Measuring Interactions of Globular and Filamentous Proteins by Nuclear Magnetic Resonance Spectroscopy (NMR) and Microscale Thermophoresis (MST)

Filamentous proteins such as vimentin provide organization within cells by providing a structural scaffold with sites that bind proteins containing plakin repeats. Here, a protocol for detecting and measuring such interactions is described using the globular plakin repeat domain of envoplakin and the helical coil of vimentin. This provides a basis for determining whether a protein binds vimentin (or similar filamentous proteins) and for measurement of the affinity of the interaction. The globular protein of interest is labeled with 15N and titrated with vimentin protein in solution. A two-dimensional NMR spectrum is acquired to detect interactions by observing changes in peak shape or chemical shifts, and to elucidate effects of solution conditions including salt levels, which influence vimentin quaternary structure. If the protein of interest binds the filamentous ligand, the binding interaction is quantified by MST using the purified proteins. The approach is a straightforward way for determining whether a protein of interest binds a filament, and for assessing how alterations, such as mutations or solution conditions, affect the interaction.

Measuring Interactions of Globular and Filamentous Proteins by Nuclear Magnetic Resonance Spectroscopy NMR and Microscale Thermophoresis MST

Here, we present a protocol for the production and purification of proteins that are labeled with stable isotopes, and subsequent characterization of protein-protein interactions using Nuclear Magnetic Resonance (NMR) spectroscopy and MicroScale Thermophoresis (MST) experiments.

Filamentous proteins such as vimentin provide organization within cells by providing a structural scaffold with sites that bind proteins containing plakin ...

Using In Vitro Fluorescence Resonance Energy Transfer to Study the Dynamics Of Protein Complexes at a Millisecond Time Scale

Proteins are the primary operators of biological systems, and they usually interact with other macro- or small molecules to carry out their biological functions. Such interactions can be highly dynamic, meaning the interacting subunits are constantly associated and dissociated at certain rates. While measuring the binding affinity using techniques such as quantitative pull-down reveals the strength of the interaction, studying the binding kinetics provides insights on how fast the interaction occurs and how long each complex can exist. Furthermore, measuring the kinetics of an interaction in the presence of an additional factor, such as a protein exchange factor or a drug, helps reveal the mechanism by which the interaction is regulated by the other factor, providing important knowledge for the advancement of biological and medical research. Here, we describe a protocol for measuring the binding kinetics of a protein complex that has a high intrinsic association rate and can be dissociated quickly by another protein. The method uses fluorescence resonance energy transfer to report the formation of the protein complex in vitro, and it enables monitoring the fast association and dissociation of the complex in real time on a stopped-flow fluorimeter. Using this assay, the association and dissociation rate constants of the protein complex are quantified.

Protein-protein interactions are critical for biological systems, and studies of the binding kinetics provide insights into the dynamics and function of protein complexes. We describe a method that quantifies the kinetic parameters of a protein complex using fluorescence resonance energy transfer and the stopped-flow technique. &#160;

Proteins are the primary operators of biological systems, and they usually interact with other macro- or small molecules to carry out their biological ...

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 ...

OaAEP1-Mediated Enzymatic Synthesis and Immobilization of Polymerized Protein for Single-Molecule Force Spectroscopy

Chemical and bio-conjugation techniques have been developed rapidly in recent years and allow the building of protein polymers. However, a controlled protein polymerization process is always a challenge. Here, we have developed an enzymatic methodology for constructing polymerized protein step by step in a rationally-controlled sequence. In this method, the C-terminus of a protein monomer is NGL for protein conjugation using OaAEP1 (Oldenlandia affinis asparaginyl endopeptidases) 1) while the N-terminus was a cleavable TEV (tobacco etch virus) cleavage site plus an L (ENLYFQ/GL) for temporary N-terminal protecting. Consequently, OaAEP1 was able to add only one protein monomer at a time, and then the TEV protease cleaved the N-terminus between Q and G to expose the NH2-Gly-Leu. Then the unit is ready for next OaAEP1 ligation. The engineered polyprotein is examined by unfolding individual protein domain using atomic force microscopy-based single-molecule force spectroscopy (AFM-SMFS). Therefore, this study provides a useful strategy for polyprotein engineering and immobilization.

