Name: Time-Resolved Fluorescence Anisotropy from Single Molecules for Characterizing Local Flexibility in Biomolecules (Video) | JoVE Journal
Uploaded: 2025-04-25T13:40:00.000+00:00
Duration: 10 min 23 s
Description: Learn time-resolved fluorescence anisotropy for studying biomolecular flexibility using single-molecule confocal microscopy. This method reveals protein dynamics and conformational changes in FoxP1.

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5 minutes

This work was supported by FONDECYT grants 11200729 and FONDEQUIP EQM200202 to E.M., NIH R15CA280699 R01GM151334, and NSF CAREER MCB 1749778 awards to HS. NK acknowledged support from the Clemson University Postdoctoral fellowship program.

NOTE: Selecting the proper fluorophore is essential for smFA experiments. Biomolecules can be labeled at site-specific positions either by modifying amino acids in proteins or nucleotide bases in nucleic acids with fluorescent markers, depending on the available reactive groups. Among organic dyes52, the Alexa Fluor, Cy, BODIPY, and Janelia Farms families are the most popular choices for smFA, thanks to their long fluorescence lifetimes, photostability, and high quantum yields. BODIPY-FL is often favored for its extended fluorescence lifetime, superior quantum yield, and short connecting linker. Additionally, alternative fluorophores are commonly used in drug screening where bulk techniques are preferred53. Chimeric fluorescent proteins can also be used for live-cell anisotropy experiments and imaging, although there is a limitation of a lower dynamic range.
1. Buffer preparation
NOTE: Wear gloves, eye protection goggles, and a laboratory coat while doing laboratory experiments.
<ol>
	<li>Standard buffer (20 mM 4- (2-hydroxyethyl)-1-piperazineethanesulfonic acid [HEPES], pH 7.8, 150 mM NaCl): Dissolve 2.38 g of HEPES, 4.38 g of NaCl in 400 mL of ultra-pure water, adjust pH to 7.8 and make the final volume to 500 mL.</li>
	<li>Lysis buffer (20 mM HEPES, pH 7.8, 150 mM NaCl, 0.1 mM phenylmethylsulfonyl fluoride [PMSF], 10 &#181;g/mL DNAse): Dissolve 2.38 g of HEPES, 4.38 g of NaCl, and 8.71 mg of PMSF and a final concentration of 10 &#181;g/mL DNAse in 400 mL of ultra-pure water, adjust pH to 7.8 and make the final volume to 500 mL.</li>
	<li>Equilibration buffer (50 mM phosphate buffered saline [PBS], 150 mM NaCl, and 10 mM Imidazole; pH 7.4): Dissolve 5.05 g of Na2HPO4, 0.85 g of NaH2PO4, 4.38 g of NaCl and 0.34 g of Imidazole in 400 mL of ultra-pure water, adjust pH to 7.4 and make the final volume to 500 mL.</li>
	<li>Wash buffer (50 mM PBS, 150 mM NaCl, and 30 mM Imidazole; pH 7.4): Dissolve 5.05 g of Na2HPO4, 0.85 g of NaH2PO4, 4.38 g of NaCl and 1.02 g of Imidazole in 400 mL of ultra-pure water, adjust pH to 7.4 and make the final volume to 500 mL.</li>
	<li>Elution buffer (50 mM PBS, 150 mM NaCl, and 250 mM Imidazole; pH 7.4): Dissolve 5.05 g of Na2HPO4, 0.85 g of NaH2PO4, 4.38 g of NaCl and 8.5 g of Imidazole in 400 mL of ultra-pure water, adjust pH to 7.4 and make the final volume to 500 mL.</li>
	<li>PBS buffer (50 mM Sodium Phosphate Buffered Saline, 150 mM NaCl, pH 7.4). Dissolve 5.05 g of Na2HPO4, 0.85 g of NaH2PO4, and 4.38 g of NaCl in 400 mL of ultra-pure water, adjust pH to 7.4, and make the final volume to 500 mL.</li>
	<li>Sterilize all buffers by filtering the solution using 0.22 &#181;m pore size filters and store the buffers at room temperature (RT).</li>
	<li>Filter the standard buffer using a charcoal filter for single-molecule acquisition.</li>
</ol>
2. Fluorescent probes
<ol>
	<li>BODIPY-FL: Dissolve 5 mg vial containing BODIPY-FL by adding a final volume of 1.29 mL of freshly opened dimethyl sulfoxide (DMSO) to get the final concentration of 10 mM BODIPY-FL.</li>
	<li>Rhodamine 110: Dissolve 3.67 mg of Rhodamine 110 by adding a final volume of 1 mL of freshly opened DMSO to get the final concentration of 10 mM Rhodamine 110. 
	NOTE: Fluorescent probes need to be avoided when exposed to light. Always use a light-sensitive tube (amber color) and wrap it with aluminum foil. If DMSO is already opened, it must be kept in the desiccator to increase its shelf life.</li>
	<li>Prepare small-volume aliquots of the prepared fluorescent probe and store it at -20 &#176;C until further use. 
	NOTE: Avoid freeze-thaw cycles to improve its labeling efficiency.</li>
</ol>
3. Calibration measurements
<ol>
	<li>Perform multiparameter fluorescence detection (MFD) experiments on a home-built polarization resolved setup with 4 detection channels and 2 pulsed lasers (blue 485 nm&#160;and red 640 nm)54. However, for anisotropy measurements, use the blue laser and two detector channels.</li>
	<li>Turn on the detector channels (parallel and perpendicular) and the blue laser power.</li>
	<li>Make sure the laser power is set to 60 &#181;W and open the time-correlated single photon counting (TCSPC) software36 control panel.</li>
	<li>Mix 1 &#181;L of 100 nM Rhodamine 110 and 49 &#181;L of distilled water to make a final concentration of 2 nM Rhodamine 110. Add 50 &#181;L of 2 nM Rhodamine 110 to the center of the cover glass for calibration measurements.</li>
