We wish to acknowledge Dr Scott Henderson for help with confocal microscopy. This research was supported by grants from the National Institutes of Health 1RO1CA127501 to W.A.B as well as pilot project funding from the Massey Cancer Center and
School of Medicine (VCU) to W.A.B. Microscopy was performed at the VCU-Dept. of Neurobiology &amp; Anatomy Microscopy Facility, supported, in part, with funding from NIHNINDS Center core grant 5P30NS047463.

The protocol consists of three major steps. The first, step cloning of your gene of interest into a mammalian expression vector amino-terminally to the monomeric versions of CFP and/or YFP will only be discussed briefly. The second and third steps; transfection of vector DNA into EA.hy926 endothelial cells, and confocal imaging with FRET, are outlined in more depth below.
1. Construction of CFP and YFP Chimeric Receptors
Receptors of interest should be cloned amino-terminally to monomeric enhanced versions of CFP and YFP. Determination of FRET is empirical and is critically dependent upon the length of linker between the transmembrane spanning region and start of the fluorophore1. Therefore, several different lengths of linker must be explored for each new receptor pair under investigation.
<ol>
<li> Into the pcDNA3.1 hygroR or neoR vector containing monomeric C- or YFP (this vector can be obtained from our lab), clone your receptor of interest into the NheI/EcoRI sites. *Note-If necessary, NheI is compatible several other enzymes including SpeI and XbaI.</li>
<li> Transform your ligation reaction into an appropriate endA1 recA competent cell line (we routinely use Mach1). Screen colonies for the presence of insert followed by DNA sequencing to verify the absence of mutations.</li>
<li> Upon obtaining conformation of successful molecular cloning, prepare transfection-grade vector DNA in sterilized water or TE using Qiagen, or similar, DNA kits. Evaluate concentration and purity using UV spectroscopy. Typical yields of DNA should be in the range of 100-200 &mu;g, which is routinely diluted to a working stock of 1 &mu;g/&mu;L.</li>
</ol>
2. Transfection of EA.hy 926 Cells
<ol>
<li> Split EA.hy 926 cells4, grown in complete-DMEM (DMEM+10% Fetal bovine serum+Pen./Strep.) into MatTek coverslip (No. 0) 35 mm dishes with 2 mL medium. You can conservatively obtain ~15 dishes (~90% confluent the following day) from a 15 cm confluent dish. Prepare one dish to evaluate expression of each individual receptor, as well as one dish for each receptor pair.</li>
<li> Allow the cells to incubate overnight in 5%CO2 atmosphere at 37&deg;C. The following day, the cultures should be ~ 70-90% confluent.</li>
<li> On the day of transfection, prepare the nucleolipid complexes by mixing 2 &mu;g of the chimeric receptor vector DNA (2 &mu;L) with 8 &mu;L of Fugene6 in 200 &mu;L of pre-warmed OptiMEM. For receptor pairs, transfect with 1 &mu;g of each receptor, for a total of 2 &mu;g (See also discussion below).</li>
<li> Mix gently by inversion.</li>
<li> Continue to incubate at room temperature for 30 minutes.</li>
<li> During the incubation period, prepare the cells for transfection. Aspirate off fresh media and wash each plate gently and briefly with warm PBS before the addition of 2 mL pre-warmed complete-DMEM without antibiotics (DMEM+FBS). It is important to avoid the presence of Penicillin and Streptomycin, as they are considerably more toxic to cells during treatment with Fugene6.</li>
<li> Add the complex drop wise to cells and incubate for an additional 20-24 hours at 37&deg;C in a 5% CO2 atmosphere.</li>
<li> After 24 hours, carefully aspirate the media from the cells and pipette fresh complete-DMEM with antibiotics. Following transfection, cell viability should remain relatively high, i.e. little to no floating cells should be observed.</li>
<li> Gene expression is typically maximal after an additional 48-60 hours post-transfection. However, imaging is typically feasible in as short as 24 hours. Time course experiments for optimal gene expression should be performed.</li>
</ol>
3. Confocal Imaging and FRET Analysis
Live cell imaging is performed on a Leica TCS-SP2 AOBS confocal laser scanning microscope equipped with blue diode (405 nm), Argon (458, 476, 488, 514 nm). green HeNe (543 nm), orange HeNe (594 nm), and red HeNe (633 nm) lasers, an HCX PI Apo 63x/1.3 n.a. glycerin-immersion objective lens, a motorized XY stage (M&auml;rzh&auml;user), and an environmentally controlled (temperature, humidity, and CO2) stage incubator (PeCon). Experimental controls are critical to eliminate fluorophore cross-talk between the emission channels as well as to evaluate expression issues and non-specific fluorophore multimerization. As such, single fluorophore transfections are utilized for setting up every image session. Negative FRET controls (using known noninteracting receptors) will need to be performed, in order to determine the background FRET that occurs due to over expression.
<ol>
<li> To eliminate crosstalk between the emission channels for CFP and YFP, place the CFP expression control into the stage incubator. Using white light, adjust the Z-plate to focus on to the cells.</li>
<li> Set SP window settings for emission detection of CFP to 465-505 nm and YFP to 525-600 nm.</li>
<li> While monitoring both emission channels, excite the CFP at 458 nm. Using the AOTF, set the laser power to eliminate appreciable bleed over of CFP into the YFP emission channel. Save this setting.</li>
<li> Perform the same control for YFP crosstalk (using the YFP expression control) in the CFP channel by adjusting the excitation power at 514 nm for cells expressing only YFP. Save this setting.</li>
<li> Next place cells co-expressing CFP and YFP into the stage incubator. Using the sequence setting for the Leica confocal software, locate cells expressing similar levels of CFP and YFP with proper membrane localization.</li>
<li> Using the Leica software acceptor photo-bleaching program, set the donor and acceptor settings to your saved CFP and YFP settings, respectably.</li>
<li> Zooming onto the cell, highlight an ROI (region of interest) in which the photo-destruction of YFP is to occur on the cellular membrane and begin the program.</li>
<li> For photo-destruction of the acceptor, adjust the AOTF to 100% at 514 nm and allow the laser bleaching to continue until 70% of the YFP emission has been depleted. To accelerate bleaching, ROIs are typically chosen in the raster axis of the laser (X-axis).</li>
<li> During photo-bleaching, monitor the cellular movement and reject measurements obtained with appreciable movement in the XY-plane, as these measurements can report false FRET efficiencies.</li>
<li> Pre- and post-bleach images are recorded for both donor (CFP) and acceptor (YFP) and FRET efficiency is calculated as: FRETEff=(Dpost-Dpre)/Dpost for all Dpost &gt; Dpre where Dpre and Dpost is the donor intensity before and after photobleaching respectively.</li>
<li> JMP Software (SAS) is used to determine the extent of experimental variability.</li>
</ol>
4. Representative Results
Transfection efficiencies are usually between 30 to 40%. Despite previous findings by others, we have not seen that transfection and expression of one vector favors that of the other. Indeed, we frequently observe exclusive expression of one fluorophore chimera or the other. Typical FRET efficiencies vary among receptor systems. For the Tie receptor system, typical values are 20-28% for epithelial cells and 19-23% in endothelial cells. For negative controls, typical efficiencies are below 2-3%. The extent of variability will decrease considerably with experience.
<img src="/files/ftp_upload/2149/2149fig1.jpg" alt="Figure 1" /> 
Figure 1. Representative images of transfected EA.hy 926 cells. A) DIC image of EA.hy 926 cell monolayer. B) Fluorescence image of cells displayed in (A). C) Overlay of (A) and (B) demonstrating ~20-30% transfection efficiency.
<img src="/files/ftp_upload/2149/2149fig2.jpg" alt="Figure 2" /> 
Figure 2. Representative acceptor photo-bleaching analysis of FRET occurring between CFP and YFP in an EA.hy926 cell. The CFP emission channel (top panels) and YFP emission channel (bottom two panels) were monitored separately prior to and post acceptor photobleaching. Photobleaching experiments were restricted to, and FRET values calculated from, the region within the green box. FRET efficiency is displayed as an absolute range from high (red-1.0) to low (purple-0.0) on a magnified overlay of a CFP/YFP merged image for orientation purposes only.

