This work was partially supported by the Deutsche Forschungsgemeinschaft (DFG) grant 254850309. The authors thank Madlen Luckner for critical reading of the manuscript.

The authors have nothing to disclose.

The experimental procedure described here allows the investigation of protein-protein trans interactions at cell-cell contacts, employing fluorescence fluctuation spectroscopy techniques, namely sFCCS and ccN&#38;B. These methods involve a statistical analysis of fluorescence fluctuations emitted by two spectrally separated FPs fused to the protein(s) of interest at a contact of two neighboring cells, each expressing one or the other fusion protein. The presence of trans complexes is quantified by probing the degree of co-diffusion of proteins in neighboring PMs. In addition to detailed protocols on sample preparation, data acquisition and analysis, this article provides experimental evidence of the successful application of the assay on the neuronal adhesion protein APLP1. We show that APLP1 undergoes specific, homotypic trans interactions at cell-cell contacts. Further, zinc ions promote the formation of APLP1 clusters at cell-cell contacts that provide a multivalent platform for trans interactions and, thus, induce enhanced trans binding.
In contrast to previous assays to detect such interactions based on disruptive biochemical methods6, the presented approach can be performed directly on living cells, with no need for fixation or isolation of protein complexes. Moreover, it provides molecular specificity and information by detection of fluorescent proteins genetically fused to the protein of interest, in contrast to previous qualitative assays8,9. Differently from other fluorescence based approaches such as FRET10 and fluorescence complementation11, there is no requirement for the fluorescent labels to be localized on the extracellular side (potentially interfering with protein-protein interactions). Nevertheless, it has to be noted that C-terminal FPs might still alter the binding to intracellular components, e.g., adaptor proteins that mediate interactions with the cytoskeleton. Notably, the assay is applicable to both homo- and heterotypic interactions.
A few requirements for a successful application of the presented assay are worth mentioning. The performed fluorescence fluctuation methods are based on temporal measurements that require bright and photostable monomeric FPs, which is a major constraint for many red FPs23. Even though the presented scanning scheme is particularly well suited for the investigation of the slow dynamics of transmembrane proteins, which are typically involved in cell-cell interactions, residual photobleaching may still occur. Therefore, we present a detailed description of correction schemes, i.e., a bleaching correction for sFCCS and boxcar filter for ccN&#38;B, which minimize such perturbations. Further, we discuss numerical alignment algorithms that effectively correct for other systematic perturbations, such as lateral movement of cells, and provide guidelines to remove transient instabilities. A major source of such instabilities is vesicular transport, i.e., intracellular vesicles carrying the protein of interest that transiently enter the focal volume. Although varying from case to case, this phenomenon can in general severely disturb data acquisition and analysis. However, the application of correction algorithms improves the stability of the acquisition, allowing extended measurement times that are needed to probe the especially slow diffusive dynamics. In this regard, it has to be underlined that the presence of diffusive dynamics is an essential condition for the method to work. The example of zinc ion mediated APLP1 clustering shows a drastic example of large, very slow protein complexes (D &#8776; 0.001 &#181;m2/s) that push the technique to its limits and prevent an accurate quantification of the underlying dynamics, due to large noise induced by the limited acquisition time30,17.
In this context, recent advances on combing sFCS and super-resolution approaches, e.g., scanning stimulated emission depletion FCS (STED-FCS), may be beneficial by providing a smaller focal volume and thus smaller effective diffusion times39,40. This could also help to resolve protein clusters in case of an inhomogeneous localization of the protein of interest at the cell-cell contacts, as observed for APLP1 in the presence of zinc. Unfortunately, a cross-correlation implementation of scanning STED-FCS, i.e., STED-FCCS, that could be directly applied here, has not been successfully demonstrated yet due to the difficulty of finding compatible dyes for two-color STED-FCS. In the case of sufficiently fast dynamics (D &#8805;&#160;~0.05 &#181;m2/s), sFCCS analysis allows to quantify the diffusion of proteins at cell-cell contacts or in other regions of the PM12.
Further, a brightness analysis can be performed for&#160;all data (sFCCS and ccN&#38;B), providing an estimate of the stoichiometry of trans protein complexes. It should be mentioned that the accuracy of the stoichiometry quantification increases with the amount of proteins that bind in trans (i.e., if there are only few cis complexes which could also affect the brightness determination). Brightness analysis does not allow resolving mixtures of different oligomeric states, e.g.,cis monomers and trans tetramers. More in general, it should be noted that N&#38;B cannot resolve mixtures of different oligomeric species homogeneously distributed in a sample. If multiple species are present within a pixel, one single value of brightness will be measured (i.e., a weighted average of the brightness of each oligomeric species). The statistical distribution of the observed brightness values (e.g., in different pixels) is not directly connected to the relative amounts of oligomeric species. To resolve such mixtures, other methods should be used (e.g., spatial intensity distribution analysis (SpIDA)41, photon counting histogram (PCH)42). Similarly, the quantification of the spectral cross-correlation in sFCCS provides an accurate quantification of the fraction of bound and unbound proteins only for simple, known stoichiometries, e.g., 1:1. For a more complex stoichiometry, further assumptions have to be made43. Overall, brightness analysis remains a powerful experimental tool, if the detector system and the fraction of non-fluorescent FPs are calibrated23 as described in the protocols.
Although the application presented here is focused on protein-protein interactions in adherent cells, the assay can be applied to a wider range of samples,&#160;e.g., suspension cells. For such systems, correction for lateral cell movement may be particularly crucial. In addition, it can be easily applied to model membrane systems such as GUVs or GPMVs, allowing the quantification of molecular interactions under well controlled conditions, e.g., different lipid compositions or membranes lacking an organized cytoskeleton. When performing sFCCS on such vesicles, vertical motion may cause additional signal fluctuations, but can be minimized when focusing on large vesicles. In this context, multiple combinations are promising, e.g., GUV-/ GPMV-cell mixing12. As shown in a recent publication, the formation of immune synapses can be successfully modeled by such systems44. Thus, the presented assay will certainly be useful for the investigation of a large variety of cell-cell interactions.