	<li>Next, add a drop of immersion liquid (either oil or water, but taking care about sharing the same refractive index) on top of the microscope objective lens to increase the resolving power of the microscope.</li>
	<li>Place the cover glass on top of the objective lens and make sure the water droplet is at the center of the microscope objective.</li>
	<li>Adjust the image plane to be inside the solution and above the glass surface.</li>
	<li>Adjust the knob to find the second bright focal point and turn to one and a half points. Focus the laser at the interface of the glass and liquid.</li>
	<li>Maximize the number of photons detected by adjusting the pinhole position (70 &#181;m) while monitoring the photon count rate.</li>
	<li>Open the software in the time-tagged time-resolved (TTTR, or T3) mode panel and click the Start button. Record the count rate for 120 s and save the Rhodamine 110.ptu file format using the acquisition software. This acquisition time should be enough considering the concentration (2 nM).</li>
	<li>For background measurements, add 50 &#181;L of distilled water to the center of the cover glass and repeat the steps 3.5-3.10. However, record the photon count rate for 300 s and save the water.ptu file.</li>
	<li>For other background measurements, add 50 &#181;L of standard buffer to the center of the cover glass and repeat steps 3.5-3.10. Record the photon count rate for 300 s and save standard buffer.ptu file. Then, analyze the data using burst integration fluorescence lifetime (BIFL) analysis software.</li>
</ol>
4. Calibration and data analysis
<ol>
	<li>Open the BIFL software and click on confirm the setup from the automatic window. Next, click on get parameters from file, then OK.</li>
	<li>For the measurement to analyze, click on data path array and select it.</li>
	<li>Next, load the water measurement water.ptu file for arrays such as Green scatter. Similarly, select the standard buffer.ptu file for Green background. For Green thick, select Rhodamine 110.ptu.</li>
	<li>Under the single molecule selection parameters, click Next, and then Adjust to display a new pop-up window. Click threshold to change the inter-photon arrival time and select single-molecule event time for the mean inter-photon arrival time (dt). Next, click min. # to select the minimum number of photons per single molecule event, then close the pop-up window by clicking Return. Then, click OK.</li>
	<li>Next, click Color fit parameters to adjust the initial fluorescence lifetime for Green, colors that are generated with fluorescence decay parameters. Adjust Prompt and Delay values by modifying From and To values. Then close the pop-up window by clicking Return. Then, click OK.</li>
	<li>Click Save to process the ASCII files and save them in a selected folder. Then, process the data for time-resolved fluorescence anisotropy decay using ChiSurf22, Photon Distribution Analysis55,56, or Burst Variance Analysis57,58. 
	NOTE: Exemplary data and step-by-step descriptions on how to use ChiSurf, PDA, and BVA are available at github.com/Fluorescence-Tools/chisurf, github.com/Fluorescence-Tools/tttrlib,&#160;www.mpc.hhu.de/en/software/mfd-fcs-and-mfis and github.com/SMB-Lab/feda_tools, respectively. The experimental data is available at Zenodo (10.5281/zenodo.13371503).</li>
</ol>
5. FoxP1 protein preparation
<ol>
	<li>Recombinant FoxP1 bacterial overexpression
	<ol>
		<li>Perform transformation into E. coli C41 bacterial cells after quick-change site-directed polymerase chain reaction (PCR) mutagenesis.</li>
		<li>Prepare LB media and autoclave to sterilize it.</li>
		<li>Pre-inoculate a single E. coli C41 colony by adding into 5 mL of LB medium containing 5 &#181;L (100 &#181;g/mL) ampicillin. Let incubate at 37 &#176;C overnight on an incubator shaker.</li>
		<li>Next day, inoculate large-scale bacterial culture by adding overnight pre-inoculum into 500&#160;mL of LB media with pre-added antibiotic with a ratio of 1:500.</li>
		<li>Monitor the growth of the culture by measuring the absorbance of the culture at 600 nm.</li>
		<li>When the optical density reaches between ~0.5-0.7, induce the protein expression by adding a final concentration of 0.5 mM isopropyl-&#946;-d-1-thiogalactopyranoside (IPTG) and keep the culture at 15 &#176;C on an incubator shaker overnight.</li>
		<li>After achieving an optical density (600 nm) in the range of 1.4-1.6, harvest the bacterial cells by centrifugation at 3000 g for 20 min at 4 &#176;C. Discard the supernatant and store the pellet at -20 &#176;C until use.</li>
	</ol>
	</li>
	<li>Recombinant FoxP1 purification
	<ol>
		<li>Lyse E. coli C41 cells by adding lysis buffer using any lysis methods such as sonication, liquid homogenization, French press, etc.</li>
		<li>Centrifuge the lysate at 14000 g for 10 min at 4 &#176;C.</li>
		<li>For His6-tagged proteins, wash the Ni2+-NTA affinity column and equilibrate with NiSO4 using fast protein liquid chromatography (FPLC).</li>
		<li>Load the His6-tagged FoxP1 protein into the equilibrated Ni2+-NTA affinity column.</li>
		<li>Elute the FoxP1 protein from the Ni2+-NTA column using a linear gradient of the elution buffer.</li>
		<li>After the protein is eluted, perform dialysis for buffer exchange. Add protein into an equilibration buffer without Imidazole and dialyze overnight using a magnetic stirrer at 4 &#176;C. 
