Authors acknowledge the financial support from CONACyT project numbers 167359 and 177138, and from DGAPA-UNAM project number IN201615.

1. SBDS-FlAsH Tag Protein Expression and Purification
NOTE: For the anisotropy experiments, a FlAsH-tag corresponding to the sequence Cys-Cys-Pro-Gly-Cys-Cys was added to the C-terminus of the human SBDS coding sequence by PCR. This construct was subcloned into the expression vector pRSET-A and transformed into Escherichia coli C41 cells to express a protein encoding a N-terminal hexahistidine tag (His-tag), the human SBDS coding sequence and a C-terminus FlAsH tag 10.
<ol>
	<li>SBDS-FlAsH protein expression
	<ol>
		<li>Transform competent E. coli C41 cells with the plasmid pRSET-HisSBDS-FlAsH using a standard heat shock protocol 19. Plate the cells in solid Luria-Bertani (LB) media supplemented with 100 µg/ml ampicillin. LB solid media composition consists of 10 g NaCl, 5 g yeast extract, 10 g tryptone and 20 g agar for 1 L volume.</li>
		<li>Culture transformed bacteria at 37 °C until absorbance at 600 nm (A600) reaches 0.5-0.7 in 1 L of LB liquid media supplemented with 100 µg/ml ampicillin.</li>
		<li>Induce protein expression by adding 0.5 mM isopropyl β-D-1-thiogalactopyranoside to the culture and continue the incubation for further 5 hr.</li>
		<li>Collect the bacterial suspension by centrifugation at 3,800 x g for 10 min at 4 °C. Remove the supernatant. At this point, either store the cell pellet at -20 °C or use immediately for protein purification.</li>
	</ol>
	</li>
	<li>SBDS-FlAsH protein purification 
	NOTE: All chromatographic steps are performed using a fast protein liquid chromatography (FPLC) system or a peristaltic pump. Ni2+-affinity chromatography uses a 5 ml fast flow column. Anionic exchange chromatography uses a 5 ml strong sulfopropyl cation exchanger column. A flow rate of 3 ml/min was used for all the chromatographic steps.
	<ol>
		<li>Resuspend the cells in 35 ml of SBDS Lysis buffer (50 mM phosphate buffer pH 7.5, 300 mM NaCl, 20 mM imidazole) supplemented with 1 mM phenylmethylsulfonyl fluoride (PMSF) and lyse by sonication for a total time of 4 min using cycles of 10 sec ON and 30 sec OFF, at 4 °C.</li>
		<li>Centrifuge the sample at 9,000 x g for 50 min at 4 °C.</li>
		<li>Keep the supernatant and discard the pellet to remove cellular debris.</li>
		<li>Equilibrate the Ni2+ affinity column with 3 column volumes (CV) of SBDS Lysis buffer and introduce the whole clarified supernatant onto the column.</li>
		<li>Remove unbound protein by washing with 3 CV of SBDS Lysis buffer and elute with 3 CV of SBDS Elution buffer (50 mM phosphate buffer pH 7.5, 300 mM NaCl, 250 mM imidazole).</li>
		<li>Dilute the eluted protein 6-fold with 50 mM Phosphate buffer pH 6.5 and remove possible aggregates by filtration through a 0.22 µm cellulose membrane.</li>
		<li>Equilibrate the sulfopropyl cation exchanger column with 3 CV of Low salt S column buffer (50 mM phosphate buffer pH 6.5, 50 mM NaCl) and introduce the protein sample from the previous step.</li>
		<li>Wash unbound material with 3 CV of Low salt S column buffer and elute the protein in one step with 50 mM phosphate buffer pH 6.5, 1 M NaCl.</li>
		<li>Dilute the eluted protein 3.3-fold with 50 mM Phosphate buffer pH 6.5. Concentrate the protein with ultrafiltration devices by centrifugation at 3,800 x g for 15 min. Flash freeze the protein in liquid nitrogen and store it at -80 °C until further use.</li>
		<li>Verify the purity of the protein by SDS-PAGE analysis and Coomassie staining 20.</li>
	</ol>
	</li>
</ol>
2. EFL1 Protein Expression and Purification
NOTE: EFL1 was expressed under the regulation of the Gal 1/10 divergent promoter 21 and the Saccharomyces cerevisiae MATA 3'UTR in the vector pRS426. The recombinant protein encodes the human EFL1 isoform 1 fused to a Tobacco Etch Virus protease (TEV) recognition site and a hexahistidine tag at the C-terminus.
<ol>
	<li>EFL1 protein expression
	<ol>
		<li>Transform S. cerevisiae BCY123 cells with the plasmid pRS426-EFL1TevHis using a standard Lithium acetate protocol 22. Plate all the transformed cells in synthetic drop out media without uracil (SD-URA) supplemented with 2% (w/v) glucose. Composition of the SD-URA media consists of 8 g yeast nitrogen base without amino acids, 11 g casamino acids, 55 mg adenine sulfate, 55 mg tyrosine, 60 mg leucine and 60 mg tryptophan for 1 L volume.</li>
		<li>Culture transformed yeast at 30 °C until A600 reaches 1.8 in 1 L of SD-URA media supplemented with 0.5% (w/v) glucose.</li>
		<li>Induce protein expression by adding 2.8% (w/v) galactose to the culture and continue the incubation for further 18 hr at 30 °C.</li>
		<li>Collect the yeast suspension by centrifugation at 3,800 x g for 10 min at 4 °C. Remove the supernatant. At this point, either store the cell pellet at -20 °C or use immediately for protein purification.</li>
	</ol>
	</li>
	<li>EFL1 protein purification 
	NOTE: All chromatographic steps are performed using a FPLC system or a peristaltic pump. Ni2+-affinity chromatography uses a 5 ml fast flow column at a flow rate of 3 ml/min. Size exclusion chromatography uses a 125 ml column pre-packed with Superdex 200 resin at a flow rate of 1 ml/min.
	<ol>
