This work was supported by the National Institutes of Health Grant HL127711 to AVK.

<ol>
	<li>Pollard, T. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Actin+and+Actin-Binding+Proteins.">Actin and Actin-Binding Proteins.</a> Cold Spring Harb Perspect Biol. 8 (8), (2016).</li><li>Hansen, M. D., Kwiatkowski, A. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Control+of+actin+dynamics+by+allosteric+regulation+of+actin+binding+proteins.">Control of actin dynamics by allosteric regulation of actin binding proteins.</a> Int Rev Cell Mol Biol. 303, 1-25 (2013).</li><li>Lappalainen, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Actin-binding+proteins:+the+long+road+to+understanding+the+dynamic+landscape+of+cellular+actin+networks.">Actin-binding proteins: the long road to understanding the dynamic landscape of cellular actin networks.</a> Mol Biol Cell. 27 (16), 2519-2522 (2016).</li><li>Mullins, R. D., Hansen, S. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+studies+of+actin+filament+and+network+dynamics.">In vitro studies of actin filament and network dynamics.</a> Curr Opin Cell Biol. 25 (1), 6-13 (2013).</li><li>Miller, P. W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Danio+rerio+alphaE-catenin+is+a+monomeric+F-actin+binding+protein+with+distinct+properties+from+Mus+musculus+alphaE-catenin.">Danio rerio alphaE-catenin is a monomeric F-actin binding protein with distinct properties from Mus musculus alphaE-catenin.</a> J Biol Chem. 288 (31), 22324-22332 (2013).</li><li>Wickline, E. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=alphaT-Catenin+Is+a+Constitutive+Actin-binding+alpha-Catenin+That+Directly+Couples+the+Cadherin.Catenin+Complex+to+Actin+Filaments.">alphaT-Catenin Is a Constitutive Actin-binding alpha-Catenin That Directly Couples the Cadherin.Catenin Complex to Actin Filaments.</a> J Biol Chem. 291 (30), 15687-15699 (2016).</li><li>Simpson, R. J., Adams, P. D., Golemis, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Basic methods in protein purification and analysis : a laboratory manual. , (2009).</li><li>Laemmli, U. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cleavage+of+structural+proteins+during+the+assembly+of+the+head+of+bacteriophage+T4.">Cleavage of structural proteins during the assembly of the head of bacteriophage T4.</a> Nature. 227 (5259), 680-685 (1970).</li><li>Dyballa, N., Metzger, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fast+and+sensitive+colloidal+coomassie+G-250+staining+for+proteins+in+polyacrylamide+gels.">Fast and sensitive colloidal coomassie G-250 staining for proteins in polyacrylamide gels.</a> J Vis Exp. (30), (2009).</li><li>Pollard, T. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+guide+to+simple+and+informative+binding+assays.">A guide to simple and informative binding assays.</a> Mol Biol Cell. 21 (23), 4061-4067 (2010).</li><li>Hansen, S. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=alphaE-catenin+actin-binding+domain+alters+actin+filament+conformation+and+regulates+binding+of+nucleation+and+disassembly+factors.">alphaE-catenin actin-binding domain alters actin filament conformation and regulates binding of nucleation and disassembly factors.</a> Mol Biol Cell. 24 (23), 3710-3720 (2013).</li><li>Perrin, B. J., Ervasti, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+actin+gene+family:+function+follows+isoform.">The actin gene family: function follows isoform.</a> Cytoskeleton (Hoboken). 67 (10), 630-634 (2010).</li><li>Tondeleir, D., Vandamme, D., Vandekerckhove, J., Ampe, C., Lambrechts, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Actin+isoform+expression+patterns+during+mammalian+development+and+in+pathology:+insights+from+mouse+models.">Actin isoform expression patterns during mammalian development and in pathology: insights from mouse models.</a> Cell Motil Cytoskeleton. 66 (10), 798-815 (2009).</li><li>Prochniewicz, E., Janson, N., Thomas, D. D., Dela Cruz, E. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cofilin+increases+the+torsional+flexibility+and+dynamics+of+actin+filaments.">Cofilin increases the torsional flexibility and dynamics of actin filaments.</a> J Mol Biol. 353 (5), 990-1000 (2005).</li><li>Kuhn, J. R., Pollard, T. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Real-time+measurements+of+actin+filament+polymerization+by+total+internal+reflection+fluorescence+microscopy.">Real-time measurements of actin filament polymerization by total internal reflection fluorescence microscopy.</a> Biophys J. 88 (2), 1387-1402 (2005).</li><li>Hansen, S. D., Zuchero, J. B., Mullins, R. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cytoplasmic+actin:+purification+and+single+molecule+assembly+assays.">Cytoplasmic actin: purification and single molecule assembly assays.</a> Methods Mol Biol. 1046, 145-170 (2013).</li></ol>