Many biological processes occur at the sites of cell-cell interactions, e.g., cell-cell adhesion1,2,3, cell-cell fusion4 and cellular recognition5. Such events are particularly important during the development of multicellular organisms and for cell-cell communication, e.g., during immune responses. These processes are typically mediated by proteins that are localized at the surface, i.e., at the plasma membrane (PM) of neighboring cells and undergo specific interactions at the cell-cell contact that are precisely regulated in space and time. In many cases, these interactions are direct homo- or heterotypic protein-protein trans interactions, but may also involve ions or ligands acting as extracellular linkers1. Although of fundamental importance, there is a lack of assays probing these specific protein-protein interactions directly in the native environment of living cells. Many methods either require cell disruption (e.g., biochemical assays such as co-immunoprecipitation6), fixation (e.g., some of the super-resolution optical microscopy techniques and electron microscopy of cell-cell contacts7), or are non-specific, e.g., aggregation/ adhesion assays8,9. To overcome this issue, fluorescence techniques have been implemented based on fluorescence resonance energy transfer (FRET)10 or fluorescence complementation11. However, to achieve sufficiently small distances between fluorophores, these methods require fluorescent labels on the extracellular side of the proteins10, potentially interfering with trans interactions.
Here, we present an alternative fluorescence-based assay for protein-protein interactions at cell-cell contacts. This approach combines fluorescence cross-correlation approaches (scanning fluorescence cross-correlation spectroscopy (sFCCS), cross-correlation number and brightness (ccN&#38;B)) and mixing of cells expressing a fusion construct of the protein of interest, e.g., an adhesion receptor. The investigated receptors in the two interacting cells are labeled with two spectrally separated fluorescent proteins (FPs), from the intracellular side (see Figure 1A).
The employed methods are based on the statistical analysis of fluorescence fluctuations induced by the diffusive motion of fluorescent fusion proteins through the focal volume of a confocal laser scanning microscope. More in detail, the assay probes the co-diffusion of the proteins of interest in both neighboring PMs at cell-cell contacts. If the proteins undergo trans interactions, these trans complexes will carry fluorescent proteins emitting in both spectral channels, causing correlated fluorescence fluctuations of both emitters. On the other hand, if no binding occurs, the number fluctuations of proteins in facing PMs will be independent, causing no correlated fluctuations. The acquisition can be performed in two ways: 1) sFCCS is based on a line-shaped scan across the cell-cell contact and effectively probes the interactions in a spot located in the contact region. Through a temporal analysis of fluorescence fluctuations, sFCCS provides also dynamics information, i.e., the diffusion coefficients of protein complexes; 2) ccN&#38;B is based on a pixel-wise analysis of a sequence of images acquired at the cell-cell contact regions. It has capability to probe and map interactions along the whole contact region (in one focal plane), but does not provide information on dynamics. Both methods can be combined with an analysis of the molecular brightness, i.e., the average fluorescence signal emitted in the time unit by single diffusing protein complexes and, thus, provide estimates of the stoichiometry of protein complexes at cell-cell contacts.
In this article, we provide detailed protocols for sample preparation, instrument calibration, data acquisition and analysis to perform the presented assay on a commercial confocal laser scanning microscope. The experiments can be performed on any instrument equipped with photon counting or analog detectors and an objective with high numerical aperture. We further discuss critical steps of the protocol and provide correction schemes for several processes causing artefactual signal fluctuations, e.g., detector noise, photobleaching or cell movement. Originally developed to probe interactions between adherent cells, the assay may be modified for suspension cells, or adapted to model membrane systems, e.g., giant unilamellar vesicles (GUVs) or giant plasma membrane vesicles (GPMVs), allowing the quantification of interactions in different lipid environments or in the absence of an organized cytoskeleton12,13.