		NOTE: If the protein is purified using a different method, do the buffer exchange accordingly.</li>
		<li>To remove the His6-tag, add tobacco etch virus (TEV) protease (1:100 ratio of TEV:FoxP1) to the dialysis overnight. 
		NOTE: His6-tag digestion can also be performed by adding on-column TEV protease digestion. This step would improve the yield of the protein of interest.</li>
		<li>Next day, repeat the steps from 5.2.3-5.2.5. However, this time, elute the protein in the wash buffer rather than the elution buffer.</li>
		<li>Concentrate the protein to an appropriate volume by adding PBS buffer and quantify the protein using absorbance at 280 nm, considering the extinction coefficient of the protein.</li>
	</ol>
	</li>
	<li>Recombinant FoxP1 labeling
	<ol>
		<li>Incubate 50-100 &#181;M FoxP1 protein with a 10-fold molar excess of either Dithiothreitol (DTT) or tris (2-carboxyethyl)phosphine (TCEP) with a final volume of 500 &#181;L at RT for 30 min.</li>
		<li>Perform buffer exchange using PD 10 desalting columns. Place the PD10 column into a 50 mL centrifuge tube using a column adapter.</li>
		<li>Equilibrate the PD10 column by adding 5 mL of PBS buffer and centrifuge at 1000&#160;&#215;&#160;g for 2 min. Repeat this step 3 times.</li>
		<li>Keep the PD10 column in a fresh 50 mL tube and add 2 mL of PBS buffer into the column. Then, add 500 &#181;L of FoxP1 protein from step 1 into the column. Elute the protein bycentrifuging 1000&#160;&#215;&#160;g for 2 min and collect the protein.</li>
		<li>Concentrate the protein by adding it into ultra centrifugal filters (10 kDa MWCO), centrifuge at 7500&#160;&#215;&#160;g for 10 min, and collect the elute.</li>
		<li>Measure the protein concentration, add BODIPY with 30% equivalent of cysteine concentration in the protein, and place it on a rotator for 2 h at 4 &#176;C.</li>
		<li>Perform buffer exchange and concentrate the protein as discussed in steps 5.3.2-5.3.5.</li>
		<li>Check the protein concentration using a spectrophotometer or a colorimetric assay, such as the Bradford or Lowry method, and the concentration of the dye by absorbance at 500 nm.</li>
		<li>Measure the degree of labeling using the following formula: 
		(ADye/&#949;Dye)/(molecular weight of protein/milligram of protein per milliliter) = moles of dye/moles of protein 
		Where ADye and &#949;Dye are the absorbance value of the dye and the molar extinction coefficient of the dye at the maximum absorbance wavelength, respectively59. When measured at 280 nm, the real absorbance of the protein must be corrected by the following formula: 
		Aprotein = A280-(ADye&#183;CF280) 
		Where CF280 is the correction factor for the specified dye, considering its intrinsic absorbance at 280 nm59. 
		NOTE: While we focus on the expression and purification of a cysteine-modified protein, careful consideration must be given to the strategic introduction of cysteine residues into the protein.</li>
	</ol>
	</li>
</ol>
6. Microscope sample chamber preparation
<ol>
	<li>Add 495 &#181;L of ultra-pure water and 5 &#181;L (0.01% v/v final concentration) of Tween-20 (nonionic surfactant) to the well of a chamber slide and mix. Incubate over 30 min at RT.</li>
	<li>Wash the chamber gently with ultra-pure water twice and dry it. 
	NOTE: The chamber is ready to use for the experiment.</li>
</ol>
7. Single-molecule fluorescence anisotropy experiment
<ol>
	<li>First, calibrate the smFA instrument. Determine the instrument response function, either by measuring Raman scattered light or by measuring the fluorescence of a dye with a short lifetime and appropriate emission spectrum (e.g., Erythrosin or Malachite green) additionally quenched by a saturated potassium iodide, KI, solution.</li>
	<li>To correct for background fluorescence, perform measurements of a buffer solution. Calibrate the relative sensitivity of the parallel and perpendicular detectors (G-factor) by acquiring data from a fast-rotating dye. Here, 2 nM Rhodamine 110 was used to calibrate the detectors60.</li>
	<li>For smFA experiments on monomeric FoxP1, add 100 pM BODIPY-labeled FoxP1 and 100 nM un-labeled FoxP1 to 500 &#181;L of standard buffer (20 mM HEPES, 20 mM NaCl, pH 7.8) in a chamber slide.</li>
	<li>For titration assay with DNA and monomeric FoxP1, add 400 nM DNA to 100 pM BODIPY-labeled/100nM un-labeled protein mix and incubate at RT for 10 min.</li>
	<li>For smFA experiments on dimeric FoxP1, add 500 nM un-labeled FoxP1 to a 100 pM BODIPY-labeled FoxP1 in the standard buffer. Next, incubate at 37 &#176;C for 30 min.</li>
	<li>For titration assay with DNA and dimeric FoxP1, add 2000 nM DNA to 100 pM BODIPY-labeled/500 nM un-labeled protein mix and incubate at RT for 10 min.</li>
	<li>Start the measurements and check in the BIFL software that the amount of bursts in a 30 s record is between 60-90. If the sample is more concentrated, dilute it until it reaches that value. This step ensures single-molecule conditions.</li>
	<li>Begin smFA each measurement for either monomeric FoxP1 or dimeric FoxP1 in the absence or presence of DNA for at least 4 h.</li>
	<li>Analyze the smFA measurements as discussed in section 4.</li>
</ol>