The authors declare that they have no competing financial interests.

The actin co-sedimentation assay is a straightforward technique that can quickly determine if a protein binds F-actin. With some modifications, the technique can also be used to measure the affinity of the interaction. In addition to points raised in the protocol above, the following issues should be considered when designing, conducting, and interpreting the assay.
Protein of Interest
Freshly prepared or frozen protein can be used in the assay. If frozen protein is used, it is highly recommended that the results be compared with fresh (never frozen) protein to ensure that freezing does not affect F-actin binding.
G-actin Source
Many pelleting experiments use G-actin isolated from muscle because of its relative abundance. There are three main actin isotypes in mammals &#8211; alpha, beta and gamma &#8211; that are remarkably similar (&#62;90% sequence identity). Nonetheless, there are functional differences between the isotypes12,13. If possible, the G-actin isotype used in the binding assay should match the in vivo isotype. For example, if testing a protein expressed in skeletal muscle, alpha-actin is the best choice; if examining a protein expressed in fibroblasts, beta-actin is recommended.
Phalloidin Use
Since phalloidin binds F-actin, it can interfere or even block the binding of some F-actin binding proteins (e.g., phalloidin blocks cofilin from binding to actin filaments)14. Thus, phalloidin should be used with caution and the results compared to non-phalloidin-treated samples when possible.
High Background
It is not uncommon for proteins to sediment in the absence of F-actin (Figure 1A, no F-actin pellet samples). However, high levels of background sedimentation can mask true actin co-sedimentation and make it difficult, if not impossible, to determine if a protein binds F-actin or to measure the affinity of the interaction. Adding polidicanol to the reaction buffer (step 4.1) can significantly reduce the background and is an easy solution. If that does not reduce the background, adjusting the reaction buffer, salt concentration, and/or incubation temperature may help.
Binding Curve
To generate a binding curve, it is necessary to vary the concentration of either the protein of interest or the F-actin over a series of reactions. In practice, it is easier and preferable to maintain F-actin at a fixed concentration and to vary the concentration of the protein of interest. Maintaining F-actin at a fixed concentration (e.g., 2 &#181;M) in the pelleting assay limits non-specific trapping at higher concentrations of F-actin and prevents depolymerization at lower (&#60;0.5 &#181;M) concentrations of F-actin. Depolymerization can be prevented using phalloidin, although this introduces a potential complicating factor into the system (see step 3.3 and above). Maintaining F-actin at a fixed concentration also allows one to compare (and normalize) the F-actin pellet across samples and to identify failed experiments (i.e., where the F-actin pellet is highly variable, preventing analysis across concentrations). Finally, maintaining F-actin at a fixed concentration allows one to determine if the binding to the actin filament is cooperative (Figure 1C).
Saturated Binding
As in all binding experiments, it is critical that the binding to F-actin is saturated and that the concentration of protein plus F-actin plateaus (Figure 1C). Without a plateau, it is not possible to calculate an accurate dissociation equilibrium constant. Thus, it is important to carefully plan the dilution series to be tested and to always include higher concentrations of protein (i.e., at least 5- to 10-fold higher than the expected Kd).
Binding Analysis
In order for the measured dissociation constants to be conclusive, the assay should be performed using an F-actin concentration that allows the concentration of binding sites on F-actin for the protein of interest to be much lower than the affinity. To check whether this criterion was met, estimate the concentration of binding sites from Bmax. For example, if [F-actin] was 2 &#181;M and Bmax = 0.5, then [binding sites] &#8776; 1 &#181;M. The Kd should be at least 5- to 10-fold greater than [binding sites]. If the measured Kd is of the same order of magnitude as [binding sites], then it is possible that the observed binding curve represents a titration of high-affinity binding sites rather than a true binding isotherm. If this is observed, repeat the assay using a 10-fold lower F-actin concentration to measure an accurate affinity. For high-affinity interactions, phalloidin stabilization (step 3.3) may be necessary to achieve an F-actin concentration low enough to accurately measure affinity.
Finally, there are fundamental limitations with the co-sedimentation assay that researchers should be aware of when performing and evaluating the assay. Most importantly, the co-sedimentation assay does not produce a true equilibrium constant. The products of binding (i.e., protein plus F-actin) are separated from the reactants during centrifugation, whereupon the products can then dissociate to create a new equilibrium. As a result, the co-sedimentation assay can miscalculate or fail to detect low-affinity interactions. Since many actin-binding proteins have a low (i.e., micromolar) affinity for F-actin, a negative result (i.e., no detectable binding) in the assay does not necessarily mean that a protein does not bind F-actin. As an alternative, TIRF microscopy-based, single-filament binding assays are more sensitive and more accurate for determining a dissociation constant (for reviews on this technique, see references15,16). Despite these limitations, the pelleting assay is within the means of most researchers and is an effective tool to determine if a protein binds F-actin and to measure the affinity of the interaction.