Scanning fluorescence cross-correlation spectroscopy is a modified version of fluorescence cross-correlation spectroscopy14 and was specifically designed to probe slow diffusive dynamics in lipid membranes15. It is based on a line scan acquisition perpendicular to the PM containing the fluorescent proteins of interest. To probe interactions of two differently labeled protein species, the acquisition is performed in two spectral channels using two laser lines and two detection windows for&#160;spectrally separated fluorophores. Due to the slow diffusion dynamics of proteins in the PM (D&#8804;~1 &#181;m2/s), a cross-talk-free measurement can be performed by alternating the excitation scheme from line to line15. The analysis starts with: 1) an alignment algorithm correcting for lateral cell movement based on block-wise averaging of ~1000 lines, 2) determination of the position with maximum fluorescence signal, i.e., the PM position, in each block and 3) shifting of all blocks to a common origin12,15, separately in each channel. Then, an automatic selection of pixels corresponding to the PM is performed by selecting the central region from a Gaussian fit of the sum of all aligned lines (i.e., center &#177; 2.5&#963;). Integration of the signal in each line yields the membrane fluorescence time series F(t) in each channel (g = green channel, r = red channel). Note that the pixel size has to be small enough, e.g., &#60;200 nm, to reconstruct the shape of the point spread function and find its center, corresponding to the position of the PM. In the presence of substantial photobleaching, the fluorescence time series in each channel may be modeled&#160;with a double-exponential function and then corrected with the following formula:16
<img alt="Equation 1" src="/files/ftp_upload/58582/58582eq2.jpg" />.&#160; &#160; (1)
It is important to note that this formula effectively corrects both the amplitudes and diffusion times obtained from correlation analysis of F(t)c, compared to parameter estimates that would be obtained from the uncorrected F(t). Then, the auto- and cross-correlation functions (ACFs/ CCFs) of the fluorescence signals are calculated:
<img alt="Equation 2" src="/files/ftp_upload/58582/58582eq3v2.jpg" />, (2)
<img alt="Equation 3" src="/files/ftp_upload/58582/58582eq4v2.jpg" />, (3)
where &#948;Fi = Fi(t) - <img alt="Image 1" src="/files/ftp_upload/58582/58582img1.jpg" />Fi(t)<img alt="Image 2" src="/files/ftp_upload/58582/58582img2.jpg" /> and i&#160;=&#160;g,r.
A two-dimensional diffusion model is then fitted to all correlation functions (CFs):
<img alt="Equation 4" src="/files/ftp_upload/58582/58582eq7.jpg" />.&#160; &#160;(4)
Here, N denotes the number of fluorescent proteins in the observation volume and &#964;d the diffusion time for each channel. This model takes into account that in the described experimental setting, diffusion of proteins in the PM occurs in the x-z plane, in contrast to the commonly used configuration of fluorescence correlation spectroscopy (FCS) experiments on membranes probing diffusion in the x-y plane of the confocal volume17. The waist w0 and the structure factor S, describing the elongation wz of the focal volume in z, S = wz/w0, are obtained from a point FCS calibration measurement performed with spectrally similar dyes and same optical settings using already available values for the diffusion coefficient Ddye:
<img alt="Equation 5" src="/files/ftp_upload/58582/58582eq8.jpg" />,&#160; &#160; (5)
where &#964;d,dye is the measured average diffusion time of the dye molecules, obtained from fitting a model for three-dimensional diffusion to the data, taking into account transitions of a fraction T of all N molecules to a triplet state with a time constant &#964;&#964;:
<img alt="Equation 6" src="/files/ftp_upload/58582/58582eq9.jpg" />.&#160; &#160;(6)
Finally, diffusion coefficients (D), molecular brightness values (&#949;) and the relative cross-correlation of sFCCS data (rel.cc.) are calculated as follows:
<img alt="Equation 7" src="/files/ftp_upload/58582/58582eq10.jpg" />,&#160; &#160; (7)
