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 &#181;g/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 &#176;C until absorbance at 600 nm (A600) reaches 0.5-0.7 in 1 L of LB liquid media supplemented with 100 &#181;g/ml ampicillin.</li>
		<li>Induce protein expression by adding 0.5 mM isopropyl &#946;-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 &#176;C. Remove the supernatant. At this point, either store the cell pellet at -20 &#176;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 &#176;C.</li>
		<li>Centrifuge the sample at 9,000 x g for 50 min at 4 &#176;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 &#181;m 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 &#176;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&#39;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 &#176;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 &#176;C.</li>
		<li>Collect the yeast suspension by centrifugation at 3,800 x g for 10 min at 4 &#176;C. Remove the supernatant. At this point, either store the cell pellet at -20 &#176;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 (&#216; = 0.5 mm) for a total time of 6 min using cycles of 2 min ON and 15 min OFF, at 4 &#176;C.</li>
		<li>Centrifuge the sample at 9,000 x g for 50 min at 4 &#176;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 &#946;-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 &#181;M. Flash freeze the protein in liquid nitrogen and store at -80 &#176;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&#39;,5&#39;-Bis(1,3,2 dithioarsolan-2-yl) Fluorescein
<ol>
	<li>Mix 3 nmol of the SBDS-FlAsH protein with 3 nmol of the 4&#39;,5&#39;-bis(1,3,2 dithioarsolan-2-yl) fluorescein dye in 5 &#181;l volume of Anisotropy buffer.</li>
	<li>Let the reaction proceed for 8 hr at 4 &#176;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="Equation1B" src="/files/ftp_upload/54640/54640eq1B.jpg" /></li>
	<li>Calculate the concentration of labeled SBDS-FlAsH protein using Equation 2. 
	<img alt="Equation2" src="/files/ftp_upload/54640/54640eq2.jpg" /></li>
	<li>Calculate the concentration of total SBDS protein using Equation 3 by substituting the calculated CSBDS-FlAsH from previous step. 
	<img alt="Equation3" src="/files/ftp_upload/54640/54640eq3.jpg" /></li>
	<li>Calculate the percentage of labeled protein using Equation 4. 
	<img alt="Equation4" src="/files/ftp_upload/54640/54640eq4.jpg" /></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&#177;25 nm. Measurements were done at 25 &#176;C in a 200 &#181;l cuvette with a 5 mm path length 10.
<ol>
	<li>In a fluorescence cuvette, place 200 &#181;l of 30 nM SBDS-FlAsH in anisotropy buffer and titrate 2 &#181;l of 30 &#181;M 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 &#181;l 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="https://cloudfront-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54640/54640table1large.jpg" target="_blank">Please click here to view a larger version of this table.</a> 
<img alt="Table 1" src="/files/ftp_upload/54640/54640table1.jpg" />