		<li>Resuspend the cells in 50 ml of EFL1 Lysis buffer (50 mM Tris-HCl pH 8, 300 mM NaCl, 20 mM imidazole, 5 mM MgCl2, 10% glycerol) supplemented with 1 mM PMSF and 1 mM benzamidine and disrupt the cells by friction on a bead beater using glass beads (Ø = 0.5 mm) for a total time of 6 min using cycles of 2 min ON and 15 min OFF, at 4 °C.</li>
		<li>Centrifuge the sample at 9,000 x g for 50 min at 4 °C.</li>
		<li>Keep the supernatant and discard the pellet to remove cellular debris.</li>
		<li>Equilibrate the Ni2+ affinity column with 3 CV of EFL1 Lysis buffer and introduce all the clarified supernatant onto the column.</li>
		<li>Remove unbound protein by washing with 3 CV of EFL1 Lysis buffer and elute with 3 CV of EFL1 Elution buffer (50 mM Tris-HCl pH 8, 300 mM NaCl, 250 mM imidazole, 5 mM MgCl2, 10% glycerol).</li>
		<li>Equilibrate the size exclusion column with 1.5 CV of Anisotropy buffer (50 mM Tris-HCl pH 7.5, 300 mM NaCl, 5 mM MgCl2, 10% glycerol, 5 mM β-mercaptoethanol).</li>
		<li>Concentrate to 1 ml the EFL1 protein eluted from the Ni2+ affinity column with ultrafiltration devices by centrifugation at 3,800 x g to the desired volume. Introduce the sample on the size exclusion column.</li>
		<li>Collect the eluted protein and concentrate by ultrafiltration to a final concentration of approximately 30 µM. Flash freeze the protein in liquid nitrogen and store at -80 °C until further use. Verify the purity of the protein by SDS-PAGE analysis and Coomassie staining 20.</li>
	</ol>
	</li>
</ol>
3. Labeling of SBDS-FlAsH with the FlAsH Fluorescent Dye 4',5'-Bis(1,3,2 dithioarsolan-2-yl) Fluorescein
<ol>
	<li>Mix 3 nmol of the SBDS-FlAsH protein with 3 nmol of the 4',5'-bis(1,3,2 dithioarsolan-2-yl) fluorescein dye in 5 µl volume of Anisotropy buffer.</li>
	<li>Let the reaction proceed for 8 hr at 4 °C. Dialyze the sample against Anisotropy buffer over night to remove the free dye.</li>
	<li>Use the Lambert-Beer law to quantify the % of labeled protein. Measure the absorbance at 280 nm and 508 nm in a spectrophotometer using a quartz cuvette of appropriate volume. NOTE: Consider the following molar absorption coefficients (M-1 cm-1): 
	<img alt="Absorption coefficients formula, ε for dye and protein, in spectroscopic analysis." src="/files/ftp_upload/54640/54640eq1B.jpg" data-altupdatedat="1747929244000" data-alt="Spectroscopy equations for absorption coefficients in dye analysis, showing ε values and wavelengths."></li>
	<li>Calculate the concentration of labeled SBDS-FlAsH protein using Equation 2. 
	<img alt="Absorption equation at 508 nm, calculating dye concentration, related to spectroscopy analysis." src="/files/ftp_upload/54640/54640eq2.jpg" data-altupdatedat="1747929244000" data-alt="Spectrophotometry equation, A508nm=C_SBDS-FLASH×l×ε_dye, related to optical absorption study."></li>
	<li>Calculate the concentration of total SBDS protein using Equation 3 by substituting the calculated CSBDS-FlAsH from previous step. 
	<img alt="Absorbance equation for protein assay; equation; spectroscopy calculation, 280 nm absorbance." src="/files/ftp_upload/54640/54640eq3.jpg" data-altupdatedat="1747929244000" data-alt="Static equilibrium equation, absorbance calculation formula for spectroscopic analysis, equation depiction."></li>
	<li>Calculate the percentage of labeled protein using Equation 4. 
	<img alt="Percentage labeling formula, shown as % Labeling = (C_SBDS-FLaSH / C_SBDS) * 100, Equation 4." src="/files/ftp_upload/54640/54640eq4.jpg" data-altupdatedat="1747929244000" data-alt="Percent labeling formula in biochemical analysis, Equation 4, for sample labeling calculation."></li>
</ol>
4. Fluorescence Anisotropy Experiments
NOTE: Anisotropy experiments were done in a spectrofluorometer equipped with a polarization toolbox and data collection was performed using the anisotropy program provided in the software of the equipment. The excitation wavelength was set at 494 nm with a spectral bandwidth of 8 nm and the emission was recorded using a band-pass filter of 530±25 nm. Measurements were done at 25 °C in a 200 µl cuvette with a 5 mm path length 10.
<ol>
	<li>In a fluorescence cuvette, place 200 µl of 30 nM SBDS-FlAsH in anisotropy buffer and titrate 2 µl of 30 µM EFL1. Mix thoroughly and let the reaction stand for 3 min before measuring the anisotropy value.</li>
	<li>Repeat step 4.1 until a total volume of 40 µl of EFL1 has been added.</li>
</ol>
5. Data Analysis
<ol>
	<li>Fit the data to the appropriate binding model using a nonlinear least squares regression algorithm. Equations for the most common binding models are presented in Table 1.</li>
	<li>Evaluate the best model that describes the interaction between the proteins by inspecting the residuals of the fit 23. Support the chosen model with additional experiments.</li>
</ol>
Table 1. Common protein-protein interaction binding models and the mathematical equations that describe them. <a href="http://cloudfront.jove.com/files/ftp_upload/54640/54640table1large.jpg" target="_blank">Please click here to view a larger version of this table.</a> 
<img alt="Binding model table showing equilibrium equations and calculations for SBDS-FIaSH analysis." src="/files/ftp_upload/54640/54640table1.jpg" data-altupdatedat="1747929244000" data-alt="Table 1">

