We acknowledge Dr Ludovic Richert for his valuable assistance on FLIM data acquisition and for the technical maintenance and development of the FLIM setup. This work was funded by grants from Fondation pour la Recherche en Chimie (https://icfrc.fr/). VN is funded by the Fondation pour la Recherche M&#233;dicale (FRM&#8208;SPF201809006906). YM is grateful to the Institut Universitaire de France (IUF) for support and providing additional time to be dedicated to research. IJS and JG acknowledge the Institute on Drug Delivery of Strasbourg for its financial support.

The authors have nothing to disclose.

FLIM-FRET offers some key advantages over intensity-based FRET imaging. Fluorescence lifetime is an intrinsic parameter of the fluorophore. As a consequence, it is not dependent on local concentrations of fluorophores neither on the intensity of the light excitation. The fluorescence lifetime is additionally also poorly affected by photo-bleaching. It is particularly interesting to evidence PPIs in cells where local proteins concentrations can be highly heterogeneous throughout the subcellular compartments or regions. FL...

Protein-protein interactions (PPIs) control various key processes in cells1. The roles of PPIs differ based on protein composition, affinities functions and locations in cells2. PPIs can be investigated via different techniques3. For example, co-immunoprecipitation is a relatively simple, robust, and inexpensive technique commonly used tool to identify or confirm PPIs. However, studying PPIs can be challenging when the interacting proteins have low expression levels or when the interactions are transient or relevant only in specific environments. Studying PPIs occurring between the different enzymes of the pyoverdine pathway in P. aeruginosa requires that the repression of the general iron-co-factored repressor Fur is relieved to allow the expression of all the proteins of the pyoverdine pathway to be expressed in the cell4,5,6. This common regulation for all the proteins of the pathway results in timely expressions in the cell expected to promote their interactions. The diversity in term of size, nature, expression levels and the number of proteins of this metabolic pathway make it difficult for study in reconstituted systems6. Exploring PPIs in their cellular environment is therefore critical to further understand the biological functions of proteins in their native context.
Only few methods including fluorescence allow exploring PPIs in living cells7. Amongst the different fluorescence parameters that can be measured, the fluorescence lifetime (i.e., the average time a fluorophore remains in its excited state before emitting a photon) is likely one of the most interesting parameters to explore in living cells. The fluorescence lifetime of a fluorophore is highly sensitive to its environment and FLIM can therefore provide chemical or physical information regarding the fluorophore surroundings8. This includes the presence of F&#246;rster resonance energy transfer (FRET) that can occur in the presence of an &#8220;acceptor&#8221; of fluorescence located at a short distance of a fluorescence &#8220;donor&#8221;. Energy transfer results in significant shortening of the donor fluorescence lifetime (Figure 1A), making Fluorescence Lifetime Imaging Microscopy (FLIM) a powerful approach to explore protein-protein interactions directly in live cells. FLIM can additionally provide spatial information about where the interactions take place in cells7,8. This approach is extremely powerful for investigating PPIs in situations where the labeling with fluorophores of the two interacting partners is possible.
For FRET to occur - critical conditions on the distance between two fluorophores are required8,9. The two fluorophores should not be distant from each other by more than 10 nm. Therefore, cautions must be taken when designing FLIM-FRET experiments to ensure that the donor and the acceptor of fluorescence have a chance to be located close to each other in the interacting complex. While this may seem constraining, it is in fact a true advantage as the distance-dependence of FRET ensures that two labelled proteins undergoing FRET have to physically interact (Figure 1A). The difficulties at getting clear answers about PPI in colocalization experiments (two colocalized proteins may not necessarily interact) are therefore not an issue using FLIM-FRET.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61602/61602fig1.jpg" /> 