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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 analysis since they are common to most proteins and would be present in both, the receptor protein and the ligand. Additionally, the lifetime of tryptophan is too short for fluorescence anisotropy experiments. For this reason, it is common to add extrinsic fluorophores to one of the studied proteins. To this end, a fluorophore was added to the C-terminus of SBDS through a coordination bond between the dye 4&#39;,5&#39;-bis(1,3,2 dithioarsolan-2-yl) fluorescein with a tetracysteine motif introduced in the protein by genetic engineering. Choosing the protein that bares the fluorophore is not a random matter. Due to the size of the proteins studied here (SBDS 29 kDa and EFL1 127 kDa), we expected a larger change in anisotropy between free labeled-SBDS and that in complex with EFL1, compared to the change expected between free labeled-EFL1 and that bound to SBDS. Bearing this in mind, a FlAsH tag was introduced into SBDS rather than to EFL1. Another parameter important to consider when setting up a fluorescence anisotropy experiment is the absorption of the solutions used. The optical density of the solution in the cuvette should not be greater than 0.1 at the excitation and emission wavelength, and should remain constant along the entire titration. Otherwise, inner filter effect can interfere in the measurement and must be corrected for. In the experiment presented here, the proteins studied do not absorb at the wavelength of excitation or emission of the 4&#39;,5&#39;-bis(1,3,2 dithioarsolan-2-yl) fluorescein dye; thus inner filter effect does not pose a problem.
Regarding data collection, and to subsequently properly fit the data, it is necessary to have well-defined lower and upper baselines with several data points through the transition along the binding curve. Otherwise, fitting of the data is inaccurate and information regarding the binding mode may be lost. To describe a decent binding curve it is necessary that the concentration of free ligand differs by two logarithmic units below and above the dissociation constant, however, experimentally this may be difficult to achieve. In the lower limit, problems with the sensitivity of the equipment may arise particularly when the affinity between the proteins is very high, while large concentrations may not be attainable due to solubility problems of the ligand. This was clearly exemplified when testing the interaction between the SBDS S143L disease mutant and EFL1. This mutant disrupted the binding to EFL1 to such an extent that it was not possible to obtain a proper binding curve since EFL1 can only be concentrated up to 70 &#181;M and the protein gets diluted &#8776; 5 fold during the titration 10. Thus, it is necessary to have a rough estimate of the affinity to optimize the titration scheme and make sure that jumps in ligand concentration do not occur at positions that would cause a less accurate fit. For example, missing the initial points of the binding curve presented in Figure 3 will make it look like a hyperbola misleading the researcher towards a single binding site model.
Once collected, data were fitted to a presumed binding model. In comparison to fluorescence data, anisotropy data can be used without further manipulation and do not need correction for dilution effects. The equations presented in Table 1 consider that [L] &#8776; [L]0 at every point of the titration. This may not happen when the affinity between the proteins is very high such that in the first points of the titration ligand depletion occurs. In this scenario, more complex quadratic equations described elsewhere 23 are needed to fit the data. In the experiment described here, the ligand EFL1 was always in large excess compared to the concentration of SBDS-FlAsH and the change in EFL1 concentration at each point of the titration can be disregarded. As discussed in the results section, a two non-identical binding sites model best described the experimental data (Figure 3C). Additional biochemical information obtained in the group supports the idea that this is the correct model to describe the interaction between EFL1 and SBDS (data not shown). This mode of interaction is common in multi-domain proteins, such as EFL1 and SBDS are, and several examples exist in the literature 24-26. However, it is also important to rule out a non-specific interaction that can appear as an interaction model of distinct binding sites with different affinities for their ligand. To this end, a Scatchard plot is very useful. Compared to other techniques, fluorescence anisotropy signal reports the amount of complex formed and thus it is possible to build a Scatchard plot with the data. A convex Scatchard curve suggests a two different binding sites model with negative cooperativity or non-specific binding, while a concave Scatchard plot implies a two different binding sites model with positive cooperativity or ligand instability (Figure 4). Analysis of the experimental data showed a Scatchard plot with a concave shape supporting the model of two independent binding sites for EFL1 and SBDS (Figure 4D). Furthermore, positive cooperativity was confirmed after performing a Hill analysis that resulted in a Hill coefficient value of 1.8&#177;0.02 (data not shown).
<img alt="Figure 4" src="/files/ftp_upload/54640/54640fig4.jpg" /> 
Figure 4. Scatchard plots for different protein-protein binding models. (A) Single binding site or multiple identical sites that do not interact. (B) Multiple different binding sites with negative cooperativity or non-specific binding. (C) Multiple different binding sites with positive cooperativity or ligand instability. (D) Scatchard plot resulting from analysis of the experimental anisotropy data obtained from the titration of EFL1 to SBDS-FlAsH. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54640/54640fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Fluorescence anisotropy is a solution-based, true-equilibrium technique that requires the studied molecules to remain in solution at the conditions tested. It can be used to study not only the interaction between full-length proteins but also the interaction between protein domains, mutant proteins or even peptides with post-translational modifications. Moreover, fluorescence anisotropy can also be used to measure the binding of proteins to lipid membranes using fluorescent sensitive membranes probes. This approach has been successfully used to study the effect that &#945;-synuclein has on synaptic vesicle packing 27. One of the main advantages of fluorescence anisotropy is that it provides quantitative information on the binding of the studied molecules such that true dissociation constants can be obtained together with mechanistic information. Although other biophysical techniques can also be used to obtain such information, fluorescence anisotropy requires small amounts of sample and can be performed not only in equilibrium, as presented here, but also using rapid kinetics, such as stopped-flow fluorometry, to obtain the association and dissociation rate constants (kon and koff, respectively) 28. A comparative table with the benefits and disadvantages of fluorescence anisotropy with respect to other biophysical techniques is presented in Table 2. All the methods described are true-equilibrium techniques since none of them include a process of mechanical separation of the free and bound fractions. As mentioned in the results section, if the fluorescence intensity of a labeled-protein largely changes upon interaction, this property can then be used to quantify the affinity of the binding. However, changes in fluorescence intensity resulting from the interaction suggest a possible interference of the extrinsic fluorophore on the molecular interaction itself; namely a possible alteration of the dissociation constant (Kd) value of the corresponding non-labeled proteins. As such, fluorescence anisotropy can be considered a less invasive technique than changes in fluorescence intensity since the former should be applied when fluorescence intensity does not change upon interaction. In this sense, though large amounts of the protein are required, Isothermal Titration Calorimetry (ITC) can provide a true Kd value of the molecular interaction since it is a label free method. On the other hand, mechanistic information is not obtained because differences in heat only report on the enthalpy of the process and not the amount of complex formed 29.
In comparison, in vivo methods such as yeast two-hybrid or fragment complementation assays do not require purified proteins but they only provide semi-quantitative information on the mode of binding. Despite the fact that in the yeast two-hybrid technique is possible to express the strength of the binding using Miller units, this is not a real equilibrium constant and many factors out of the control of the researcher can alter it.
Table 2. Advantages and disadvantages of commonly used biophysical techniques used to study protein-protein interactions. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54640/54640table2large.jpg" target="_blank">Please click here to view a larger version of this table.</a> 
<img alt="Table 2" src="/files/ftp_upload/54640/54640table2.jpg" />