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The authors have nothing to disclose and they have no competing financial interests.

Most biochemical experiments with proteins require not only pure protein but also large amounts of them, irrespective of the technique used. For this reason, the proteins used for this type of experiments are obtained by heterologous expression, as it was the case presented here. Florescence spectroscopy requires the presence of a fluorophore in the studied molecule. Aromatic residues constitute the intrinsic fluorophores of a protein, however, using their signal to study protein-protein interactions complicates the anal...

Cells contain a multitude of biomacromolecules that constantly interact with each other. This association gives rise to complexes that participate in the cellular pathways responsible for their functioning in signal transduction, regulation of gene expression and cell migration amongst others. All protein-protein interactions that occur in a cell comprise a network known as the interactome. In Saccharomyces cerevisiae more than 70% of its proteins have been shown to have interacting partners 1. Understanding the interactome of a cell and their functions provide relevant information on the complexity and diversity of living organisms. Several methodologies have been described to identify and characterize protein-protein interactions. Different high through put methods such as yeast two-hybrid 2, protein-fragment complementation assays 3, affinity purification 4 coupled to mass spectrometry and protein microarrays are used to identify an interaction 5,6. Once identified, it is necessary to validate it and this may vary on a case-by-case basis. Typically, these experiments involve disrupting the interaction itself at the level of the individual members of the interaction pair, e.g., by gene deletion or overexpression of one of the proteins, and then looking for changes in the properties or function of the other member at the cellular level. Subsequently, biophysical techniques 7 are used to characterize the protein-protein interaction at the molecular level. To this end, the structure of protein complexes are determined by X-ray crystallography, nuclear magnetic resonance and cryo-electron microscopy while calorimetry and fluorescence spectroscopy are used to quantitatively and mechanistically describe them.
In this work, fluorescence anisotropy was used as a technique to characterize the interaction between the GTPase EFL1 and the SBDS protein. These proteins participate in the synthesis of ribosomes by promoting the release of eukaryotic initiation factor 6 from the surface of the 60S ribosomal subunit 8. The SBDS protein is mutated in a disease known as the Shwachman-Diamond Syndrome 9 and acts as a guanine nucleotide exchange factor for EFL1 decreasing its affinity for guanosine diphosphate 10,11. Disease mutations in SBDS abolish the interaction with EFL1 and thus prevent its activation.
Fluorescence anisotropy is commonly used in biological applications to study protein-peptide or protein-nucleic acid interactions. It is based on the principle that a fluorophore excited with polarized light results in a partially polarized emission. Fluorescence anisotropy is defined by Equation 1:
<img alt="anisotropy equation r=IVV-IVH/IVV+2IVH formula, molecular fluorescence polarization" src="/files/ftp_upload/54640/54640eq1.jpg" data-altupdatedat="1747929244000" data-alt="anisotropy equation r=(I&lt;sub&gt;VV&lt;/sub&gt;-I&lt;sub&gt;VH&lt;/sub&gt;)/(I&lt;sub&gt;VV&lt;/sub&gt;+2I&lt;sub&gt;VH&lt;/sub&gt;); formula, optics.">
where IVV and IVH are the fluorescence intensities of the vertically (VV) and horizontally (VH) polarized emission when the sample is excited with vertically polarized light 12. Fluorescence anisotropy is sensitive to factors that affect the rate of the rotational diffusion of the fluorophore and thus depends on the temperature, the viscosity of the solution and the apparent molecular size of the fluorophore. The apparent size of a protein containing a fluorophore increases when it interacts with another protein and such change can then be evaluated as a change in anisotropy. More specifically, a fluorophore that rotates slowly in solution relative to its fluorescent lifetime will have a large IVV value and small IVH value and therefore will exhibit a relatively large anisotropy. For fluorophores that tumble rapidly relative to their fluorescent lifetime, IVV and IVH will be similar and their anisotropy value will be small 12 (Figure 1). In addition, for a good anisotropy signal to noise measurement, it is necessary to have a fluorophore with a fluorescence lifetime similar to the rotational correlation time of the molecule of interest. Otherwise, it is not possible to accurately record the difference in anisotropy between the free protein and that in the complex. For example, the anisotropy of a fluorescent probe with a lifetime close to 4 nsec such as fluorescein or rhodamine attached to a low molecular weight compound of 100 Da is 0.05. Binding to a molecule of 160 kDa will increase its anisotropy value to 0.29; a difference that can be accurately measured. In contrast, the same fluorescent probe involved in a binding reaction whose increase in molecular size varies from 65 to 1,000 kDa will only result in an anisotropy change of 0.28 to 0.3, which is too small to be accurately measured. In this scenario, a probe with a lifetime of 400 nsec would be more suitable 12.