<img alt="Equation 8" src="/files/ftp_upload/58582/58582eq11.jpg" />&#160;,&#160; &#160;(8)
<img alt="Equation 9" src="/files/ftp_upload/58582/58582eq12.jpg" />,&#160; &#160;(9)
where Gcross(0) is the amplitude of the cross-correlation function and <img alt="Equation 14" src="/files/ftp_upload/58582/58582eq14.jpg" /> is the amplitude of the autocorrelation function in the i-th channel.
This definition of the relative cross-correlation, i.e. using max instead of mean in Equation 9, takes into account that the maximum number of complexes of two protein species present at different concentrations is limited by the species present in a lower number.
Cross-correlation number and brightness is based on a moment analysis of the fluorescence intensity for each pixel of an image stack acquired over time at a fixed position in the sample, typically consisting of ~100-200 frames, with two spectral channels (g = green channel, r = red channel). From the temporal mean <img alt="Image 1" src="/files/ftp_upload/58582/58582img1.jpg" />I<img alt="Image 2" src="/files/ftp_upload/58582/58582img2.jpg" />i and variance&#160; <img alt="Equation 16" src="/files/ftp_upload/58582/58582eq16.jpg" />, the molecular brightness&#160;&#949;i and number ni are calculated in each pixel and spectral channel (i = g, r)18:
<img alt="Equation 10" src="/files/ftp_upload/58582/58582eq20.jpg" />,&#160; &#160;(10)
<img alt="Equation 11" src="/files/ftp_upload/58582/58582eq21.jpg" />. (11)
It is important to note that the given equations apply to the ideal case of a true photon-counting detector. For analog detection systems, the following equations apply19,20:
<img alt="Equation 12" src="/files/ftp_upload/58582/58582eq22.jpg" />,&#160; &#160;(12)
<img alt="Equation 13" src="/files/ftp_upload/58582/58582eq23.jpg" />.&#160; &#160;(13)
Here, S is the conversion factor between detected photons and the recorded digital counts, <img alt="Equation 24" src="/files/ftp_upload/58582/58582eq24.jpg" /> is the readout noise and offset refers to the detector intensity offset. Generally, these quantities should be calibrated, for any detector type, based on measuring the detector variance as a function of intensity for steady illumination19, e.g., a reflective metal surface or dried dye solution. The offset can be determined by measuring the count rate for a sample without excitation light. By performing a linear regression of the detector-associated variance <img alt="Equation 25" src="/files/ftp_upload/58582/58582eq25.jpg" /> versus intensity (I) plot, S and <img alt="Equation 24" src="/files/ftp_upload/58582/58582eq24.jpg" /> can be determined19:
<img alt="Equation 14" src="/files/ftp_upload/58582/58582eq27.jpg" />.&#160; &#160;(14)
Finally, the cross-correlation brightness is calculated in each pixel and is defined in general as21
<img alt="Equation 15" src="/files/ftp_upload/58582/58582eq28.jpg" />,&#160; &#160;(15)
where <img alt="Equation 29" src="/files/ftp_upload/58582/58582eq29.jpg" /> is the cross-variance <img alt="Equation 30" src="/files/ftp_upload/58582/58582eq30.jpg" />.
In order to filter long-lived fluctuations, all ccN&#38;B calculations are performed following a boxcar filtering, independently for each pixel22. Briefly, ni, &#949;i (i = g, r) and Bcc are calculated in sliding segments of e.g., 8-15 frames. The values thus obtained can be then averaged to obtain the final pixel number and brightness values.
Stoichiometry analysis 
In order to estimate the stoichiometry of protein complexes at cell-cell contacts, the molecular brightness can be separately analyzed in each spectral channel for the sFCCS or ccN&#38;B data. In sFCCS, one brightness value is obtained per measurement in each channel. In ccN&#38;B, a brightness histogram of all pixels corresponding to the cell-cell contact is obtained and the average (or median) value can be used as representative brightness for the measurement. By performing the same analysis on a monomeric reference, all brightness values can be normalized to directly obtain the average oligomeric state of the detected protein complexes. At this point, it is important to correct for the presence of non-fluorescent FPs that may result in an underestimation of the oligomeric state. This is typically performed by measuring the brightness of a homo-dimeric reference protein23,24 using one-color sFCS or number and brightness (N&#38;B).