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="Equation1" src="/files/ftp_upload/54640/54640eq1.jpg" />
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="Figure 1" src="/files/ftp_upload/54640/54640fig1.jpg" /> 
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="https://cloudflare-jove-com.remotexs.ntu.edu.sg/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&#39;,5&#39;-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.

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</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="Equation5" src="/files/ftp_upload/54640/54640eq5.jpg" />
where Fi represents the fractional fluorescence of each component and ri is the corresponding anisotropy. The fractional fluorescence of each component depends on both, its concentration and its relative fluorescence, which is defined by Equation 6:
<img alt="Equation6" src="/files/ftp_upload/54640/54640eq6.jpg" />
where Xi represents the mole fraction of the ith specie and &#216;i is the quantum yield of the ith specie.
Thus, if fluorescence intensity changes dramatically along the titration it is better to, either measure complex formation as a function of the change in fluorescence signal and not of the anisotropy signal, or correct the observed anisotropy value by fitting both the fluorescence intensity and the anisotropy data. The fluorescence of SBDS-FlAsH does not noticeable change upon binding to EFL1 (Figure 2) suggesting that fluorescence anisotropy is adequate for measuring the binding between the two proteins.
<img alt="Figure 2" src="/files/ftp_upload/54640/54640fig2.jpg" /> 
Figure 2. Fluorescence emission of SBDS-FlAsH upon titration of increasing concentrations of EFL1. Changes in the fluorescence emission of the fluorophore are negligible and random along the titration. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54640/54640fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
An example of the fluorescence anisotropy binding curve obtained from the titration of EFL1 to SBDS-FlAsH is shown in Figure 3. Qualitatively, the shape of the graph clearly shows that the two proteins interact as evidenced by an increase in the anisotropy signal upon addition of EFL1. Furthermore, anisotropy values reached a plateau suggesting that at the end of the titration all the SBDS-FlAsH protein was present in a complex with EFL1. It is possible then, to quantitatively describe the interaction between these proteins and fit the data using nonlinear least squares regression to a presumed binding model and obtain the corresponding dissociation equilibrium constant. Fitting to a one binding site model did not appropriately describe the experimental data (Figure 3A). This is evident not only from the fitting trace in the binding curve but also from the deviations of the residuals of the fit (the difference between the theoretical and experimental data) that are appreciable and nonrandom. This agrees well with the fact that a single binding site model is described mathematically by a hyperbolic curve rather than a sigmoidal curve as that observed for the experimental data presented in Figure 3. On the other hand, models such as two identical or two different binding sites describe better the experimental data and the residuals of the fit show no systematic deviation supporting a two-site association model (Figure 3B-C). In the absence of any other experimental information is difficult to establish which of the two models correspond to the binding mechanism between EFL1 and SBDS. Nevertheless, SBDS and EFL1 are both monomeric proteins, so it is difficult to envisage a two identical binding sites model.
<img alt="Figure 3" src="/files/ftp_upload/54640/54640fig3.jpg" /> 
Figure 3. Fluorescence anisotropy binding isotherm of the interaction between EFL1 and SBDS-FlAsH. The continuous line in each of the upper plots corresponds to the fit of the data to different binding models: (A) single site, (B) two identical sites and (C) two non-identical sites. Lower plots represent the residuals of the fit to the corresponding model. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54640/54640fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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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30.3K Views.  Universidad Nacional Autónoma de México. 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).

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 ...