<img alt="Static equilibrium diagram; spectroscopy setup; DNA separation; optical excitation measurement." src="/files/ftp_upload/54640/54640fig1.jpg" data-altupdatedat="1747929244000" data-alt="Polarization-resolved spectroscopy diagram; optical excitation, monochromator, detector setup."> 
Figure 1. Schematic representation of the equipment used to measure fluorescence anisotropy and the procedure. Schematic representation of the equipment used to perform a protein-protein interaction experiment measuring fluorescence anisotropy. Fluorophores that tumble fast display small anisotropy that increases upon binding to an interaction partner. <a href="http://cloudflare.jove.com/files/ftp_upload/54640/54640fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Fluorescence applications require the presence of a fluorophore in any of the molecules studied. To study protein-protein interactions there are three type of fluorophores: 1) the tryptophan residues present in the proteins, 2) chemically attached fluorophores and 3) fluorescent fusion partners such as green fluorescent protein (GFP) and its derivatives. Most proteins have tryptophan residues on its structure, thus the easiest way to measure an interaction is by monitoring the changes in the corresponding fluorescence spectra or by monitoring changes in the fluorescence intensity of the tryptophan residues. However, tryptophan residues may be present in both proteins complicating the analysis. On the other hand, for a fluorophore to change its fluorescent properties due to an interaction it needs to be located on or near the binding site and it could interfere with the interaction itself. This needs special attention when using bulky fluorophores such as GFP. If none of these fluorophores can be used for binding studies it is necessary, then, to introduce extrinsic fluorophores to the one of the proteins involved. Many chemically synthesized fluorophores exist and can be covalently attached to proteins through their reactive groups such as the amine groups (side chain of lysines or N-terminus) and the thiol groups in cysteine. Fluorophore derivatives with isothiocyanate and succinimidyl esters react with amide groups while iodoacetamide and maleimide are thiol-reactive groups 13. The most common dyes used in fluorescence applications are derivatives of the fluorescein and the rhodamine green dyes, coumarins, BODIPY fluorophores and Alexa Fluor dyes. A detailed list of commercially available fluorophores and their use can be found in references 14,15. For successful labeling, the reactive group must be exposed on the surface of the protein, but due to the large number of reactive functional groups typically present in polypeptides it is very hard to get site-specific modification. The protein of interest in this study, SBDS, contains 5 free cysteines and 33 lysines that may result in multiple site labeling. Non-uniform labeling may affect the binding and will complicate data analysis as different fluorophore molecules may elicit different fluorescent intensity signals upon binding. To overcome this problem, we used the FlAsH fluorophore, 4',5'-bis(1,3,2 dithioarsolan-2-yl) fluorescein to site-direct label the SBDS protein. This is an arsenoxide dye with a high affinity for four spaced cysteines in a motif know as FlAsH-tag consisting of the sequence CCXXCC where X is any amino acid other than cysteine 16,17. This tetracysteine motif is added to the N- or C-terminus of the protein by genetic engineering together with an appropriate linker to prevent the disruption of the overall fold of the protein. The pair consisting of FlAsH dye and FlAsH-tag was originally designed to site-specific label proteins in living cells 17 but it can also be used to label purified proteins in vitro as it is exemplified here. Additionally, enzymatic strategies have also been developed to enable site-specific functionalization of proteins 18.
In this manuscript we describe the usefulness of fluorescence anisotropy as a tool to study protein-protein interactions. Binding can be assessed by simple inspection of the binding curve shape while quantitative information can be obtained from the fit of the experimental data.