Figure 1: FLIM-FRET analysis principle. Each pixel of the FLIM-FRET multidimensional image contains information about the fluorescence decay recorded at this particular location (#counts = number of detected photons in the channel t). (A) The classical representation of the FLIM image is usually a false-color lifetime encoded 2D image (left). A decrease in the mean fluorescence lifetime of the donor - as seen by a change in the color scale - can be observed in the presence of FRET and is informative about the presence of PPIs in this spatial area. (B) Overlap between the donor emission spectrum and the acceptor absorption spectrum is necessary for FRET to occur. <a href="https://www.jove.com/files/ftp_upload/61602/61602fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
A second requirement for FRET is that the emission spectrum of the donor and the absorption spectra of the acceptor should overlap8 (Figure 1B). The fluorescence excitation of the donor should be at wavelengths that contribute very little to the direct fluorescence excitation of the acceptor. Not all combinations of fluorophores are possible and we additionally recommend to preferentially use donors with monoexponential fluorescence decays to facilitate FLIM-FRET interpretations10. Several couples of fluorescence proteins meet these requirements, including the popular eGFP-mCherry couple11 (for a review on the palette of available fluorescent protein FRET pairs see12,13).
FLIM-FRET allows measuring the fluorescence lifetime decay of a FRET donor at every pixel of a FLIM image (Figure 1A). There are two major techniques to determine fluorescence lifetime that differ in acquisition and analysis: frequency-domain (FD)14 and time-domain (TD). TD FLIM is more widespread and is performed using a pulsed illumination combined with different possible detection configurations including gating methods15, streak camera16 or time-correlated single photon counting (TCSPC) techniques8. For both FD and TD techniques, fluorescence lifetime is not directly measured but requires an analysis of the measured data to estimate the lifetime(s) or the presence of interactions. For TCSPC techniques, the most widely used analysis relies on fitting the decays with single or multi exponential functions using least square iterative re-convolutions that minimize the weighted sum of the residuals.
Finally, FLIM-FRET can be performed both by using single photon or multiphoton excitations. The latest have several advantages like reducing autofluorescence and photodamage out of the focal plane. Multiphoton excitations allow also a longer excitation depth if working in thick 3D samples8. On the contrary, single photon excitation is usually more efficient as the two-photon absorption cross sections of fluorescent proteins are limited17.
Here, we propose a protocol for FLIM-FRET measurements of PPIs in live P. aeruginosa in the particular case of two interacting proteins (PvdA and PvdL) expressed with highly different numbers of copies to demonstrate the quality and robustness of the technique at revealing critical features of PPIs. PvdA and PvdL proteins are involved in pyoverdine biosynthesis. PvdA is a L-ornithine N5-oxygenase and synthesizes the L-N5-formyl-N5-hydroxyornithine from L-ornithine by hydroxylation (PvdA) and formylation (PvdF)18. PvdL is a non-ribosomal peptide synthesis (NRPS) enzyme composed of four modules. The first module catalyzes the acylation of myristic acid. The second module catalyzes the activation of L-Glu and its condensation to the myristic-coA. Then, the third module condenses a L-Tyr amino acid that is then isomerized in D-Tyr. Finally, the fourth module binds a L-Dab (Diaminobutyric acid) amino acid to form the acylated tripeptide L-Glu/D-Tyr/L-Dab6. PvdL is thus responsible for the synthesis of the three first amino acids of the pyoverdine precursor. The interaction of PvdA protein with PvdL is surprising as PvdL, on the contrary to PvdI and PvdJ, does not carry a module specific for the L-N5-formyl-N5-hydroxyornithine. This interaction suggest that all the enzymes responsible for the pyoverdine precursor biosynthesis are arranged in large transient and dynamic multi-enzymatic complexes19,20.
In this report we explain in detail how to construct the bacterial strains expressing natively the two interacting eGFP and mCherry labelled proteins. We also describe sample preparation and conditions for efficient FLIM-FRET cell imaging. Finally, we propose a step-by-step tutorial for image analysis including a recently developed tool providing advanced visualization possibilities for straightforward interpretation of complex FLIM-FRET data. With this report, we would like to convince not only adventurous but most biologists that FRET-FLIM is an accessible and powerful technique able to address their questions about PPIs directly in the native cellular environment.