Actin is an essential cytoskeletal protein that plays a critical role in multiple cellular processes, including motility, contraction, adhesion, and morphology1. Actin exists in two forms: monomeric globular actin (G-actin) and polymerized filamentous actin (F-actin). Within cells, F-actin organization is controlled by a large collection of proteins that regulate the nucleation, growth, cross-linking, and disassembly of actin filaments2,3,4. However, how multiple actin-binding proteins function in concert to regulate actin network organization is still largely unclear.
The measurement of protein-protein interactions is an important approach for understanding how proteins exert their effects on cellular behavior at the biochemical level. Many different assays can be used to detect interactions between purified proteins. Common approaches for soluble proteins include pull-downs, fluorescence polarization, isothermal titration calorimetry, and surface plasmon resonance. Importantly, all of these assays require proteins to be soluble, and are thus difficult to adapt for use with a polymeric, filamentous protein such as F-actin. Here, we describe a technique &#8211; the actin co-sedimentation, or pelleting, assay &#8211; to determine if a protein or protein domain binds F-actin and to measure the affinity of the interaction.
The actin pelleting assay is a relatively straightforward technique that does not require specialized equipment, aside from an ultracentrifuge. All reagents can be made, assuming knowledge of basic biochemistry, or purchased. Once binding to F-actin is established, the assay can be used to measure the apparent affinity (i.e., the dissociation equilibrium constant)5. Also, once an affinity is established, the pelleting assay is a useful tool to measure how changes to the protein of interest (i.e., post-translational modifications, mutations, or ligand binding) affect its interaction with F-actin6. The technique does have limitations (see the Discussion) that the researcher should be aware of before attempting the assay.