<table><tbody><tr><td>0.5 mm Glass beads</td><td>Biospec Products</td><td>11079105</td><td></td></tr><tr><td>Tris Base</td><td>Formedium</td><td>TRIS01</td><td>Ultra pure</td></tr><tr><td>Glycerol</td><td>Sigma-Aldrich</td><td>G5516</td><td></td></tr><tr><td>dye 4&amp;rsquo;,5&amp;rsquo;-bis(1,3,2 dithioarsolan-2-yl) fluorescein</td><td>ThermoFischer Scientific</td><td>LC6090</td><td>This kit contains the dye to label a FlAsH tag</td></tr><tr><td>Ampiciline</td><td>IBI Shelton Scientific, Inc</td><td>IB02040</td><td></td></tr><tr><td>D(+)-Glucose Anhydrous</td><td>Formedium</td><td>GLU03</td><td></td></tr><tr><td>D(+)-Galactose</td><td>Formedium</td><td>GAL03</td><td></td></tr><tr><td>L-Leucine</td><td>Formedium</td><td>DOC0157</td><td></td></tr><tr><td>L-Tryptofan&amp;nbsp;</td><td>Formedium</td><td>DOC0189</td><td></td></tr><tr><td>Bezamidine hydrochloride</td><td>Sigma-Aldrich</td><td>B6506-5G</td><td></td></tr><tr><td>PMSF</td><td>Gold Biotechnology, Inc</td><td>P-470-25</td><td>Phenylmethylsulfonyl fluoride</td></tr><tr><td>NaCl</td><td>Formedium</td><td>NAC02</td><td>Sodium Chloride&amp;nbsp;</td></tr><tr><td>Glycerol</td><td>Tecsiquim, S.A. de C.V.</td><td>GT1980-6</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Merck Millipore Corporation</td><td>1725711000</td><td>Magnesium Chloride</td></tr><tr><td>Imidazole</td><td>Sigma-Aldrich</td><td>I2399-500G</td><td></td></tr><tr><td>2-Mercaptoethanol</td><td>Sigma-Aldrich</td><td>M6250-100ML</td><td></td></tr><tr><td>K&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>P3786-500G</td><td>Potassium phosphate dibasic</td></tr><tr><td>NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>S3139-500G</td><td>Sodium phosphate monobasic</td></tr><tr><td>Yeast nitrogen base without amino acids</td><td>Formedium</td><td>CYN0410</td><td></td></tr><tr><td>Yeast extract</td><td>Formedium</td><td>YEM03</td><td>Micro Granulated</td></tr><tr><td>L-Tyroisne</td><td>Formedium</td><td>DOC0193</td><td></td></tr><tr><td>Adenine sulphate</td><td>Formedium</td><td>DOC0230</td><td></td></tr><tr><td>Casamino acids</td><td>Formedium</td><td>CAS03</td><td></td></tr><tr><td>Tryptone</td><td>IBI Shelton Scientific, Inc</td><td>IB49182</td><td></td></tr><tr><td>IPTG</td><td>Formedium</td><td>IPTG025</td><td></td></tr><tr><td>Filtration units</td><td>Merck Millipore Corporation</td><td>UFC901096</td><td>Amicon Ultra-15, membrana PLGC Ultracel-PL, 10&amp;nbsp;kDa</td></tr><tr><td>Membrane Filter</td><td>Merck Millipore Corporation</td><td>GSWP04700</td><td>Membrane Filter, mixed cellulose esters, Hydrophilic, 0.22&amp;nbsp;&amp;micro;m, 47&amp;nbsp;mm, white, plain</td></tr><tr><td>Ni&lt;sup&gt;2+&lt;/sup&gt; affinity column</td><td>QIAGEN</td><td>30760</td><td>Cartridge pre-filled with 5&amp;nbsp;ml Ni-NTA Superflow</td></tr><tr><td>Strong Sulfopropyl cation exchanger column</td><td>GE Healthcare Life Science</td><td>17-5157-01</td><td>HiTrap SP Sepharose FF 5 ml</td></tr><tr><td>Size Exclusion column</td><td>GE Healthcare Life Science</td><td>28989335</td><td>HiLoad 16/600 Superdex 200 PG</td></tr><tr><td>Fluorescence cell</td><td>Hellma Analytics</td><td>111-057-40</td><td></td></tr><tr><td>Spectrophotometer</td><td>Agilent Technologies</td><td>G6860AA</td><td>Cary 60 UV-Vis</td></tr><tr><td>Shaker</td><td>ThermoFischer Scientific</td><td>SHKA4000-7</td><td>MaxQ 4000 Benchtop temperature range Ambient-15&amp;deg; to 60&amp;deg;C</td></tr><tr><td>Centrifuge</td><td>ThermoFischer Scientific</td><td>75004271</td><td>Heraeus Megafuge 16R</td></tr><tr><td>FPLC</td><td>Pharmacia Biotech</td><td>Discontinued</td><td>FPLC system conductivity UV-MM II monitor P500 pump fraction</td></tr><tr><td>Spectrofluorometer</td><td>Olis</td><td>No applicable</td><td>Olis DM 45 with Polarization Toolbox</td></tr></tbody></table>