<table><tbody><tr><td>525/50 nm band-pass filter</td><td>F37-516, AHF, Germany</td><td></td><td></td></tr><tr><td>680 nm short pass filter</td><td>F75-680, AHF, Germany</td><td></td><td></td></tr><tr><td>Agarose</td><td>Sigma-Aldrich</td><td>A9539</td><td></td></tr><tr><td>Ammonium Sulfate (NH4)2SO4</td><td>Sigma-Aldrich</td><td>A4418</td><td></td></tr><tr><td>DreamTaq DNA polymerase 5U/&amp;mu;L</td><td>ThermoFisher Scientific</td><td>EP0714</td><td></td></tr><tr><td>E. coli TOP10</td><td>Invitrogen</td><td>C404010</td><td></td></tr><tr><td>Fiber-coupled avalanche photo-diode</td><td>SPCM-AQR-14- FC, Perkin Elmer</td><td></td><td></td></tr><tr><td>Glass coverslips (Thickness No. 1.5, 20&amp;times;20mm</td><td>Knitel glass</td><td>MS0011</td><td></td></tr><tr><td>High-Fidelity DNA polymerase Phusion 2U/&amp;mu;L</td><td>ThermoFisher Scientific</td><td>F530S</td><td></td></tr><tr><td>Lysogeny broth (LB)</td><td>Millipore</td><td>1.10285</td><td></td></tr><tr><td>Magnesium Sulfate Heptahydrate (MgSO4 . 7H2O)</td><td>Sigma-Aldrich</td><td>10034-99-8</td><td></td></tr><tr><td>Microscope slides (25&amp;times;75mm)</td><td>Knitel glass</td><td>MS0057</td><td></td></tr><tr><td>NucleoSpin Gel and PCR Clean-up</td><td>Macherey-Nagel</td><td>740609.50</td><td></td></tr><tr><td>NucleoSpin Plasmid</td><td>Macherey-Nagel</td><td>740588.10</td><td></td></tr><tr><td>Potassium Phosphate Dibasic (K2HPO4)</td><td>Sigma-Aldrich</td><td>RES20765</td><td></td></tr><tr><td>Potassium Phosphate Monobasic (KH2PO4)</td><td>Sigma-Aldrich</td><td>P5655</td><td></td></tr><tr><td>Sodium Succinate (Disodium)</td><td>Sigma-Aldrich</td><td>14160</td><td></td></tr><tr><td>SPCImage, SPCM software</td><td>Becker &amp;amp; Hickl</td><td></td><td></td></tr><tr><td>Sterile inoculating loop</td><td>Nunc</td><td>7648-1PAK</td><td></td></tr><tr><td>T4 DNA ligase 1U/&amp;mu;L</td><td>ThermoFisher Scientific</td><td>15224017</td><td></td></tr><tr><td>TCSPC module</td><td>SPC830, Becker &amp;amp; Hickl, Germany</td><td></td><td></td></tr><tr><td>Ti:Sapphire laser</td><td>Insight DeepSee, Spectra Physics</td><td></td><td></td></tr><tr><td>Tubes 50mL</td><td>Falcon</td><td>352070</td><td></td></tr></tbody></table>

flim-fret measurements of protein-protein interactions in live bacteria.