<table><tbody><tr><td>Sorvall MTX 150 Micro-Ultracentrifuge</td><td>ThermoFisher Scientific</td><td>46960</td><td></td></tr><tr><td>S100-AT3 rotor</td><td>ThermoFisher Scientific</td><td>45585</td><td></td></tr><tr><td>Ultracentrifuge tubes - 0.2 mL</td><td>ThermoFisher Scientific</td><td>45233</td><td></td></tr><tr><td>Actin, rabbit skeletal muscle</td><td>Cytoskeleton</td><td>AKL99</td><td></td></tr><tr><td>Bovine Serum Albumin</td><td>Sigma</td><td>A8531</td><td></td></tr><tr><td>Polidicanol (Thesit)&amp;nbsp;</td><td>Sigma</td><td>88315</td><td></td></tr><tr><td>Phalloidin</td><td>ThermoFisher Scientific</td><td>P3457</td><td></td></tr><tr><td>Dithiothreitol (DTT)</td><td>ThermoFisher Scientific</td><td>R0862</td><td></td></tr><tr><td>Adenosine triphosphate (ATP)</td><td>Sigma</td><td>A2383</td><td></td></tr><tr><td>Imidazole</td><td>Fisher Scientific</td><td>O3196</td><td></td></tr><tr><td>Sodium Chloride (NaCl)</td><td>Fisher Scientific</td><td>BP358</td><td></td></tr><tr><td>Magnesium Chloride (MgCl&lt;sub&gt;2&lt;/sub&gt;)</td><td>Fisher Scientific</td><td>M33</td><td></td></tr><tr><td>Potassium Chloride (KCl)</td><td>Fisher Scientific</td><td>P217</td><td></td></tr><tr><td>Ethylene glycol-bis(&amp;beta;-aminoethyl ether)-N,N,N&#039;,N&#039;-tetraacetic acid (EGTA)</td><td>Sigma</td><td>3779</td><td></td></tr><tr><td>Odyssey CLx Imaging System</td><td>LI-COR</td><td></td><td></td></tr><tr><td>Coomassie Brilliant Blue R-250 Dye</td><td>ThermoFisher Scientific</td><td>20278</td><td></td></tr><tr><td>Colloidal Blue Staining Kit</td><td>ThermoFisher Scientific</td><td>LC6025</td><td></td></tr></tbody></table>

measuring protein binding to f-actin by co-sedimentation

Filamentous actin (F-actin) organization within cells is regulated by a large number of actin-binding proteins that control actin nucleation, growth, cross-linking and/or disassembly. This protocol describes a technique &#8211; the actin co-sedimentation, or pelleting, assay &#8211; to determine whether a protein or protein domain binds F-actin and to measure the affinity of the interaction (i.e., the dissociation equilibrium constant). In this technique, a protein of interest is first incubated with F-actin in solution. Then, differential centrifugation is used to sediment the actin filaments, and the pelleted material is analyzed by SDS-PAGE. If the protein of interest binds F-actin, it will co-sediment with the actin filaments. The products of the binding reaction (i.e., F-actin and the protein of interest) can be quantified to determine the affinity of the interaction. The actin pelleting assay is a straightforward technique for determining if a protein of interest binds F-actin and for assessing how changes to that protein, such as ligand binding, affect its interaction with F-actin.