<table><tbody><tr><td>DMEM growth medium</td><td>PAN-Biotech</td><td>P04-01548</td><td></td></tr><tr><td>DPBS w/o: Ca2+ and Mg2+</td><td>PAN-Biotech</td><td>P04-36500</td><td></td></tr><tr><td>DPBS w: Ca2+ and Mg2+</td><td>PAN-Biotech</td><td>P04-35500</td><td></td></tr><tr><td>Trypsin EDTA</td><td>PAN-Biotech</td><td>P10-023100</td><td></td></tr><tr><td>TurboFect Transfection Reagent</td><td>Thermo Fisher Scientific</td><td>R0531</td><td></td></tr><tr><td>HEK 293T cells</td><td>DSMZ</td><td>ACC 635</td><td></td></tr><tr><td>Alexa Fluor 488 NHS Ester</td><td>Thermo Fisher Scientific</td><td>A20000</td><td></td></tr><tr><td>Rhodamine B</td><td>Sigma-Alderich</td><td>83689-1G</td><td></td></tr><tr><td>Plasmid DNA</td><td>Addgene</td><td>NA</td><td>See reference 12 (Dunsing et. al., MBoC 2017),for a detailed description of all plasmids</td></tr><tr><td>6-well plate</td><td>Starlab</td><td>CC7672-7506</td><td></td></tr><tr><td>35-mm glass bottom dishes</td><td>CellVis</td><td>D35-14-1.5-N</td><td></td></tr><tr><td>Zeiss LSM780 confocal</td><td>Carl Zeiss</td><td>NA</td><td></td></tr><tr><td>MATLAB software package</td><td>MathWorks</td><td>&amp;nbsp;2015b&amp;nbsp;</td><td></td></tr><tr><td>Neubauer cell counting chamber</td><td>Marienfeld</td><td>640110</td><td></td></tr></tbody></table>