Protein-protein interactions (PPIs) control various key processes in cells. Fluorescence lifetime imaging microscopy (FLIM) combined with F&#246;rster resonance energy transfer (FRET) provide accurate information about PPIs in live cells. FLIM-FRET relies on measuring the fluorescence lifetime decay of a FRET donor at each pixel of the FLIM image, providing quantitative and accurate information about PPIs and their spatial cellular organizations. We propose here a detailed protocol for FLIM-FRET measurements that we applied to monitor PPIs in live Pseudomonas aeruginosa in the particular case of two interacting proteins expressed with highly different copy numbers to demonstrate the quality and robustness of the technique at revealing critical features of PPIs. This protocol describes in detail all the necessary steps for PPI characterization - starting from bacterial mutant constructions up to the final analysis using recently developed tools providing advanced visualization possibilities for a straightforward interpretation of complex FLIM-FRET data.

Empirical cumulative distribution functions (ecdf) of the fluorescence lifetimes measured for the different bacterial strains are shown in Figure 8. If FRET occurs, the ecdfs are shifted towards the shorter-lived lifetimes (Figure 8A,8B). Note that when the interaction of the two proteins results in a long distance between the two fluorophores, no FRET can occur (Figure 8C). This situation cannot be distinguished fr...

Watch this Scientific Journal Video about FLIM-FRET Measurements of Protein-Protein Interactions in Live Bacteria. at JoVE.com

FLIM-FRET Measurements of Protein-Protein Interactions in Live Bacteria.

We describe here a protocol to characterize protein-protein interactions between two highly-differently expressed proteins in live Pseudomonas aeruginosa using FLIM-FRET measurements. The protocol includes bacteria strain constructions, bacteria immobilization, imaging and post-imaging data analysis routines.

Protein-protein interactions (PPIs) control various key processes in cells. Fluorescence lifetime imaging microscopy (FLIM) combined with F&#246;rster ...

flim-fret-measurements-protein-protein-interactions-live

Research

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Biology

9.2K Views.  Laboratoire de Bioimagerie et Pathologies, UMR CNRS 7021. We describe here a protocol to characterize protein-protein interactions between two highly-differently expressed proteins in live Pseudomonas aeruginosa using FLIM-FRET measurements. The protocol includes bacteria strain constructions, bacteria immobilization, imaging and post-imaging data analysis routines.

Video: FLIM-FRET Measurements of Protein-Protein Interactions in Live Bacteria.

Using In Vitro Fluorescence Resonance Energy Transfer to Study the Dynamics Of Protein Complexes at a Millisecond Time Scale

Proteins are the primary operators of biological systems, and they usually interact with other macro- or small molecules to carry out their biological functions. Such interactions can be highly dynamic, meaning the interacting subunits are constantly associated and dissociated at certain rates. While measuring the binding affinity using techniques such as quantitative pull-down reveals the strength of the interaction, studying the binding kinetics provides insights on how fast the interaction occurs and how long each complex can exist. Furthermore, measuring the kinetics of an interaction in the presence of an additional factor, such as a protein exchange factor or a drug, helps reveal the mechanism by which the interaction is regulated by the other factor, providing important knowledge for the advancement of biological and medical research. Here, we describe a protocol for measuring the binding kinetics of a protein complex that has a high intrinsic association rate and can be dissociated quickly by another protein. The method uses fluorescence resonance energy transfer to report the formation of the protein complex in vitro, and it enables monitoring the fast association and dissociation of the complex in real time on a stopped-flow fluorimeter. Using this assay, the association and dissociation rate constants of the protein complex are quantified.

Protein-protein interactions are critical for biological systems, and studies of the binding kinetics provide insights into the dynamics and function of protein complexes. We describe a method that quantifies the kinetic parameters of a protein complex using fluorescence resonance energy transfer and the stopped-flow technique. &#160;

Proteins are the primary operators of biological systems, and they usually interact with other macro- or small molecules to carry out their biological ...