We examined &#945;E-catenin homodimer binding to F-actin in the co-sedimentation assay. Since past experiments have shown that the affinity of &#945;E-catenin homodimer for F-actin is around 1 &#181;M and the Bmax near 111, we performed the assay with a low concentration of F-actin (0.2 &#181;M rather than 2 &#181;M; see the Discussion). Since 0.2 &#181;M is below the critical concentration, phalloidin was added to stabilize the F-actin polymerized from rabbit skeletal muscle G-actin (step 3.3). Increasing concentrations of &#945;E-catenin homodimer (0.125-12.0 &#181;M) were incubated in the presence or absence of 0.2 &#181;M F-actin. The samples were centrifuged, and the resulting pellets were analyzed (Figure 1A). As expected, the &#945;E-catenin homodimer co-sedimented with F-actin above background (Figure 1A, compare the F-actin pellet samples to the no-F-actin pellet samples). BSA was run as a negative control (Figure 1B). Bound protein was quantified and plotted over free protein to calculate the affinity of the interaction (Figure 1C). The plotted data best fit a Hill equation. The calculated Kd was 2.9 &#181;M, the Bmax was 0.2, and the Hill coefficient (h) was 4.8. Thus, &#945;E-catenin homodimer binds F-actin cooperatively with a low micromolar affinity, consistent with previous work (a Kd of 2.9 &#181;M versus ~1.0 &#181;M)11.
<img alt="Figure 1" src="/files/ftp_upload/55613/55613fig1.jpg" /> 
Figure 1: High-speed F-actin co-sedimentation assay. (A) Increasing concentrations (0.125-12.0 &#181;M) of &#945;E-catenin homodimer were incubated with (left panels) or without (right panels) 0.2 &#181;M F-actin stabilized with phalloidin. They were incubated for 30 min at RT and centrifuged. The total (7.5% of the starting material) and pelleted material (50% of the pelleted material) were separated by SDS-PAGE and stained with Coomassie dye. (B) 4 &#181;M BSA was run as a negative control. Total and pellet samples, with (+) or without (-) F-actin, were separated by SDS-PAGE and stained with Coomassie dye. (C) Bound &#945;E-catenin (&#181;M/&#181;M actin) from A was plotted against free &#945;E-catenin (&#181;M), and the data fit to a Hill equation (red line). The Kd, Bmax, and Hill coefficient (h) are listed. <a href="http://cloudflare.jove.com/files/ftp_upload/55613/55613fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/55613/55613fig2.jpg" /> 
Figure 2: Actin pelleting quantification&#160;&#8211; flow chart. This schematic outlines key steps in section 5, with examples of Total and Pellet samples (A, C-E) and the standard curve (B) used for quantification. 5.13 steps: 1) Measure the amount of protein of interest in the Total samples (A). 2) Generate a standard curve by plotting the band intensity versus protein mass (B). 3) Measure the amount of protein of interest that co-sedimented with F-actin (C). 4) Measure the amount of protein of interest that pelleted in the absence of F-actin (D). 5) Subtract D from C to determine the amount of protein bound to F-actin. 6) Measure the amount of F-actin in each pellet (E), calculate the average amount of F-actin per sample, and divide each sample by the average (the numbers below show the ratio). 7) For each sample, divide the amount of bound protein (calculated in step 5) by the F-actin pellet ratio (calculated in step 6) to adjust for differences in the pellet. 8) Use the standard curve (B) to calculate the amount (mass) of normalized bound protein in each sample (step 7). 9) Determine the concentration of free protein and bound protein to create a binding curve. <a href="http://cloudflare.jove.com/files/ftp_upload/55613/55613fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Measuring Protein Binding to F-actin by Co-sedimentation at JoVE.com

Measuring Protein Binding to F-actin by Co-sedimentation

This protocol describes a method to test the ability of a protein to co-sediment with filamentous actin (F-actin) and, if binding is observed, to measure the affinity of the interaction.

Filamentous actin (F-actin) organization within cells is regulated by a large number of actin-binding proteins that control actin nucleation, growth, ...

measuring-protein-binding-to-f-actin-by-co-sedimentation

Research

JoVE Journal

Biochemistry

16.4K Views.  University of Pittsburgh School of Medicine.  Dit protocol beschrijft een methode om het vermogen van een eiwit om te co-sedimenteren met filamenteuze actine (F-actine) te testen en, als de binding wordt waargenomen, de affiniteit van de interactie te meten. 

Video: Measuring Protein Binding to F-actin by Co-sedimentation

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.

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 ...

Sedimentation Equilibrium of a Small Oligomer-forming Membrane Protein: Effect of Histidine Protonation on Pentameric Stability

Analytical ultracentrifugation (AUC) can be used to study reversible interactions between macromolecules over a wide range of interaction strengths and under physiological conditions. This makes AUC a method of choice to quantitatively assess stoichiometry and thermodynamics of homo- and hetero-association that are transient and reversible in biochemical processes. In the modality of sedimentation equilibrium (SE), a balance between diffusion and sedimentation provides a profile as a function of radial distance that depends on a specific association model. Herein, a detailed SE protocol is described to determine the size and monomer-monomer association energy of a small membrane protein oligomer using an analytical ultracentrifuge. AUC-ES is label-free, only based on physical principles, and can be used on both water soluble and membrane proteins. An example is shown of the latter, the small hydrophobic (SH) protein in the human respiratory syncytial virus (hRSV), a 65-amino acid polypeptide with a single &#945;-helical transmembrane (TM) domain that forms pentameric ion channels. NMR-based structural data shows that SH protein has two protonatable His residues in its transmembrane domain that are oriented facing the lumen of the channel. SE experiments have been designed to determine how pH affects association constant and the oligomeric size of SH protein. While the pentameric form was preserved in all cases, its association constant was reduced at low pH. These data are in agreement with a similar pH dependency observed for SH channel activity, consistent with a lumenal orientation of the two His residues in SH protein. The latter may experience electrostatic repulsion and reduced oligomer stability at low pH. In summary, this method is applicable whenever quantitative information on subtle protein-protein association changes in physiological conditions have to be measured.&#160;&#160;

