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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.1 &#181;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 
			.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 ...

1 Our lab aims to understand the interaction behaviors<v>2 of intrinsically disordered protein regions 3 and how they play roles in transcriptional regulation 4 in healthy and diseased human cells. 5 In our lab, we develop and use novel 6 single molecule microscopy techniques 7 in combination with molecular biology, 8 biochemical and proteomic approaches 9 to study biomolecular condensates. 10 This protocol enables the quantification of the dynamics 11 by which a specific protein 12 binds to a particular type of condensate 13 in live human cells, 14 and is broadly applicable 15 to measuring the interaction dynamics of any protein 16 that participates in liquid-liquid phase separation.

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Research

JoVE Journal

Biology

2.7K Views.  California Institute of Technology. 1 Our lab aims to understand the interaction behaviors2 of intrinsically disordered protein regions 3 and how they play roles in transcriptional regulation 4 in healthy and diseased human cells. 5 In our lab, we develop and use novel 6 single molecule microscopy techniques 7 in combination with molecular biology, 8 biochemical and proteomic approaches 9 to study biomolecular condensates. 10 This protocol enables the quantification of the dynamics 11 by which a specific protein 12 binds to ...

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

Visualizing Single-molecule DNA Replication with Fluorescence Microscopy

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA template is attached to a functionalized glass coverslip and replicated extensively after introduction of replication proteins and nucleotides (Figure 1). The growing product double-strand DNA (dsDNA) is extended with laminar flow and visualized by using an intercalating dye. Measuring the position of the growing DNA end in real time allows precise determination of replication rate (Figure 2). Furthermore, the length of completed DNA products reports on the processivity of replication. This experiment can be performed very easily and rapidly and requires only a fluorescence microscope with a reasonably sensitive camera.


This protocol demonstrates a simple single-molecule fluorescence microscopy technique for visualizing DNA replication by individual replisomes in real time.

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA ...

Automated System for Single Molecule Fluorescence Measurements of Surface-immobilized Biomolecules

Fluorescence Resonance Energy Transfer (FRET) microscopy has been widely used to study the structure and dynamics of molecules of biological interest, such as nucleic acids and proteins. Single molecule FRET (sm-FRET) measurements on immobilized molecules permit long observations of the system -effectively until both dyes photobleach- resulting in time-traces that report on biomolecular dynamics with a broad range of timescales from milliseconds to minutes. To facilitate the acquisition of large number of traces for statistical analyses, the process must be automated and the sample environment should be tightly controlled over the entire measurement time (~12 hours). This is accomplished using an automated scanning confocal microscope that allows the interrogation of thousands of single molecules overnight, and a microfluidic cell that permits the controlled exchange of buffer, with restricted oxygen content and maintains a constant temperature throughout the entire measuring period. Here we show how to assemble the microfluidic device and how to activate its surface for DNA immobilization. Then we explain how to prepare a buffer to maximize the photostability and lifetime of the fluorophores. Finally, we show the steps involved in preparing the setup for the automated acquisition of time-resolved single molecule FRET traces of DNA molecules.

In this article we describe how we obtain FRET traces from individual DNA molecules immobilized to a surface using an automated scanning confocal microscope.

Fluorescence Resonance Energy Transfer (FRET) microscopy has been widely used to study the structure and dynamics of molecules of biological interest, ...

Single-Molecule Imaging of Nuclear Transport

The utility of single molecule fluorescence microscopy approaches has been proven to be of a great avail in understanding biological reactions over the last decade. The investigation of molecular interactions with high temporal and spatial resolutions deep within cells has remained challenging due to the inherently weak signals arising from individual molecules. Recent works by Yang et al. demonstrated that narrow-field epifluorescence microscopy allows visualization of nucleocytoplasmic transport at the single molecule level. By the single molecule approach, important kinetics, such as nuclear transport time and efficiency, for signal-dependent and independent cargo molecules have been obtained. Here we described a protocol for the methodological approach with an improved spatiotemporal resolution of 0.4 ms and 12 nm. The improved resolution enabled us to capture transient active transport and passive diffusion events through the nuclear pore complexes (NPC) in semi-intact cells. We expect this method to be used in elucidating other binding and trafficking events within cells.

Single molecule microscopy approch provided novel insights into nuclear transport.

The utility of single molecule fluorescence microscopy approaches has been proven to be of a great avail in understanding biological reactions over the ...