Biochemistry

Measuring TCR-pMHC Binding In Situ using a FRET-based Microscopy Assay

T-cells are remarkably specific and effective when recognizing antigens in the form of peptides embedded in MHC molecules (pMHC) on the surface of Antigen Presenting Cells (APCs). This is despite T-cell antigen receptors (TCRs) exerting usually a moderate affinity (&#181;M range) to antigen when binding is measured in vitro1. In view of the molecular and cellular parameters contributing to T-cell antigen sensitivity, a microscopy-based methodology has been developed as a means to monitor TCR-pMHC binding in situ, as it occurs within the synapse of a live T-cell and an artificial and functionalized glass-supported planar lipid bilayer (SLB), which mimics the cell membrane of an Antigen presenting Cell (APC) 2. Measurements are based on F&#246;rster Resonance Energy Transfer (FRET) between a blue- and red-shifted fluorescent dye attached to the TCR and the pMHC. Because the efficiency of FRET is inversely proportional to the sixth power of the inter-dye distance, one can employ FRET signals to visualize synaptic TCR-pMHC binding. The sensitive of the microscopy approach supports detection of single molecule FRET events. This allows to determine the affinity and off-rate of synaptic TCR-pMHC interactions and in turn to interpolate the on-rate of binding. Analogous assays could be applied to measure other receptor-ligand interactions in their native environment.

Measuring TCR-pMHC Binding In Situ using a FRET-based Microscopy Assay

This manuscript describes how to conduct (single molecule) F&#246;rster Resonance Energy Transfer (FRET)- based assays to measure the binding dynamics between T-cell antigen receptor (TCR) and antigenic peptide-loaded MHC molecules as they occur within the immunological synapse of a T-cell in contact with a functionalized planar supported lipid bilayer.

T-cells are remarkably specific and effective when recognizing antigens in the form of peptides embedded in MHC molecules (pMHC) on the surface of Antigen ...

Bioengineering

A Protocol for Using F&#246;rster Resonance Energy Transfer (FRET)-force Biosensors to Measure Mechanical Forces across the Nuclear LINC Complex

The LINC complex has been hypothesized to be the critical structure that mediates the transfer of mechanical forces from the cytoskeleton to the nucleus. Nesprin-2G is a key component of the LINC complex that connects the actin cytoskeleton to membrane proteins (SUN domain proteins) in the perinuclear space. These membrane proteins connect to lamins inside the nucleus. Recently, a F&#246;rster Resonance Energy Transfer (FRET)-force probe was cloned into mini-Nesprin-2G (Nesprin-TS (tension sensor)) and used to measure tension across Nesprin-2G in live NIH3T3 fibroblasts. This paper describes the process of using Nesprin-TS to measure LINC complex forces in NIH3T3 fibroblasts. To extract FRET information from Nesprin-TS, an outline of how to spectrally unmix raw spectral images into acceptor and donor fluorescent channels is also presented. Using open-source software (ImageJ), images are pre-processed and transformed into ratiometric images. Finally, FRET data of Nesprin-TS is presented, along with strategies for how to compare data across different experimental groups.

A number of FRET-based force biosensors have recently been developed, enabling the protein-specific resolution of intracellular force. In this protocol, we demonstrate how one of these sensors, designed for the linker of the nucleoskeleton-cytoskeleton (LINC) complex protein Nesprin-2G can be used to measure actomyosin forces on the nuclear LINC complex.

The LINC complex has been hypothesized to be the critical structure that mediates the transfer of mechanical forces from the cytoskeleton to the nucleus. ...

Imaging Protein-protein Interactions in vivo

Protein-protein interactions are a hallmark of all essential cellular processes. However, many of these interactions are transient, or energetically weak, preventing their identification and analysis through traditional biochemical methods such as co-immunoprecipitation. In this regard, the genetically encodable fluorescent proteins (GFP, RFP, etc.) and their associated overlapping fluorescence spectrum have revolutionized our ability to monitor weak interactions in vivo using F&ouml;rster resonance energy transfer (FRET)1-3. Here, we detail our use of a FRET-based proximity assay for monitoring receptor-receptor interactions on the endothelial cell surface.

Imaging Protein-protein Interactions in vivo

This protocol describes how to image protein-protein interactions using a FRET-based proximity assay.