All the authors declare that they have no competing financial interests with the contents of this article.

For single-molecule fluorescence anisotropy experiments, it is crucial to consider the chosen fluorophore's photophysical properties carefully. These properties include the emission wavelength, which must align with the detection system, and the excitation wavelength, which should be compatible with the available pulsed lasers. To optimize the dynamic range, the fluorophore should have a long fluorescence lifetime relative to the molecule's rotational diffusion time. This is critical for tracking rotational dynamics and the linkage/orientation of the fluorophore's dipole relative to the biomolecule of interest. Additionally, brightness, photostability, and quantum yield are essential for producing strong signals with a stable signal-to-noise ratio. For these reasons, BODIPY-FL has been chosen as the fluorophore in several studies39,40,42.
Screening the backbone dynamics of biomolecules often requires protein labeling, typically achieved through site-specific labeling. This is usually done by introducing a residue for targeted chemical modification. The most common approach is introducing cysteines at positions of interest, where their thiol side chains can be selectively modified with reagents such as maleimides or iodoacetamides. Less commonly, benzylic halides and bromomethyl ketones are used to form thioether bonds. Other amino acid side chains can also be targeted, but their abundance in proteins is less commonly used. However, alternative approaches, like unnatural amino acids, can also be used73. Proper site selection for labeling is crucial to minimize interference with the biomolecule under study, and appropriate controls must be in place. For example, if the labeled molecule is used in binding assays, complementary label-free methods should verify that the fluorophores do not impact binding affinity. 
After identifying the appropriate sample and implementing the optimal labeling strategy, the next step is to ensure the confocal microscope is properly aligned and calibrated for single-molecule experiments. The protocol describes how to determine the required factor for further analysis. Once the instrument is calibrated, the next step is to measure the sample and process the data to extract as much information as possible from the detected photons. The key parameters, such as micro-time, macro-time, and channel type, as shown in Figure 2, can be used for further analysis and visualization using typical TCSPC electronics.
Recent advances in single-molecule fluorescence spectroscopy can be extensively used to study structural information from the heterogeneous ensembles of biomolecules. However, relatively few studies leverage the insights provided by fluorescence anisotropy, and a complete protein model is required to derive the structural dynamics of biomolecules. Therefore, unraveling the dynamics of interdomain and protein-protein interactions of several transcription factors is challenging.
In conclusion, single-molecule fluorescence anisotropy experiments offer complementary information about the local and global motions of the biomolecular backbone, which are critical for understanding its function.