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.

Watch this Scientific Journal Video about A Fluorescence Fluctuation Spectroscopy Assay of Protein-Protein Interactions at Cell-Cell Contacts at JoVE.com

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

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 ...

a-fluorescence-fluctuation-spectroscopy-assay-protein-protein

Nanyang Technological University

Research

JoVE Journal

Biochemistry

11.3K Views.  University of Potsdam. Deze methode kan helpen om belangrijke vragen in het cel-naar-cel interactieveld te begrijpen, zoals welke receptoren bemiddelen hechting of herkenning in cel-naar-celcontacten en of specifieke liganden hun interacties activeren. Het belangrijkste voordeel van deze techniek is dat het mogelijk maakt om dergelijke interacties direct te onderzoeken in levende cellen zonder behoefte aan celfixatie of verstoring. Hoewel deze methode wordt gebruikt om interacties tussen gekweekte cellen te onderzo...

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

A Fluorescence-based Assay of Phospholipid Scramblase Activity

Scramblases translocate phospholipids across the membrane bilayer bidirectionally in an ATP-independent manner. The first scramblase to be identified and biochemically verified was opsin, the apoprotein of the photoreceptor rhodopsin. Rhodopsin is a G protein-coupled receptor localized in rod photoreceptor disc membranes of the retina where it is responsible for the perception of light. Rhodopsin&#39;s scramblase activity does not depend on its ligand 11-cis-retinal, i.e., the apoprotein opsin is also active as a scramblase. Although constitutive and regulated phospholipid scrambling play an important role in cell physiology, only a few phospholipid scramblases have been identified so far besides opsin. Here we describe a fluorescence-based assay of opsin&#39;s scramblase activity. Opsin is reconstituted into large unilamellar liposomes composed of phosphatidylcholine, phosphatidylglycerol and a trace quantity of fluorescent NBD-labeled PC (1-palmitoyl-2-{6-[7-nitro-2-1,3-benzoxadiazole-4-yl)amino]hexanoyl}-sn-glycero-3-phosphocholine). Scramblase activity is determined by measuring the extent to which NBD-PC molecules located in the inner leaflet of the vesicle are able to access the outer leaflet where their fluorescence is chemically eliminated by a reducing agent that cannot cross the membrane. The methods we describe have general applicability and can be used to identify and characterize scramblase activities of other membrane proteins.

We describe a fluorescence-based assay to measure phospholipid scrambling in large unilamellar liposomes reconstituted with opsin.

Scramblases translocate phospholipids across the membrane bilayer bidirectionally in an ATP-independent manner. The first scramblase to be identified and ...

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 ...

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 ...

Fluorescence Biomembrane Force Probe: Concurrent Quantitation of Receptor-ligand Kinetics and Binding-induced Intracellular Signaling on a Single Cell

Membrane receptor-ligand interactions mediate many cellular functions. Binding kinetics and downstream signaling triggered by these molecular interactions are likely affected by the mechanical environment in which binding and signaling take place. A recent study demonstrated that mechanical force can regulate antigen recognition by and triggering of the T-cell receptor (TCR). This was made possible by a new technology we developed and termed fluorescence biomembrane force probe (fBFP), which combines single-molecule force spectroscopy with fluorescence microscopy. Using an ultra-soft human red blood cell as the sensitive force sensor, a high-speed camera and real-time imaging tracking techniques, the fBFP is of ~1 pN (10-12 N), ~3 nm and ~0.5 msec in force, spatial and temporal resolution. With the fBFP, one can precisely measure single receptor-ligand binding kinetics under force regulation and simultaneously image binding-triggered intracellular calcium signaling on a single live cell. This new technology can be used to study other membrane receptor-ligand interaction and signaling in other cells under mechanical regulation.