Here, we present a protocol to conjugate protein monomer by enzymes forming protein polymer with a controlled sequence and immobilize it on the surface for single-molecule force spectroscopy studies.

Chemical and bio-conjugation techniques have been developed rapidly in recent years and allow the building of protein polymers. However, a controlled ...

Utilizing Time-Resolved Protein-Induced Fluorescence Enhancement to Identify Stable Local Conformations One &#945;-Synuclein Monomer at a Time

Using spectroscopic rulers to track multiple conformations of single biomolecules and their dynamics have revolutionized the understanding of structural dynamics and its contributions to biology. While the FRET-based ruler reports on inter-dye distances in the 3-10 nm range, other spectroscopic techniques, such as protein-induced fluorescence enhancement (PIFE), report on the proximity between a dye and a protein surface in the shorter 0-3 nm range. Regardless of the method of choice, its use in measuring freely-diffusing biomolecules one at a time retrieves histograms of the experimental parameter yielding separate centrally-distributed sub-populations of biomolecules, where each sub-population represents either a single conformation that stayed unchanged within milliseconds, or multiple conformations that interconvert much faster than milliseconds, and hence an averaged-out sub-population. In single-molecule FRET, where the reported parameter in histograms is the inter-dye FRET efficiency, an intrinsically disordered protein, such as the &#945;-Synuclein monomer in buffer, was previously reported as exhibiting a single averaged-out sub-population of multiple conformations interconverting rapidly. While these past findings depend on the 3-10 nm range of the FRET-based ruler, we sought to put this protein to the test using single-molecule PIFE, where we track the fluorescence lifetime of site-specific sCy3-labeled &#945;-Synuclein proteins one at a time. Interestingly, using this shorter range spectroscopic proximity sensor, sCy3-labeled &#945;-Synuclein exhibits several lifetime sub-populations with distinctly different mean lifetimes that interconvert in 10-100 ms. These results show that while &#945;-Synuclein might be disordered globally, it nonetheless attains stable local structures. In summary, in this work we highlight the advantage of using different spectroscopic proximity sensors that track local or global structural changes one biomolecule at a time.

Time-resolved single-molecule protein-induced fluorescence enhancement is a useful fluorescence spectroscopic proximity sensor sensitive to local structural changes in proteins. Here we show it can be used to uncover stable local conformations in &#945;-Synuclein, which is otherwise known as globularly unstructured and unstable when measured using the longer range FRET ruler.

Using spectroscopic rulers to track multiple conformations of single biomolecules and their dynamics have revolutionized the understanding of structural ...

Expanding the Mass Photometry Analysis Concentration Range with Microfluidics

Analysis of Protein Complex Formation at Micromolar Concentrations by Coupling Microfluidics with Mass Photometry

Mass photometry is a versatile mass measurement technology that enables the study of biomolecular interactions and complex formation in solution without labels. Mass photometry is generally suited to analyzing samples in the 100 pM-100 nM concentration range. However, in many biological systems, it is necessary to measure more concentrated samples to study low-affinity or transient interactions. Here, we demonstrate a method that effectively expands the range of sample concentrations that can be analyzed by mass photometry from nanomolar to tens of micromolar. 
In this protocol, mass photometry is combined with a novel microfluidics system to investigate the formation of protein complexes in solution in the micromolar concentration range. With the microfluidics system, users can maintain a sample at a desired higher concentration followed by dilution to the nanomolar range - several milliseconds prior to the mass photometry measurement. Due to the speed of the dilution, data is obtained before the equilibrium of the sample has shifted (i.e., dissociation of the complex). 
The technique is applied to measure interactions between an immunoglobulin G (IgG) antibody and the neonatal Fc receptor, showing the formation of high-order complexes that were not quantifiable with static mass photometry measurements. 
In conclusion, the combination of mass photometry and microfluidics makes it possible to characterize samples in the micromolar concentration range and is proficient in measuring biomolecular interactions with weaker affinities. These capabilities can be applied in a range of contexts - including the development and design of biotherapeutics - enabling thorough characterization of diverse protein-protein interactions.