The functional activity of biomolecules depends on their molecular flexibility and structural dynamics1,2,3. Naturally, biomolecules experience constant thermal fluctuations, ranging from rapid movements to long-term conformational changes influencing their function (Figure 1)4. In biomolecules, local backbone motions contribute to larger-scale global movements, including hinge bending in enzymes and significant conformational changes in motor proteins. Structure determination methods such as nuclear magnetic resonance (NMR)5, X-ray crystallography6, and cryogenic electron microscopy (cryo-EM)7 have revealed multiple conformations in various biomolecules. Nevertheless, connecting the local fluctuations to large conformational dynamics of biomolecules and their role in function are mostly unexplored. Relating dynamics and structure can be challenging, especially for intrinsically disordered proteins (IDPs)8,9,10. Unlike structured proteins, IDPs do not maintain a stable tertiary structure. Instead, they undergo extensive conformational changes with similar free energy levels, enabling a wide range of biological activities11,12.
Several experimental approaches have been employed to investigate the conformational dynamics of proteins by probing their molecular flexibility1,13,14,15,16. Among these, NMR stands out for its ability to provide atomic-level resolution across various timescales, from tens of picoseconds to several hours12. However, determining macromolecular flexibility remains challenging due to the high degrees of freedom and for large-size proteins; thus, NMR is often limited to studying biomolecules of around 100 kDa17.
Given the structural complexity of highly dynamic proteins like IDPs, additional methodological advancements have been developed to explore local and long-range conformational space to understand their function11. Single-molecule multiparameter Fluorescence Spectroscopy (smMFS)18,19,20,21,22 offers extensive information on biomolecules, providing crucial insights into their function, conformational dynamics, binding states, and stoichiometry. However, interpreting the vast amount of structural data obtained from biomolecules is challenging, and factors such as molecular dynamics, fluorophore behavior, and the complex behavior of molecules can further complicate data analysis23,24,25,26,27,28.
We employ single-molecule fluorescence anisotropy (smFA) as a robust method for assessing local and global dynamics along the backbone of biomolecules (Figure 1A). Fluorescence anisotropy, first described by Perrin29 and introduced by Weber30,31 as a bioanalytical tool32, was later adapted for single-molecule studies with the advent of time-resolved fluorescence techniques and the increment of detectors&#39; sensitivity33,34,35,36,37. smFA spans a broad range of timescales -- from picoseconds to several hours -- and complements the data obtained from single-molecule F&#246;rster Resonance Energy Transfer (smFRET) experiments38.
smFA can be visualized in various formats to extract critical information about biomolecular dynamics (Figure 1B). Time-resolved fluorescence anisotropy decays are one-dimensional histograms that capture dynamics on the picosecond to nanosecond timescale39,40. Two-dimensional single-molecule histograms, which correlate fluorescence lifetime with anisotropy for individual molecules, can reveal anisotropy state heterogeneity and provide visual insights into potential dynamics within the observation time in confocal experiments (~ms)41,42. For studying sub-millisecond dynamics, dynamic anisotropy photon distribution analysis (daPDA) can be used, while time-resolved anisotropy Burst Variance Analysis (traBVA) offers a robust method for confirming specific dynamics around milliseconds43 (Figure 1B).
These methods complement more traditional tools, such as polarization-resolved fluorescence correlation spectroscopy (pFCS), which has a broader spectrum44,45,46,47. Overall, multiple data analysis tools for smFA facilitate identifying local and global conformational changes, provided proper calibration is considered.
Here, we apply smFA to study the DNA binding of the human FoxP1 transcription factor48,49,50,51. This&#160;protein&#160;adopts domain-swapped dimer&#160;due to the intrinsically disordered nature of its polypeptide chain, which is notably affected depending on the quaternary state of the protein and the presence of DNA. We generated different single-cysteine mutants to label with BODIPY-FL, performed smFA experiments, and employed daPDA and trBVAa. This approach allowed us to relate dynamics and heterogeneity to FoxP1 dimerization and the DNA binding, highlighting the complex mechanism of action that characterizes this transcription factor.