fluorescence anisotropy as a tool to study protein-protein interactions

Protein-protein interactions play an essential role in the function of a living organism. Once an interaction has been identified and validated it is necessary to characterize it at the structural and mechanistic level. Several biochemical and biophysical methods exist for such purpose. Among them, fluorescence anisotropy is a powerful technique particularly used when the fluorescence intensity of a fluorophore-labeled protein remains constant upon protein-protein interaction. In this technique, a fluorophore-labeled protein is excited with vertically polarized light of an appropriate wavelength that selectively excites a subset of the fluorophores according to their relative orientation with the incoming beam. The resulting emission also has a directionality whose relationship in the vertical and horizontal planes defines anisotropy (r) as follows: r=(IVV-IVH)/(IVV+2IVH), where IVV and IVH are the fluorescence intensities of the vertical and horizontal components, respectively. Fluorescence anisotropy is sensitive to the rotational diffusion of a fluorophore, namely the apparent molecular size of a fluorophore attached to a protein, which is altered upon protein-protein interaction. In the present text, the use of fluorescence anisotropy as a tool to study protein-protein interactions was exemplified to address the binding between the protein mutated in the Shwachman-Diamond Syndrome (SBDS) and the Elongation factor like-1 GTPase (EFL1). Conventionally, labeling of a protein with a fluorophore is carried out on the thiol groups (cysteine) or in the amino groups (the N-terminal amine or lysine) of the protein. However, SBDS possesses several cysteines and lysines that did not allow site directed labeling of it. As an alternative technique, the dye 4&#39;,5&#39;-bis(1,3,2 dithioarsolan-2-yl) fluorescein was used to specifically label a tetracysteine motif, Cys-Cys-Pro-Gly-Cys-Cys, genetically engineered in the C-terminus of the recombinant SBDS protein. Fitting of the experimental data provided quantitative and mechanistic information on the binding mode between these proteins.

To perform any anisotropy experiment it is important to rule out large changes in the fluorescence intensity of the fluorophore since the observed anisotropy of a mixture of species is represented by Equation 5:
<img alt="Static equilibrium; r=ΣFi*ri, Equation 5; equation for moment calculation; concise educational use." src="/files/ftp_upload/54640/54640eq5.jpg" data-altupdatedat="1747929244000" data-alt="static equilibrium eq...

Watch this Scientific Journal Video about Fluorescence Anisotropy as a Tool to Study Protein-protein Interactions at JoVE.com

Fluorescence Anisotropy as a Tool to Study Protein-protein Interactions | Biochemistry | Video

Fluorescence Anisotropy as a Tool to Study Protein-protein Interactions

Protein interactions are at the heart of a cell&#39;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).

Protein-protein interactions play an essential role in the function of a living organism. Once an interaction has been identified and validated it is ...

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30.5K Views.  Universidad Nacional Autónoma de México.  Eiwit interacties vormen de kern van de functie van een cel. Calorimetrische en spectroscopische technieken worden gewoonlijk gebruikt om ze te karakteriseren. Hier beschrijven we fluorescentie anisotropie als middel om de interactie tussen het eiwit gemuteerd in de Shwachman-Diamond Syndroom (SBDS) en elongatiefactor-1 achtige GTPase (EFL1) te bestuderen. 

Video: Fluorescence Anisotropy as a Tool to Study Protein-protein Interactions

Utilizing the Ethylene-releasing Compound, 2-Chloroethylphosphonic Acid, as a Tool to Study Ethylene Response in Bacteria

Ethylene (C2H4) is a gaseous phytohormone that is involved in numerous aspects of plant development, playing a dominant role in senescence and fruit ripening. Exogenous ethylene applied during early plant development triggers the triple response phenotype; a shorter and thicker hypocotyl with an exaggerated apical hook. Despite the intimate relationship between plants and bacteria, the effect of exogenous ethylene on bacteria has been greatly overlooked. This is partly due to the difficulty of controlling gaseous ethylene within the laboratory without specialized equipment. 2-Chloroethylphosphonic acid (CEPA) is a compound that decomposes into ethylene, chlorine, and phosphate in a 1:1:1:1 molar ratio when dissolved in an aqueous medium of pH 3.5 or greater. Here we describe the use of CEPA to produce in situ ethylene for the investigation of ethylene response in bacteria using the fruit-associated, cellulose-producing bacterium Komagataeibacter xylinus as a model organism. The protocols described herein include both the verification of ethylene production from CEPA via the Arabidopsis thaliana triple response assay and the effects of exogenous ethylene on K. xylinus cellulose production, pellicle properties and colonial morphology. These protocols can be adapted to examine the effect of ethylene on other microbes using appropriate growth media and phenotype analyses. The use of CEPA provides researchers with a simple and efficient alternative to pure ethylene gas for the routine determination of bacterial ethylene response.

The protocols outlined herein facilitate the convenient investigation of bacterial ethylene responses by utilizing 2-chloroethylphosphonic acid (CEPA). Ethylene is produced in situ through the decomposition of CEPA in an aqueous bacterial growth medium, circumventing the requirement for pure ethylene gas.