Identifying Protein-protein Interaction in Drosophila Adult Heads by Tandem Affinity Purification (TAP)

Genetic screens conducted using Drosophila melanogaster (fruit fly) have made numerous milestone discoveries in the advance of biological sciences. However, the use of biochemical screens aimed at extending the knowledge gained from genetic analysis was explored only recently. Here we describe a method to purify the protein complex that associates with any protein of interest from adult fly heads. This method takes advantage of the Drosophila GAL4/UAS system to express a bait protein fused with a Tandem Affinity Purification (TAP) tag in fly neurons in vivo, and then implements two rounds of purification using a TAP procedure similar to the one originally established in yeast1 to purify the interacting protein complex. At the end of this procedure, a mixture of multiple protein complexes is obtained whose molecular identities can be determined by mass spectrometry. Validation of the candidate proteins will benefit from the resource and ease of performing loss-of-function studies in flies. Similar approaches can be applied to other fly tissues. We believe that the combination of genetic manipulations and this proteomic approach in the fly model system holds tremendous potential for tackling fundamental problems in the field of neurobiology and beyond.

Identifying Protein-protein Interaction in Drosophila Adult Heads by Tandem Affinity Purification TAP

Drosophila is famous for its powerful genetic manipulation, but not for its suitability of in-depth biochemical analysis. Here we present a TAP-based procedure to identify interacting partners of any protein of interest from the fly brain. This procedure can potentially lead to new avenues of research.

Genetic screens conducted using Drosophila melanogaster (fruit fly) have made numerous milestone discoveries in the advance of biological sciences. ...

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 ...

Studying DNA Looping by Single-Molecule FRET

Bending of double-stranded DNA (dsDNA) is associated with many important biological processes such as DNA-protein recognition and DNA packaging into nucleosomes. Thermodynamics of dsDNA bending has been studied by a method called cyclization which relies on DNA ligase to covalently join short sticky ends of a dsDNA. However, ligation efficiency can be affected by many factors that are not related to dsDNA looping such as the DNA structure surrounding the joined sticky ends, and ligase can also affect the apparent looping rate through mechanisms such as nonspecific binding. Here, we show how to measure dsDNA looping kinetics without ligase by detecting transient DNA loop formation by FRET (Fluorescence Resonance Energy Transfer). dsDNA molecules are constructed using a simple PCR-based protocol with a FRET pair and a biotin linker. The looping probability density known as the J factor is extracted from the looping rate and the annealing rate between two disconnected sticky ends. By testing two dsDNAs with different intrinsic curvatures, we show that the J factor is sensitive to the intrinsic shape of the dsDNA.

This study presents a detailed experimental procedure to measure looping dynamics of double-stranded DNA using single-molecule Fluorescence Resonance Energy Transfer (FRET). The protocol also describes how to extract the looping probability density called the J factor.

Bending of double-stranded DNA (dsDNA) is associated with many important biological processes such as DNA-protein recognition and DNA packaging into ...

Identifying Protein-protein Interaction Sites Using Peptide Arrays

Protein-protein interactions mediate most of the processes in the living cell and control homeostasis of the organism. Impaired protein interactions may result in disease, making protein interactions important drug targets. It is thus highly important to understand these interactions at the molecular level. Protein interactions are studied using a variety of techniques ranging from cellular and biochemical assays to quantitative biophysical assays, and these may be performed either with full-length proteins, with protein domains or with peptides. Peptides serve as excellent tools to study protein interactions since peptides can be easily synthesized and allow the focusing on specific interaction sites. Peptide arrays enable the identification of the interaction sites between two proteins as well as screening for peptides that bind the target protein for therapeutic purposes. They also allow high throughput SAR studies. For identification of binding sites, a typical peptide array usually contains partly overlapping 10-20 residues peptides derived from the full sequences of one or more partner proteins of the desired target protein. Screening the array for binding the target protein reveals the binding peptides, corresponding to the binding sites in the partner proteins, in an easy and fast method using only small amount of protein.
In this article we describe a protocol for screening peptide arrays for mapping the interaction sites between a target protein and its partners. The peptide array is designed based on the sequences of the partner proteins taking into account their secondary structures. The arrays used in this protocol were Celluspots arrays prepared by INTAVIS Bioanalytical Instruments. The array is blocked to prevent unspecific binding and then incubated with the studied protein. Detection using an antibody reveals the binding peptides corresponding to the specific interaction sites between the proteins.

Peptide array screening is a high throughput assay for identifying protein-protein interaction sites. This allows mapping multiple interactions of a target protein and can serve as a method for identifying sites for inhibitors that target a protein. Here we describe a protocol for screening and analyzing peptide arrays.

Protein-protein interactions mediate most of the processes in the living cell and control homeostasis of the organism. Impaired protein interactions may ...