<table><tbody><tr><td>Alexa Fluor 488</td><td>ThermoFisher Scienctific</td><td>A20100</td><td>https://www.thermofisher.com/order/catalog/product/A20100?SID=srch-srp-A20100</td></tr><tr><td>Amicon Ultra Centrifugal Filter, 10 kDa MWCO</td><td>Millipore Sigma</td><td>UFC901008</td><td>https://www.sigmaaldrich.com/US/en/product/mm/ufc9010</td></tr><tr><td>Ampicillin sodium salt</td><td>Millipore Sigma</td><td>A0166-5G</td><td>https://www.sigmaaldrich.com/US/en/product/sigma/a0166</td></tr><tr><td>BODIP FL Maleimide (BODIPY FL N-(2-Aminoethyl))Maleimide)</td><td>ThermoFisher Scienctific</td><td>B10250</td><td>https://www.thermofisher.com/order/catalog/product/B10250?SID=srch-srp-B10250</td></tr><tr><td>Disposable PD 10 Desalting Columns</td><td>Millipore Sigma</td><td>GE17-0851-01</td><td>https://www.sigmaaldrich.com/US/en/product/sigma/ge17085101</td></tr><tr><td>Dithiothreitol</td><td>Millipore Sigma</td><td>10197777001</td><td>https://www.sigmaaldrich.com/US/en/product/roche/dttro</td></tr><tr><td>DMSO, Anhydrous</td><td>ThermoFisher Scienctific</td><td>D12345</td><td>https://www.thermofisher.com/order/catalog/product/D12345?SID=srch-srp-D12345</td></tr><tr><td>DNAse</td><td>Millipore Sigma</td><td>10104159001</td><td>https://www.sigmaaldrich.com/US/en/product/roche/10104159001</td></tr><tr><td>&lt;em&gt;E. coli&lt;/em&gt; C41 bacterial cells&amp;nbsp;</td><td>Invitrogen</td><td></td><td></td></tr><tr><td>Foresigh Nuvi Ni-Charged IMAC, 5 mL column</td><td>Bio-Rad</td><td>12004037</td><td>https://www.bio-rad.com/en-us/sku/12004037-foresight-nuvia-ni-charged-imac-5-ml-column?ID=12004037</td></tr><tr><td>HEPES</td><td>Millipore Sigma</td><td>7365-45-9</td><td>https://www.sigmaaldrich.com/US/en/product/sigma/h3375</td></tr><tr><td>Imidazole</td><td>Millipore Sigma</td><td>288-32-4</td><td>https://www.sigmaaldrich.com/US/en/product/sigma/i5513</td></tr><tr><td>IPTG</td><td>Millipore Sigma</td><td>I6758-1G</td><td>https://www.sigmaaldrich.com/US/en/product/sial/i6758</td></tr><tr><td>MCE Membrane Filter, 0.22 &amp;mu;m Pore Size</td><td>Millipore Sigma</td><td>GSWP02500</td><td>https://www.sigmaaldrich.com/US/en/product/mm/gswp02500</td></tr><tr><td>NaCl</td><td>Millipore Sigma</td><td>7647-14-5</td><td>https://www.sigmaaldrich.com/US/en/product/sigma/s3014</td></tr><tr><td>Nunc Lab-Tek II Chamber Slide System</td><td>ThermoFisher Scienctific</td><td>154534</td><td>https://www.thermofisher.com/order/catalog/product/154534</td></tr><tr><td>OverExpress C41(DE3) Chemically Competent Cells</td><td>Millipore Sigma</td><td>CMC0017-20X40UL</td><td>https://www.sigmaaldrich.com/US/en/product/sigma/cmc0017</td></tr><tr><td>PMSF</td><td>Millipore Sigma</td><td>329-98-6</td><td>https://www.sigmaaldrich.com/US/en/product/sigma/78830</td></tr><tr><td>Rhodamine 110</td><td>ThermoFisher Scienctific</td><td>419075000</td><td>https://www.thermofisher.com/order/catalog/product/419075000?SID=srch-hj-419075000</td></tr><tr><td>Sodium phosphate dibasic</td><td>Millipore Sigma</td><td>7558-79-4</td><td>https://www.sigmaaldrich.com/US/en/product/sigma/s3264</td></tr><tr><td>Sodium phosphate monobasic dihydrate</td><td>Millipore Sigma</td><td>13472-35-0</td><td>https://www.sigmaaldrich.com/US/en/product/sigma/71505</td></tr><tr><td>TCEP, Hydrochloride, Reagent Grade</td><td>Millipore Sigma</td><td>580567-5GM</td><td>https://www.sigmaaldrich.com/US/en/product/mm/580567</td></tr><tr><td>Tween 20</td><td>Millipore Sigma</td><td>11332465001</td><td>https://www.sigmaaldrich.com/US/en/product/roche/11332465001</td></tr></tbody></table>