Biotinylated Cell-penetrating Peptides to Study Intracellular Protein-protein Interactions

Here we present a protocol to study intracellular protein-protein interactions that is based on the widely used biotin-avidin pull-down system. The modification presented includes the combination of this technique with cell-penetrating sequences. We propose to design cell-penetrating baits that can be incubated with living cells instead of cell lysates and therefore the interactions found will reflect those that occur within the intracellular context. Connexin43 (Cx43), a protein that forms gap junction channels and hemichannels is down-regulated in high-grade gliomas. The Cx43 region comprising amino acids 266-283 is responsible for the inhibition of the oncogenic activity of c-Src in glioma cells. Here we use TAT as the cell-penetrating sequence, biotin as the pull-down tag and the region of Cx43 comprised between amino acids 266-283 as the target to find intracellular interactions in the hard-to-transfect human glioma stem cells. One of the limitations of the proposed method is that the molecule used as bait could fail to fold properly and, consequently, the interactions found could not be associated with the effect. However, this method can be especially interesting for the interactions involved in signal transduction pathways because they are usually carried out by intrinsically disordered regions and, therefore, they do not require an ordered folding. In addition, one of the advantages of the proposed method is that the relevance of each residue on the interaction can be easily studied. This is a modular system; therefore, other cell-penetrating sequences, other tags, and other intracellular targets can be employed. Finally, the scope of this protocol is far beyond protein-protein interaction because this system can be applied to other bioactive cargoes such as RNA sequences, nanoparticles, viruses or any molecule that can be transduced with cell-penetrating sequences and fused to pull-down tags to study their intracellular mechanism of action.

This is a protocol to study intracellular protein-protein interactions based on the biotin-avidin pull-down system with the novelty of combining cell-penetrating sequences. The main advantage is that the target sequence is incubated with living cells instead of cell lysates and therefore the interactions will occur within the cellular context.

Here we present a protocol to study intracellular protein-protein interactions that is based on the widely used biotin-avidin pull-down system. The ...

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 ...

A Yeast 2-Hybrid Screen in Batch to Compare Protein Interactions

Screening for protein-protein interactions using the yeast 2-hybrid assay has long been an effective tool, but its use has largely been limited to the discovery of high-affinity interactors that are highly enriched in the library of interacting candidates. In a traditional format, the yeast 2-hybrid assay can yield too many colonies to analyze when conducted at low stringency where low affinity interactors might be found. Moreover, without a comprehensive and complete interrogation of the same library against different bait plasmids, a comparative analysis cannot be achieved. Although some of these problems can be addressed using arrayed prey libraries, the cost and infrastructure required to operate such screens can be prohibitive. As an alternative, we have adapted the yeast 2-hybrid assay to simultaneously uncover dozens of transient and static protein interactions within a single screen utilizing a strategy termed DEEPN (Dynamic Enrichment for Evaluation of Protein Networks), which incorporates high-throughput DNA sequencing and computation to follow the evolution of a population of plasmids that encode interacting partners. Here, we describe customized reagents and protocols that allow a DEEPN screen to be executed easily and cost-effectively.

Batch processing of yeast 2-hybrid screens allows for direct comparison of the interaction profiles of multiple bait proteins with a highly complex set of prey fusion proteins. Here, we describe refined methods, new reagents, and how to implement their use for such screens.

Screening for protein-protein interactions using the yeast 2-hybrid assay has long been an effective tool, but its use has largely been limited to the ...

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 ...

Application of Biolayer Interferometry (BLI) for Studying Protein-Protein Interactions in Transcription

A transcription factor&#160;(TF) is a protein that regulates gene expression by interacting with the RNA polymerase, another TF, and/or template DNA. GrgA is a novel transcription activator found specifically in the obligate intracellular bacterial pathogen Chlamydia. Protein pulldown assays using affinity beads have revealed that GrgA binds two &#963; factors, namely &#963;66 and &#963;28, which recognize different sets of promoters for genes whose products are differentially required at developmental stages. We have used BLI to confirm and further characterize the interactions. BLI demonstrates several advantages over pulldown: 1) It reveals real-time association and dissociation between binding partners, 2) It generates quantitative kinetic parameters, and 3) It can detect bindings that pulldown assays often fail to detect. These characteristics have enabled us to deduce the physiological roles of GrgA in gene expression regulation in Chlamydia, and possible detailed interaction mechanisms. We envision that this relatively affordable technology can be extremely useful for studying transcription and other biological processes.

Application of Biolayer Interferometry BLI for Studying Protein-Protein Interactions in Transcription

Interactions of transcription factors (TFs) with the RNA polymerase are usually studied using pulldown assays. We apply a Biolayer Interferometry (BLI) technology to characterize the interaction of GrgA with the chlamydial RNA polymerase. Compared to pulldown assays, BLI detects real-time association and dissociation, offers higher sensitivity, and is highly quantitative.