Luciferase Complementation Imaging Assay in Nicotiana benthamiana Leaves for Transiently Determining Protein-protein Interaction Dynamics

Protein-protein interactions are fundamental mechanisms for relaying signal transduction in most cellular processes; therefore, identification of novel protein-protein interaction pairs and monitoring protein interaction dynamics are of particular interest for revealing how plants respond to environmental factors and/or developmental signals. A plethora of approaches have been developed to examine protein-protein interactions, either in vitro or in vivo. Among them, the recently established luciferase complementation imaging (LCI) assay is the simplest and fastest method for demonstrating in vivo protein-protein interactions. In this assay, protein A or protein B is fused with the amino-terminal or carboxyl-terminal half of luciferase, respectively. When protein A interacts with protein B, the two halves of luciferase will be reconstituted to form a functional and active luciferase enzyme. Luciferase activity can be recorded with a luminometer or CCD-camera. Compared with other approaches, the LCI assay shows protein-protein interactions both qualitatively and quantitatively. Agrobacterium infiltration in Nicotiana benthamiana leaves is a widely used system for transient protein expression. With the combination of LCI and transient expression, these approaches show that the physical interaction between COP1 and SPA1 was gradually reduced after jasmonate treatment.

Luciferase Complementation Imaging Assay in Nicotiana benthamiana Leaves for Transiently Determining Protein-protein Interaction Dynamics

This manuscript describes an easy and rapid experimental procedure for determining protein-protein interactions based on the measurement of luciferase activity.

Protein-protein interactions are fundamental mechanisms for relaying signal transduction in most cellular processes; therefore, identification of novel ...

Visualizing the Interaction Between the Qdot-labeled Protein and Site-specifically Modified &#955; DNA at the Single Molecule Level

The fluorescence microscopy&#160;has made great contributions in dissecting the mechanisms of complex biological processes at the single molecule level. In single molecule assays for studying DNA-protein interactions, there are two important factors for consideration: the DNA substrate with enough length for easy observation and labeling a protein with a suitable fluorescent probe. 48.5 kb &#955; DNA is a good candidate for the DNA substrate. Quantum dots (Qdots), as a class of fluorescent probes, allow long-time observation (minutes to hours) and high-quality image acquisition. In this paper, we present a protocol to study DNA-protein interactions at the single-molecule level, which includes preparing a site-specifically modified &#955; DNA and labeling a target protein with streptavidin-coated Qdots. For a proof of concept, we choose ORC (origin recognition complex) in budding yeast as a protein of interest and visualize its interaction with an ARS (autonomously replicating sequence) using TIRFM. Compared with other fluorescent probes, Qdots have obvious advantages in single molecule studies due to its high stability against photobleaching, but it should be noted that this property limits its application in quantitative assays.

Visualizing the Interaction Between the Qdot-labeled Protein and Site-specifically Modified &amp;#955; DNA at the Single Molecule Level

Here, we present a protocol to study DNA-protein interactions by total internal reflection fluorescence microscopy (TIRFM) using a site-specifically modified &#955; DNA substrate and a Quantum-dot labeled protein.

The fluorescence microscopy&#160;has made great contributions in dissecting the mechanisms of complex biological processes at the single molecule level. ...

Chemical Dimerization-Induced Protein Condensates on Telomeres

Chromatin-associated condensates are implicated in many nuclear processes, but the underlying mechanisms remain elusive. This protocol describes a chemically-induced protein dimerization system to create condensates on telomeres. The chemical dimerizer consists of two linked ligands that can each bind to a protein: Halo ligand to Halo-enzyme and trimethoprim (TMP) to E. coli dihydrofolate reductase (eDHFR), respectively. Fusion of Halo enzyme to a telomere protein anchors dimerizers to telomeres through covalent Halo ligand-enzyme binding. Binding of TMP to eDHFR recruits eDHFR-fused phase separating proteins to telomeres and induces condensate formation. Because TMP-eDHFR interaction is non-covalent, condensation can be reversed by using excess free TMP to compete with the dimerizer for eDHFR binding. An example of inducing promyelocytic leukemia (PML) nuclear body formation on telomeres and determining condensate growth, dissolution, localization and composition is shown. This method can be easily adapted to induce condensates at other genomic locations by fusing Halo to a protein that directly binds to the local chromatin or to dCas9 that is targeted to the genomic locus with a guide RNA. By offering the temporal resolution required for single cell live imaging while maintaining phase separation in a population of cells for biochemical assays, this method is suitable for probing both the formation and function of chromatin-associated condensates.

This protocol illustrates a chemically induced protein dimerization system to create condensates on chromatin.&#160; The formation of promyelocytic leukemia (PML) nuclear body on telomeres with chemical dimerizers is demonstrated. Droplet growth, dissolution, localization and composition are monitored with live cell imaging, immunofluorescence (IF) and fluorescence in situ hybridization (FISH).