time-resolved fluorescence anisotropy from single molecules for characterizing local flexibility in biomolecules

Time-Resolved Fluorescence Anisotropy from Single Molecules for Characterizing Local Flexibility in Biomolecules

We describe a protocol for conducting time-resolved fluorescence anisotropy at the single-molecule level using confocal microscopy to investigate the local flexibility and dynamics of the deoxyribonucleic acid (DNA)-binding forkhead (FKH) domain of the FoxP1 transcription factor. FoxP1 dimerizes through a three-dimensional domain-swapping (3D-DS) mechanism, forming a disordered intermediate with or without DNA. Since 3D-DS involves an intrinsically disordered region, understanding its behavior is crucial for elucidating the structural and functional properties of FoxP1. Using a single-cysteine-labeled FoxP1, we conducted single-molecule fluorescence anisotropy (smFA) experiments, applying dynamic anisotropy Photon Distribution Analysis (daPDA) and time-resolved anisotropy Burst Variance Analysis (traBVA) approaches to probe local flexibility and dynamics. This protocol provides a detailed, step-by-step guide for smFA measurements, emphasizing time-resolved analyses, variance, and probability distribution techniques to capture structural dynamics across different timescales. This approach enabled us to relate dynamics and heterogeneity to FoxP1 dimerization and DNA binding, highlighting the complex action mechanism that characterizes this transcription factor.