Ethylene (C2H4) is a gaseous phytohormone that is involved in numerous aspects of plant development, playing a dominant role in senescence and fruit ...

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating these studies, reincorporation of detergent-solubilized membrane proteins into liposomes is a stochastic process where protein topology is impossible to enforce. This paper offers an alternative method to these challenging techniques that utilizes a liposome-based scaffold. Protein solubility is enhanced by deletion of the transmembrane domain, and these amino acids are replaced with a tethering moiety, such as a His-tag. This tether interacts with an anchoring group (Ni2+ coordinated by nitrilotriacetic acid (NTA(Ni2+)) for His-tagged proteins), which enforces a uniform protein topology at the surface of the liposome. An example is presented wherein the interaction between Dynamin-related protein 1 (Drp1) with an integral membrane protein, Mitochondrial Fission Factor (Mff), was investigated using this scaffold liposome method. In this work, we have demonstrated the ability of Mff to efficiently recruit soluble Drp1 to the surface of liposomes, which stimulated its GTPase activity. Moreover, Drp1 was able to tubulate the Mff-decorated lipid template in the presence of specific lipids. This example demonstrates the effectiveness of scaffold liposomes using structural and functional assays and highlights the role of Mff in regulating Drp1 activity.

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

This paper describes a method for assessing the interactions and assemblies of integral membrane proteins in vitro with various partner factors in a lipid-proximal environment.

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating ...

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

Single-molecule F&#246;rster resonance energy transfer (smFRET) has become a widely used biophysical technique to study the dynamics of biomolecules. For many molecular machines in a cell proteins have to act together&#160;with interaction partners in a functional cycle to fulfill their task. The extension of two-color to multi-color smFRET makes it possible to simultaneously probe more than one interaction or conformational change. This not only adds a new dimension to smFRET experiments but it also offers the unique possibility to directly study the sequence of events and to detect correlated interactions when using an immobilized sample and a total internal reflection fluorescence microscope (TIRFM). Therefore, multi-color smFRET is a versatile tool for studying biomolecular complexes in a quantitative manner and in a previously unachievable detail.
Here, we demonstrate how to overcome the special challenges of multi-color smFRET experiments on proteins. We present detailed protocols for obtaining the data and for extracting kinetic information. This includes trace selection criteria, state separation, and the recovery of state trajectories from the noisy data using a 3D ensemble Hidden Markov Model (HMM). Compared to other methods, the kinetic information is not recovered from dwell time histograms but directly from the HMM. The maximum likelihood framework allows us to critically evaluate the kinetic model and to provide meaningful uncertainties for the rates.
By applying our method to the heat shock protein 90 (Hsp90), we are able to disentangle the nucleotide binding and the global conformational changes of the protein. This allows us to directly observe the cooperativity between the two nucleotide binding pockets of the Hsp90 dimer.

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.

Single-molecule F&#246;rster resonance energy transfer (smFRET) has become a widely used biophysical technique to study the dynamics of biomolecules. For ...

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 ...

A Fluorescence Fluctuation Spectroscopy Assay of Protein-Protein Interactions at Cell-Cell Contacts

A variety of biological processes involves cell-cell interactions, typically mediated by proteins that interact at the interface between neighboring cells. Of interest, only few assays are capable of specifically probing such interactions directly in living cells. Here, we present an assay to measure the binding of proteins expressed at the surfaces of neighboring cells, at cell-cell contacts. This assay consists of two steps: mixing of cells expressing the proteins of interest fused to different fluorescent proteins, followed by fluorescence fluctuation spectroscopy measurements at cell-cell contacts using a confocal laser scanning microscope. We demonstrate the feasibility of this assay in a biologically relevant context by measuring the interactions of the amyloid precursor-like protein 1 (APLP1) across cell-cell junctions. We provide detailed protocols on the data acquisition using fluorescence-based techniques (scanning fluorescence cross-correlation spectroscopy, cross-correlation number and brightness analysis) and the required instrument calibrations. Further, we discuss critical steps in the data analysis and how to identify and correct external, spurious signal variations, such as those due to photobleaching or cell movement.
In general, the presented assay is applicable to any homo- or heterotypic protein-protein interaction at cell-cell contacts, between cells of the same or different types and can be implemented on a commercial confocal laser scanning microscope. An important requirement is the stability of the system, which needs to be sufficient to probe diffusive dynamics of the proteins of interest over several minutes.

This protocol describes a fluorescence fluctuation spectroscopy-based approach to investigate interactions among proteins mediating cell-cell interactions, i.e. proteins localized in cell junctions, directly in living cells. We provide detailed guidelines on instrument calibration, data acquisition and analysis, including corrections to possible artefact sources.

A variety of biological processes involves cell-cell interactions, typically mediated by proteins that interact at the interface between neighboring ...

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 ...

Fluorescence Anisotropy to Determine Transcription Factor-DNA Binding Affinity

Source: Jung, C. et al., <a href="https://app.jove.com/v/58763">High Sensitivity Measurement of Transcription Factor-DNA Binding Affinities by Competitive Titration Using Fluorescence Microscopy.</a> J. Vis. Exp. (2019).
This video describes high-performance fluorescence anisotropy that helps to monitor the interaction between transcription factors and DNA. The assay monitors the interactions of a fluorophore-labeled DNA molecule with its specific transcription factor by measuring the degree of polarization due to molecular rotation or anisotropy of the fluorophore-labeled DNA.