Sedimentation equilibrium (SE) can be used to study protein-protein interactions in a physiological environment. This manuscript describes the use of this technique to determine the effect of pH on the stability of a homo-pentamer formed by the small hydrophobic (SH) protein encoded by the human syncytial respiratory virus (hRSV).

Analytical ultracentrifugation (AUC) can be used to study reversible interactions between macromolecules over a wide range of interaction strengths and ...

Chemistry

Actin Co-Sedimentation Assay; for the Analysis of Protein Binding to F-Actin

The actin cytoskeleton within the cell is a network of actin filaments that allows the movement of cells and cellular processes, and that generates tension and helps maintains cellular shape.  Although the actin cytoskeleton is a rigid structure, it is a dynamic structure that is constantly remodeling.  A number of proteins can bind to the actin cytoskeleton.  The binding of a particular protein to F-actin is often desired to support cell biological observations or to further understand dynamic processes due to remodeling of the actin cytoskeleton. The actin co-sedimentation assay is an in vitro assay routinely used to analyze the binding of specific proteins or protein domains with F-actin.  The basic principles of the assay involve an incubation of the protein of interest (full length or domain of) with F-actin, ultracentrifugation step to pellet F-actin and analysis of the protein co-sedimenting with F-actin.  Actin co-sedimentation assays can be designed accordingly to measure actin binding affinities and in competition assays.

Proteins bind to filamentous actin (F-actin) through distinct actin binding modules. In this video we demonstrate the procedure of actin co-sedimentation, which is an in vitro assay routinely used to analyze proteins or specific domains that bind F-actin.

The actin cytoskeleton within the cell is a network of actin filaments that allows the movement of cells and cellular processes, and that generates ...

Biology

Investigating Receptor-ligand Systems of the Cellulosome with AFM-based Single-molecule Force Spectroscopy

Cellulosomes are discrete multienzyme complexes used by a subset of anaerobic bacteria and fungi to digest lignocellulosic substrates. Assembly of the enzymes onto the noncatalytic scaffold protein is directed by interactions among a family of related receptor-ligand pairs comprising interacting cohesin and dockerin modules. The extremely strong binding between cohesin and dockerin modules results in dissociation constants in the low picomolar to nanomolar range, which may hamper accurate off-rate measurements with conventional bulk methods. Single-molecule force spectroscopy (SMFS) with the atomic force microscope measures the response of individual biomolecules to force, and in contrast to other single-molecule manipulation methods (i.e.&#160;optical tweezers), is optimal for studying high-affinity receptor-ligand interactions because of its ability to probe the high-force regime (&#62;120 pN). Here we present our complete protocol for studying cellulosomal protein assemblies at the single-molecule level. Using a protein topology derived from the native cellulosome, we worked with enzyme-dockerin and carbohydrate binding module-cohesin (CBM-cohesin) fusion proteins, each with an accessible free thiol group at an engineered cysteine residue. We present our site-specific surface immobilization protocol, along with our measurement and data analysis procedure for obtaining detailed binding parameters for the high-affinity complex. We demonstrate how to quantify single subdomain unfolding forces, complex rupture forces, kinetic off-rates, and potential widths of the binding well. The successful application of these methods in characterizing the cohesin-dockerin interaction responsible for assembly of multidomain cellulolytic complexes is further described.

Cellulosomes are multienzyme complexes designed for digesting cellulose. AFM-based SMFS was used to study the mechanical properties and folding configuration of cellulosome-associated protein assemblies. We present a complete workflow for protein immobilization, data acquisition, and data analysis to study the interactions of individual receptor-ligand complexes involved in cellulosome assembly.