Here, we present the protocol to study the local flexibility and dynamics of biomolecules using time-resolved fluorescence anisotropy at the single-molecule level in confocal microscopy mode.

We describe a protocol for conducting time-resolved fluorescence anisotropy at the single-molecule level using confocal microscopy to investigate the ...

time-resolved-fluorescence-anisotropy-from-single-molecules-for

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Protein interactions are at the heart of a cell's function. Calorimetric and spectroscopic techniques are commonly used to characterize them. Here we describe fluorescence anisotropy as a tool to study the interaction between the protein mutated in the Shwachman-Diamond Syndrome (SBDS) and the Elongation factor-like 1 GTPase (EFL1).

Fluorescence Anisotropy as a Tool to Study Protein-protein Interactions

We present the setup and experimental procedure to obtain smFRET data from large donor-acceptor networks with a TIRF microscope. The step-by-step analysis of these measurements with the Bayesian inference software Fast-NPS yields high-resolved structural information via the application of adapted dye models.

Structural Information from Single-molecule FRET Experiments Using the Fast Nano-positioning System

A protocol for high-precision FRET experiments at the single molecule level is presented here. Additionally, this methodology can be used to identify three conformational states in the ligand-binding domain of the N-methyl-D-aspartate (NMDA) receptor. Determining precise distances is the first step towards building structural models based on FRET experiments.

High Precision FRET at Single-molecule Level for Biomolecule Structure Determination

Here, we present a protocol to obtain three-color smFRET data and its analysis with a 3D ensemble Hidden Markov Model. With this approach, scientists can extract kinetic information from complex protein systems, including cooperativity or correlated interactions.

Using Three-color Single-molecule FRET to Study the Correlation of Protein Interactions

Presented here is a protocol for performing single-molecule Förster resonance energy transfer to study HJ resolution. Two-color alternating excitation is used for determining the dissociation constants. Single-color time lapse smFRET is then applied in real-time cleavage assays to obtain the dwell time distribution prior to HJ resolution.

Single-Molecule F&ouml;rster Resonance Energy Transfer Methods for Real-Time Investigation of the Holliday Junction Resolution by GEN1

This article provides step-by-step instructions for making fully-corrected accurate FRET measurements on individual, freely diffusing biomolecules using the open-source, inexpensive smfBox, from switch on, through alignment and focusing, to data collection and analysis.

Making Precise and Accurate Single-Molecule FRET Measurements using the Open-Source smfBox

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 α-Synuclein, which is otherwise known as globularly unstructured and unstable when measured using the longer range FRET ruler.

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

We present an experimental protocol and data analysis workflow to perform live cell dual-color fluorescence cross correlation spectroscopy (FCCS) combined with Förster Resonance Energy transfer (FRET) to study membrane receptor dynamics in live cells using modern fluorescence labeling techniques.

Dual-Color Fluorescence Cross-Correlation Spectroscopy to Study Protein-Protein Interaction and Protein Dynamics in Live Cells

This article presents a method for spatiotemporal analysis of mobile, single-molecule Förster resonance energy transfer (smFRET)-based probes using widefield fluorescence microscopy. The newly developed software toolkit allows the determination of smFRET time traces of moving probes, including the correct FRET efficiency and the molecular positions, as functions of time.

Automated Two-dimensional Spatiotemporal Analysis of Mobile Single-molecule FRET Probes

This study presents a detailed procedure to perform single-molecule fluorescence resonance energy transfer (smFRET) experiments on G protein-coupled receptors (GPCRs) using site-specific labeling via unnatural amino acid (UAA) incorporation. The protocol provides a step-by-step guide for smFRET sample preparation, experiments, and data analysis.

Visualizing the Conformational Dynamics of Membrane Receptors Using Single-Molecule FRET

Learn time-resolved fluorescence anisotropy for studying biomolecular flexibility using single-molecule confocal microscopy. This method reveals protein dynamics and conformational changes in FoxP1.

Time-Resolved Fluorescence Anisotropy from Single Molecules for Characterizing Local Flexibility in Biomolecules

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