A transcription factor&#160;(TF) is a protein that regulates gene expression by interacting with the RNA polymerase, another TF, and/or template DNA. GrgA ...

Evaluation of Protein&#8211;Protein Interactions using an On-Membrane Digestion Technique

Numerous intracellular proteins physically interact in accordance with their intracellular and extracellular circumstances. Indeed, cellular functions largely depend on intracellular protein&#8211;protein interactions. Therefore, research regarding these interactions is indispensable to facilitating the understanding of physiologic processes. Co-precipitation of associated proteins, followed by mass spectrometry (MS) analysis, enables the identification of novel protein interactions. In this study, we have provided details of the novel technique of immunoprecipitation-liquid chromatography (LC)-MS/MS analysis combined with on-membrane digestion for the analysis of protein&#8211;protein interactions. This technique is suitable for crude immunoprecipitants and can improve the throughput of proteomic analyses. Tagged recombinant proteins were precipitated using specific antibodies; next, immunoprecipitants blotted onto polyvinylidene difluoride membrane pieces were subjected to reductive alkylation. Following trypsinization, the digested protein residues were analyzed using LC-MS/MS. Using this technique, we were able to identify several candidate associated proteins. Thus, this method is convenient and useful for the characterization of novel protein&#8211;protein interactions.

Here, we present a protocol for an on-membrane digestion technique for the preparation of samples for mass spectrometry. This technique facilitates the convenient analysis of protein&#8211;protein interactions.

Numerous intracellular proteins physically interact in accordance with their intracellular and extracellular circumstances. Indeed, cellular functions ...

In Vivo Hydroxyl Radical Protein Footprinting for the Study of Protein Interactions in Caenorhabditis elegans

Fast oxidation of proteins (FPOP) is a hydroxyl radical protein footprinting (HRPF) method used to study protein structure, protein-ligand interactions, and protein-protein interactions. FPOP utilizes a KrF excimer laser at 248 nm for photolysis of hydrogen peroxide to generate hydroxyl radicals which in turn oxidatively modify solvent-accessible amino acid side chains. Recently, we expanded the use of FPOP of in vivo oxidative labeling in Caenorhabditis elegans (C. elegans), entitled IV-FPOP. The transparent nematodes have been used as model systems for many human diseases. Structural studies in C. elegans by IV-FPOP is feasible because of the animal&#8217;s ability to uptake hydrogen peroxide, their transparency to laser irradiation at 248 nm, and the irreversible nature of the modification. The assembly of a microfluidic flow system for IV-FPOP labeling, IV-FPOP parameters, protein extraction, and LC-MS/MS optimized parameters are described herein.

In Vivo Hydroxyl Radical Protein Footprinting for the Study of Protein Interactions in Caenorhabditis elegans

In vivo fast photochemical oxidation of proteins (IV-FPOP) is a hydroxyl radical protein footprinting technique that allows for mapping of protein structure in their native environment. This protocol describes the assembly and set-up of the IV-FPOP microfluidic flow system.

Fast oxidation of proteins (FPOP) is a hydroxyl radical protein footprinting (HRPF) method used to study protein structure, protein-ligand interactions, ...

Optical Tweezers to Study RNA-Protein Interactions in Translation Regulation

RNA adopts diverse structural folds, which are essential for its functions and thereby can impact diverse processes in the cell. In addition, the structure and function of an RNA can be modulated by various trans-acting factors, such as proteins, metabolites or other RNAs. Frameshifting RNA molecules, for instance, are regulatory RNAs located in coding regions, which direct translating ribosomes into an alternative open reading frame, and thereby act as gene switches. They may also adopt different folds after binding to proteins or other trans-factors. To dissect the role of RNA-binding proteins in translation and how they modulate RNA structure and stability, it is crucial to study the interplay and mechanical features of these RNA-protein complexes simultaneously.&#160;This work illustrates how to employ single-molecule-fluorescence-coupled optical tweezers to explore the conformational and thermodynamic landscape of RNA-protein complexes at a high resolution. As an example, the interaction of the SARS-CoV-2 programmed ribosomal frameshifting element with the trans-acting factor short isoform of zinc-finger antiviral protein is elaborated. In addition, fluorescence-labeled ribosomes were monitored using the confocal unit, which would ultimately enable the study of translation elongation. The fluorescence coupled OT assay can be widely applied to explore diverse RNA-protein complexes or trans-acting factors regulating translation and could facilitate studies of RNA-based gene regulation.

This protocol presents a complete experimental workflow for studying RNA-protein interactions using optical tweezers. Several possible experimental setups are outlined including the combination of optical tweezers with confocal microscopy.