Cellulosomes are discrete multienzyme complexes used by a subset of anaerobic bacteria and fungi to digest lignocellulosic substrates. Assembly of the ...

Bioengineering

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 ...

From a 2DE-Gel Spot to Protein Function: Lesson Learned From HS1 in Chronic Lymphocytic Leukemia

The identification of molecules involved in tumor initiation and progression is fundamental for understanding disease&#8217;s biology and, as a consequence, for the clinical management of patients. In the present work we will describe an optimized proteomic approach for the identification of molecules involved in the progression of Chronic Lymphocytic Leukemia (CLL). In detail, leukemic cell lysates are resolved by 2-dimensional Electrophoresis (2DE) and visualized as &#8220;spots&#8221; on the 2DE gels. Comparative analysis of proteomic maps allows the identification of differentially expressed proteins (in terms of abundance and post-translational modifications) that are picked, isolated and identified by Mass Spectrometry (MS). The biological function of the identified candidates can be tested by different assays (i.e. migration, adhesion and F-actin polymerization), that we have optimized for primary leukemic cells.

Here we describe a protocol that couples two proteomic techniques, namely 2-dimensional Electrophoresis (2DE) and Mass Spectrometry (MS), to identify differentially expressed/post-translational modified proteins among two or more groups of primary samples. This approach, together with functional experiments, allows the identification and characterization of prognostic markers/therapeutic targets.

The identification of molecules involved in tumor initiation and progression is fundamental for understanding disease&#8217;s biology and, as a ...

Medicine

Concanavalin A-Based Sedimentation Assay to Measure Substrate Binding of Glucan Phosphatases

Glucan phosphatases belong to the larger family of dual specificity phosphatases (DSP) that dephosphorylate glucan substrates, such as glycogen in animals and starch in plants. The crystal structures of glucan phosphatase with model glucan substrates reveal distinct glucan-binding interfaces made of DSP and carbohydrate-binding domains. However, quantitative measurements of glucan-glucan phosphatase interactions with physiologically relevant substrates are fundamental to the biological understanding of the glucan phosphatase family of enzymes and the regulation of energy metabolism. This manuscript reports a Concanavalin A (ConA)-based in vitro sedimentation assay designed to detect the substrate binding affinity of glucan phosphatases against different glucan substrates.&#160;As a proof of concept, the dissociation constant (KD) of glucan phosphatase Arabidopsis thaliana Starch Excess4 (SEX4) and amylopectin was determined. The characterization of SEX4 mutants and other members of the glucan phosphatase family of enzymes further demonstrates the utility of this assay to assess the differential binding of protein- carbohydrate interactions. These data demonstrate the suitability of this assay to characterize a wide range of starch and glycogen interacting proteins.

This method describes a lectin-based in vitro sedimentation assay to quantify the binding affinity of glucan phosphatase and amylopectin. This co-sedimentation assay is reliable for measuring glucan phosphatase substrate binding and can be applied to various solubilized glucan substrates.

Glucan phosphatases belong to the larger family of dual specificity phosphatases (DSP) that dephosphorylate glucan substrates, such as glycogen in animals ...

Reconstitution of Actin-Based Motility with Commercially Available Proteins

Many cell movements and shape changes and certain types of intracellular bacterial and organelle motility are driven by the biopolymer actin that forms a dynamic network at the surface of the cell, organelle, or bacterium. The biochemical and mechanical basis of force production during this process can be studied by reproducing actin-based movement in an acellular manner on inert surfaces such as beads that are functionalized and incubated with a controlled set of components. Under the appropriate conditions, an elastic actin network assembles at the bead surface and breaks open due to the stress generated by network growth, forming an "actin comet" that propels the bead forward. However, such experiments require the purification of a host of different actin-binding proteins, often putting them beyond the reach of non-specialists. This article details a protocol for reproducibly obtaining actin comets and motility of beads using commercially available reagents. Bead coating, bead size, and motility mixture can be altered to observe the effect on bead speed, trajectories, and other parameters. This assay can be used for testing the biochemical activities of different actin-binding proteins, and for performing quantitative physical measurements that shed light on active matter properties of actin networks. This will be a useful tool for the community, enabling the study of in vitro actin-based motility without expert knowledge in actin-binding protein purification.