Source: Jung, C. et al., High Sensitivity Measurement of Transcription Factor-DNA Binding Affinities by Competitive Titration Using Fluorescence ...

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Fluorescence Anisotropy-Based Detection of Protein-Protein Interactions

Source: Gijsbers, A., et al. <a href="https://app.jove.com/v/54640">Fluorescence Anisotropy as a Tool to Study Protein-protein Interactions.</a> J. Vis. Exp. (2016).
In this video, we describe the fluorescence anisotropy technique to study the interactions between the fluorophore-tagged Shwachman-diamond syndrome (SBDS) protein and the elongation factor-like 1 GTPase (EFL1). On incubating SBDS proteins with gradually increasing concentrations of EFL1, a steady increase in anisotropy is observed, indicating a successful interaction between the two proteins.

Source: Gijsbers, A., et al. Fluorescence Anisotropy as a Tool to Study Protein-protein Interactions. J. Vis. Exp. (2016).
In this video, we describe the ...

Protein-protein interactions

Fluorescence Fluctuation Spectroscopy to Study Protein Homo-Oligomerization

To set up the multiwell plate array, first, prepare a solution of 100 nanomolar purified FKBP12 in the same buffer used for size exclusion chromatography. Sonicate and centrifuge with a quick spin of 13,000 RPM to prevent the formation of aggregates.
Now, pipette 100 to 200 microliters of the diluted protein into an 8-well observation chamber with a glass bottom. Add the BB dimerizer to final concentrations of 10, 20, 40, 80, 100, 150, 300, and 500 nanomolar. As a reference, prepare solution of 100 nanomolar mVenus alone to evaluate potential aggregation and precipitation effects, and to recover a brightness value for the monomer with the same acquisition settings.
Any light scanning microscope confocal system equipped with digital detectors or well-characterized analog detectors, and capable of keeping a constant dwell time for every pixel acquired can be used. Select the collar correction water immersion objective designed for fluorescence correlation spectroscopy.
Now add a drop of water to the collar correction water immersion objective. Mount the 8-well observation chamber into the stage. Set the excitation beam path by turning on the 514-nanometer laser, and setting it at 20 to 100 nanowatts power at the exit of the objective. Turn on one HyD detector. Detectors capable of photon-counting are preferable. Select the emission window from 520 to 560 nanometers.
For the acquisition mode, use 16 by 16 pixels.
Set the pixel dwell time such that the frame time is longer than the protein diffusion, and the pixel dwell time is much shorter. This corresponded to setting the dwell time to approximately 13 microseconds for the system used in this demonstration.
Set the pinhole at one Airy unit for the corresponding emission of approximately 545 nanometers. Select the xy-time acquisition mode and select the number of frames to be acquired per acquisition and well. Now, set the pixel size at approximately 120 nanometers.
If the system is equipped with high-throughput mode, introduce the coordinates of each well, and the number of acquisitions per well to automate the process. If the system is equipped with a perfusion system, load the BB solution, and program the perfusion to start right after the 5,000th frame to evaluate the kinetics of dimerization while acquiring 10,000 images.
Select the correct well, and focus on the solution. Then, start the acquisition and save the resulting stack of images in TIFF format.

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Fluorescence Fluctuation Spectroscopy to Study Protein Interaction at Cell Contacts

Transfer the cell solution of one well to the corresponding well. Mix gently by pipetting a few times up and down, then, seed the mixed cells on a 35-milliliter glass-bottomed dish, and culture the seeded cells at 37 degrees Celsius and 5% carbon dioxide for one day.
In the laser-scanning confocal microscope software, set up the optical path. To avoid spectral cross-talk, select two separate tracks to excite and detect mEGFP or mEYFP and mCherry or mCardinal sequentially, and select &#34;Switch Tracks Every Line.&#34; For the detection, use appropriate filters for both channels.
Place the dish containing the mixed cells on the sample holder. After waiting 10 minutes to ensure temperature equilibration and to reduce focus drift, focus on the cells using the Transmission Light in the &#34;Locate&#34; menu. Search for a pair of &#39;red&#39; and &#39;green&#39; cells in contact with each other.
Next, select a scan path perpendicular to cell-cell contact using the &#34;Crop&#34; button. &#34;Zoom&#34; to achieve a pixel size of 50 to 200 nanometers and select &#34;Line&#34; in &#34;Scan Mode.&#34; Set &#34;Frame Size&#34; to 128 by 1 pixels. Set &#34;Scan Speed&#34; to the maximum allowed value, then set cycles to 100,000 to 500,000.
Following this, choose the appropriate laser powers. Set the detectors to &#34;Photon Counting&#34; Mode. Press &#34;Start Experiment&#34; to start the acquisition.

Fluorescence Anisotropy as a Tool to Study Protein-protein Interactions

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