Protein-protein interactions are a hallmark of all essential cellular processes. However, many of these interactions are transient, or energetically weak, ...

Measurement of Force-Sensitive Protein Dynamics in Living Cells Using a Combination of Fluorescent Techniques

Cells sense and respond to physical cues in their environment by converting mechanical stimuli into biochemically-detectable signals in a process called mechanotransduction. A crucial step in mechanotransduction is the transmission of forces between the external and internal environments. To transmit forces, there must be a sustained, unbroken physical linkage created by a series of protein-protein interactions. For a given protein-protein interaction, mechanical load can either have no effect on the interaction, lead to faster disassociation of the interaction, or even stabilize the interaction. Understanding how molecular load dictates protein turnover in living cells can provide valuable information about the mechanical state of a protein, in turn elucidating its role in mechanotransduction. Existing techniques for measuring force-sensitive protein dynamics either lack direct measurements of protein load or rely on the measurements performed outside of the cellular context. Here, we describe a protocol for the F&#246;rster resonance energy transfer-fluorescence recovery after photobleaching (FRET-FRAP) technique, which enables the measurement of force-sensitive protein dynamics within living cells. This technique is potentially applicable to any FRET-based tension sensor, facilitating the study of force-sensitive protein dynamics in variety of subcellular structures and in different cell types.

Here, we present a protocol for the simultaneous use of F&#246;rster resonance energy transfer-based tension sensors to measure protein load and fluorescence recovery after photobleaching to measure protein dynamics enabling the measurement of force-sensitive protein dynamics within living cells.

Cells sense and respond to physical cues in their environment by converting mechanical stimuli into biochemically-detectable signals in a process called ...

FRET Microscopy for Real-time Monitoring of Signaling Events in Live Cells Using Unimolecular Biosensors

F&ouml;rster resonance energy transfer (FRET) microscopy continues to gain increasing interest as a technique for real-time monitoring of biochemical and signaling events in live cells and tissues. Compared to classical biochemical methods, this novel technology is characterized by high temporal and spatial resolution. FRET experiments use various genetically-encoded biosensors which can be expressed and imaged over time in situ or in vivo1-2. Typical biosensors can either report protein-protein interactions by measuring FRET between a fluorophore-tagged pair of proteins or conformational changes in a single protein which harbors donor and acceptor fluorophores interconnected with a binding moiety for a molecule of interest3-4. Bimolecular biosensors for protein-protein interactions include, for example, constructs designed to monitor G-protein activation in cells5, while the unimolecular sensors measuring conformational changes are widely used to image second messengers such as calcium6, cAMP7-8, inositol phosphates9 and cGMP10-11. Here we describe how to build a customized epifluorescence FRET imaging system from single commercially available components and how to control the whole setup using the Micro-Manager freeware. This simple but powerful instrument is designed for routine or more sophisticated FRET measurements in live cells. Acquired images are processed using self-written plug-ins to visualize changes in FRET ratio in real-time during any experiments before being stored in a graphics format compatible with the build-in ImageJ freeware used for subsequent data analysis. This low-cost system is characterized by high flexibility and can be successfully used to monitor various biochemical events and signaling molecules by a plethora of available FRET biosensors in live cells and tissues. As an example, we demonstrate how to use this imaging system to perform real-time monitoring of cAMP in live 293A cells upon stimulation with a &beta;-adrenergic receptor agonist and blocker.

F&#246;rster resonance energy transfer (FRET) microscopy is a powerful technique for real-time monitoring of signaling events in live cells using various biosensors as reporters. Here we describe how to build a customized epifluorescence FRET imaging system from commercially available components and how to use it for FRET experiments.

F&ouml;rster resonance energy transfer (FRET) microscopy  continues to gain increasing interest as a technique for real-time monitoring  of biochemical ...