This protocol describes how to produce actin comets on the surfaces of beads using commercially available protein ingredients. Such systems mimic the protrusive structures found in cells, and can be used to examine physiological mechanisms of force production in a simplified way.

Many cell movements and shape changes and certain types of intracellular bacterial and organelle motility are driven by the biopolymer actin that forms a ...

FtsZ Polymerization Assays: Simple Protocols and Considerations

During bacterial cell division, the essential protein FtsZ assembles in the middle of the cell to form the so-called Z-ring. FtsZ polymerizes into long filaments in the presence of GTP in vitro, and polymerization is regulated by several accessory proteins. FtsZ polymerization has been extensively studied in vitro using basic methods including light scattering, sedimentation, GTP hydrolysis assays and electron microscopy. Buffer conditions influence both the polymerization properties of FtsZ, and the ability of FtsZ to interact with regulatory proteins. Here, we describe protocols for FtsZ polymerization studies and validate conditions and controls using Escherichia coli and Bacillus subtilis FtsZ as model proteins. A low speed sedimentation assay is introduced that allows the study of the interaction of FtsZ with proteins that bundle or tubulate FtsZ polymers. An improved GTPase assay protocol is described that allows testing of GTP hydrolysis over time using various conditions in a 96-well plate setup, with standardized incubation times that abolish variation in color development in the phosphate detection reaction. The preparation of samples for light scattering studies and electron microscopy is described. Several buffers are used to establish suitable buffer pH and salt concentration for FtsZ polymerization studies. A high concentration of KCl is the best for most of the experiments. Our methods provide a starting point for the in vitro characterization of FtsZ, not only from E. coli and B. subtilis but from any other bacterium. As such, the methods can be used for studies of the interaction of FtsZ with regulatory proteins or the testing of antibacterial drugs which may affect FtsZ polymerization.

Polymerization of FtsZ is essential for bacterial cell division. In this report, we detail simple protocols to monitor FtsZ polymerization activity and discuss the influence of buffer composition. The protocols can be used to study the interaction of FtsZ with regulatory proteins or antibacterial drugs that affect FtsZ polymerization.

During bacterial cell division, the essential protein FtsZ assembles in the middle of the cell to form the so-called Z-ring. FtsZ polymerizes into long ...

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

Single-Molecule Measurement of Protein Interaction Dynamics Within Biomolecular Condensates

Biomolecular condensates formed via liquid-liquid phase separation (LLPS) have been considered critical in cellular organization and an increasing number of cellular functions. Characterizing LLPS in live cells is also important because aberrant condensation has been linked to numerous diseases, including cancers and neurodegenerative disorders. LLPS is often driven by selective, transient, and multivalent interactions between intrinsically disordered proteins. Of great interest are the interaction dynamics of proteins participating in LLPS, which are well-summarized by measurements of their binding residence time (RT), that is, the amount of time they spend bound within condensates. Here, we present a method based on live-cell single-molecule imaging that allows us to measure the mean RT of a specific protein within condensates. We simultaneously visualize individual protein molecules and the condensates with which they associate, use single-particle tracking (SPT) to plot single-molecule trajectories, and then fit the trajectories to a model of protein-droplet binding to extract the mean RT of the protein. Finally, we show representative results where this single-molecule imaging method was applied to compare the mean RTs of a protein at its LLPS condensates when fused and unfused to an oligomerizing domain. This protocol is broadly applicable to measuring the interaction dynamics of any protein that participates in LLPS.

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

Many intrinsically disordered proteins have been shown to participate in the formation of highly dynamic biomolecular condensates, a behavior important for numerous cellular processes. Here, we present a single-molecule imaging-based method for quantifying the dynamics by which proteins interact with each other in biomolecular condensates in live cells.

Biomolecular condensates formed via liquid-liquid phase separation (LLPS) have been considered critical in cellular organization and an increasing number ...

Measuring Protein Binding to F-actin by Co-sedimentation

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