We describe a technique for concurrently measuring force-regulated single receptor-ligand binding kinetics and real-time imaging of calcium signaling in a single T lymphocyte.

Membrane receptor-ligand interactions mediate many cellular functions. Binding kinetics and downstream signaling triggered by these molecular interactions ...

Bioengineering

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

We present a protocol and workflow to perform live cell dual-color fluorescence cross-correlation spectroscopy (FCCS) combined with F&#246;rster Resonance Energy transfer (FRET) to study membrane receptor dynamics in live cells using modern fluorescence labeling techniques. In dual-color FCCS, where the fluctuations in fluorescence intensity represent the dynamic &#34;fingerprint&#34; of the respective fluorescent biomolecule, we can probe co-diffusion or binding of the receptors. FRET, with its high sensitivity to molecular distances, serves as a well-known &#34;nanoruler&#34; to monitor intramolecular changes. Taken together, conformational changes and key parameters such as local receptor concentrations and mobility constants become accessible in cellular settings.
Quantitative fluorescence approaches are challenging in cells due to high noise levels and the vulnerability of the sample. Here we show how to perform this experiment, including the calibration steps using dual-color labeled &#946;2-adrenergic receptor (&#946;2AR) labeled with eGFP and SNAP-tag-TAMRA. A step-by-step data analysis procedure is provided using open-source software and templates that are easy to customize.
Our guideline enables researchers to unravel molecular interactions of biomolecules in live cells in situ with high reliability despite the limited signal-to-noise levels in live cell experiments. The operational window of FRET and particularly FCCS at low concentrations allows quantitative analysis at near-physiological conditions.

We present an experimental protocol and data analysis workflow to perform live cell dual-color fluorescence cross correlation spectroscopy (FCCS) combined with F&#246;rster Resonance Energy transfer (FRET) to study membrane receptor dynamics in live cells using modern fluorescence labeling techniques.

We present a protocol and workflow to perform live cell dual-color fluorescence cross-correlation spectroscopy (FCCS) combined with F&#246;rster Resonance ...

Detecting and Characterizing Protein Self-Assembly In Vivo by Flow Cytometry

Protein self-assembly governs protein function and compartmentalizes cellular processes in space and time. Current methods to study it suffer from low-sensitivity, indirect read-outs, limited throughput, and/or population-level rather than single-cell resolution. We designed a flow cytometry-based single methodology that addresses all of these limitations: Distributed Amphifluoric FRET or DAmFRET. DAmFRET detects and quantifies protein self-assemblies by sensitized emission FRET in vivo, enables deployment across model systems-from yeast to human cells-and achieves sensitive, single-cell, high-throughput read-outs irrespective of protein localization or solubility.

This article describes a FRET-based flow cytometry protocol to quantify protein self-assembly in both S. cerevisiae and HEK293T cells.

Protein self-assembly governs protein function and compartmentalizes cellular processes in space and time. Current methods to study it suffer from ...

Biology

Assessing Protein Interactions in Live-Cells with FRET-Sensitized Emission

F&#246;rster Resonance Energy Transfer (FRET) is the radiationless transfer of energy from an excited donor to an acceptor molecule and depends upon the distance and orientation of the molecules as well as the extent of overlap between the donor emission and acceptor absorption spectra. FRET permits to study the interaction of proteins in the living cell over time and in different subcellular compartments. Different intensity-based algorithms to measure FRET using microscopy have been described in the literature. Here, a protocol and an algorithm are provided to quantify FRET efficiency based on measuring both the sensitized emission of the acceptor and quenching of the donor molecule. The quantification of ratiometric FRET in the living cell not only requires the determination of the crosstalk (spectral spill-over, or bleed-through) of the fluorescent proteins but also the detection efficiency of the microscopic setup. The protocol provided here details how to assess these critical parameters.