Investigating Protein-protein Interactions in Live Cells Using Bioluminescence Resonance Energy Transfer

Assays based on Bioluminescence Resonance Energy Transfer (BRET) provide a sensitive and reliable means to monitor protein-protein interactions in live cells. BRET is the non-radiative transfer of energy from a 'donor' luciferase enzyme to an 'acceptor' fluorescent protein. In the most common configuration of this assay, the donor is Renilla reniformis luciferase and the acceptor is Yellow Fluorescent Protein (YFP). Because the efficiency of energy transfer is strongly distance-dependent, observation of the BRET phenomenon requires that the donor and acceptor be in close proximity. To test for an interaction between two proteins of interest in cultured mammalian cells, one protein is expressed as a fusion with luciferase and the second as a fusion with YFP. An interaction between the two proteins of interest may bring the donor and acceptor sufficiently close for energy transfer to occur. Compared to other techniques for investigating protein-protein interactions, the BRET assay is sensitive, requires little hands-on time and few reagents, and is able to detect interactions which are weak, transient, or dependent on the biochemical environment found within a live cell. It is therefore an ideal approach for confirming putative interactions suggested by yeast two-hybrid or mass spectrometry proteomics studies, and in addition it is well-suited for mapping interacting regions, assessing the effect of post-translational modifications on protein-protein interactions, and evaluating the impact of mutations identified in patient DNA.

Interactions between proteins are fundamental to all cellular processes. Using Bioluminescence Resonance Energy Transfer, the interaction between a pair of proteins can be monitored in live cells and in real time. Furthermore, the effects of potentially pathogenic mutations can be assessed.

Assays based on Bioluminescence Resonance Energy Transfer (BRET) provide a sensitive and reliable means to monitor protein-protein interactions in live ...

Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing molecules of interest to fluorescent proteins such as GFP to study their expression, localization and function. However, GFP and its derivatives are significantly larger and less photostable than organic fluorophores generally used for in vitro experiments, and this can limit the scope of investigation. 
We recently introduced a straightforward, versatile and high-throughput method based on electroporation, allowing the internalization of biomolecules labeled with organic fluorophores into living microorganisms. Here we describe how to use electroporation to internalize labeled DNA fragments or proteins into Escherichia coli and Saccharomyces cerevisi&#230;, how to quantify the number of internalized molecules using fluorescence microscopy, and how to quantify the viability of electroporated cells. Data can be acquired at the single-cell or single-molecule level using fluorescence or FRET. The possibility of internalizing non-labeled molecules that trigger a physiological observable response in vivo is also presented. Finally, strategies of optimization of the protocol for specific biological systems are discussed.

Studies of biomolecules in vivo are crucial for understanding molecular function in a biological context. Here we describe a novel method allowing the internalization of fluorescent biomolecules, such as DNA or proteins, into living microorganisms. Analysis of in vivo data recorded by fluorescence microscopy is also presented and discussed.

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing ...

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 ...

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 ...

FLIM-FRET Measurements of Protein-Protein Interactions in Live Bacteria.

Summary

Explore More Videos

Chapters in this video

Using In Vitro Fluorescence Resonance Energy Transfer to Study the Dynamics Of Protein Complexes at a Millisecond Time Scale

Measuring TCR-pMHC Binding In Situ using a FRET-based Microscopy Assay

A Protocol for Using Förster Resonance Energy Transfer (FRET)-force Biosensors to Measure Mechanical Forces across the Nuclear LINC Complex

Imaging Protein-protein Interactions in vivo

Measurement of Force-Sensitive Protein Dynamics in Living Cells Using a Combination of Fluorescent Techniques

FRET Microscopy for Real-time Monitoring of Signaling Events in Live Cells Using Unimolecular Biosensors

Investigating Protein-protein Interactions in Live Cells Using Bioluminescence Resonance Energy Transfer

Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation

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

Single-Molecule Measurement of Protein Interaction Dynamics Within Biomolecular Condensates