Author Spotlight: Characterization of Low-Affinity Protein Interactions in Solution Using MassFluidix Technology

This protocol combines mass photometry with a novel microfluidics system to investigate low-affinity protein-protein interactions. This approach is based on the rapid dilution of highly concentrated complexes in solution, which enables low-affinity measurements and broadens the applicability of mass photometry.

Mass photometry is a versatile mass measurement technology that enables the study of biomolecular interactions and complex formation in solution without ...

Measurement of Force-Sensitive Protein Dynamics in Living Cells Using a Combination of Fluorescent Techniques

Cells sense and respond to physical cues in their environment by converting mechanical stimuli into biochemically-detectable signals in a process called mechanotransduction. A crucial step in mechanotransduction is the transmission of forces between the external and internal environments. To transmit forces, there must be a sustained, unbroken physical linkage created by a series of protein-protein interactions. For a given protein-protein interaction, mechanical load can either have no effect on the interaction, lead to faster disassociation of the interaction, or even stabilize the interaction. Understanding how molecular load dictates protein turnover in living cells can provide valuable information about the mechanical state of a protein, in turn elucidating its role in mechanotransduction. Existing techniques for measuring force-sensitive protein dynamics either lack direct measurements of protein load or rely on the measurements performed outside of the cellular context. Here, we describe a protocol for the F&#246;rster resonance energy transfer-fluorescence recovery after photobleaching (FRET-FRAP) technique, which enables the measurement of force-sensitive protein dynamics within living cells. This technique is potentially applicable to any FRET-based tension sensor, facilitating the study of force-sensitive protein dynamics in variety of subcellular structures and in different cell types.

Here, we present a protocol for the simultaneous use of F&#246;rster resonance energy transfer-based tension sensors to measure protein load and fluorescence recovery after photobleaching to measure protein dynamics enabling the measurement of force-sensitive protein dynamics within living cells.

Cells sense and respond to physical cues in their environment by converting mechanical stimuli into biochemically-detectable signals in a process called ...

Bioengineering

Combining Single-molecule Manipulation and Imaging for the Study of Protein-DNA Interactions

The paper describes the combination of optical tweezers and single molecule fluorescence detection for the study of protein-DNA interaction. The method offers the opportunity of investigating interactions occurring in solution (thus avoiding problems due to closeby surfaces as in other single molecule methods), controlling the DNA extension and tracking interaction dynamics as a function of both mechanical parameters and DNA sequence. The methods for establishing successful optical trapping and nanometer localization of single molecules are illustrated. We illustrate the experimental conditions allowing the study of interaction of lactose repressor (lacI), labeled with Atto532, with a DNA molecule containing specific target sequences (operators) for LacI binding. The method allows the observation of specific interactions at the operators, as well as one-dimensional diffusion of the protein during the process of target search. The method is broadly applicable to the study of protein-DNA interactions but also to molecular motors, where control of the tension applied to the partner track polymer (for example actin or microtubules) is desirable.

Here we describe the instrumentation and methods for detecting single fluorescently-labeled protein molecules interacting with a single DNA molecule suspended between two optically trapped microspheres.

The paper describes the combination of optical tweezers and single molecule fluorescence detection for the study of protein-DNA interaction. The method ...

Single-Molecule Measurement of Protein Interaction Dynamics Within Biomolecular Condensates

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