F&#246;rster Resonance Energy Transfer (FRET) between two fluorophore molecules can be used for studying protein interactions in the living cell. Here, a protocol is provided as to how to measure FRET in live cells by detecting sensitized emission of the acceptor and quenching of the donor molecule using confocal laser scanning microscopy.

F&#246;rster Resonance Energy Transfer (FRET) is the radiationless transfer of energy from an excited donor to an acceptor molecule and depends upon the ...

Fluorescence Fluctuation Spectroscopy to Study Protein Interaction at Cell Contacts

Source: Dunsing, V. et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/58582">A Fluorescence Fluctuation Spectroscopy Assay of Protein-Protein Interactions at Cell-Cell Contacts.</a> J. Vis. Exp.&#160;(2018).
This video demonstrates the use of fluorescence fluctuation spectroscopy to detect the interaction among cell surface proteins at cell-cell contacts. By expressing the transmembrane adhesion receptor of interest labeled with a fluorescent protein and mixing two different cell populations harboring two spectrally separated fluorescent labels, the trans-interaction between the receptors of two neighboring cells with different-colored fluorescence is assessed via cross-correlation in the fluctuations of fluorescence intensity.

Source: Dunsing, V. et al. A Fluorescence Fluctuation Spectroscopy Assay of Protein-Protein Interactions at Cell-Cell Contacts. J. Vis. Exp.&#160;(2018).
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JoVE Encyclopedia of Experiments

Biological Techniques

Biomolecular Interaction Detection Techniques

Protein-protein interactions

Fluorescence Fluctuation Spectroscopy to Study Protein Homo-Oligomerization

Source: Nolan, R., et al. Calibration-free In Vitro <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/58157">Quantification of Protein Homo-oligomerization Using Commercial Instrumentation and Free, Open Source Brightness Analysis Software.</a> J. Vis. Exp. (2018)
This video demonstrates the technique of fluorescence fluctuation spectroscopy (FFS) to study protein homo-oligomerization. Fluorescently-labeled proteins in a sample are dimerized using an agent when studying protein oligomerization using FFS. Using a confocal microscope, as the proteins move in and out of the small observation volume, the analysis of the fluctuations in the brightness of the fluorescent molecules is performed to determine the oligomeric state of the proteins.

Source: Nolan, R., et al. Calibration-free In Vitro Quantification of Protein Homo-oligomerization Using Commercial Instrumentation and Free, Open Source ...

Fluorescence Anisotropy-Based Detection of Protein-Protein Interactions

Mix 3 nanomoles of the SBDS-FlAsH protein with 3 nanomoles of the Lumio Green dye in a 5-microliter volume of anisotropy buffer. Let the reaction proceed for 8 hours at 4 degrees Celsius. After 8 hours, dialyze the sample against the anisotropy buffer overnight to remove the free dye. Measure the absorbance at 280 nanometers and 508 nanometers in a spectrophotometer using a quartz cuvette. Then, use the Lambert-Beer law to quantify the percent of labeled protein as described in the text protocol.
Before measuring the anisotropy value, each titration step of the protein-ligand should be done carefully, ensuring that the entire sample is dispensed into the solution, and becomes homogeneous.
In a fluorescence cuvette, place 200 microliters of 30 nanomolar SBDS-FlAsH in anisotropy buffer, and titrate 2 microliters of 30 micromolar EFL1. Mix thoroughly, and let the reaction stand for 3 minutes before measuring the anisotropy and fluorescence value. Repeat this process until a total volume of 40 microliters of EFL1 has been added. As a final step, fit the data to a presumed binding model, as described in the text protocol.

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

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