<ol>
	<li>Kim, M., Carman, C. V., Springer, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bidirectional+transmembrane+signaling+by+cytoplasmic+domain+separation+in+integrins.">Bidirectional transmembrane signaling by cytoplasmic domain separation in integrins.</a> Science. 301, 1720-1725 (2003).</li><li>Zacharias, D. A., Violin, J. D., Newton, A. C., Tsien, R. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Partitioning+of+lipidmodified+monomeric+GFPs+into+membrane+microdomains+of+live+cells.">Partitioning of lipidmodified monomeric GFPs into membrane microdomains of live cells.</a> Science. 296, 913-916 (2002).</li><li>Seegar, T. C. M., Eller, B., Tzvetkova-Robev, D., Kolev, M., Henderson, S. C., Nikolov, D. B., Barton, W. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tie1-Tie2+interactions+mediate+functional+differences+between+angiopoietin+ligands.">Tie1-Tie2 interactions mediate functional differences between angiopoietin ligands.</a> Molecular Cell. 37 (5), 643-655 (2010).</li><li>Edgell, C. J., McDonald, C. C., Graham, J. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Permanent+cell+line+expressing+human+factor+VIII-related+antigen+established+by+hybridization.">Permanent cell line expressing human factor VIII-related antigen established by hybridization.</a> Proc Natl Acad Sci U S A. 80 (12), 3734-3737 (1983).</li></ol>

No conflicts of interest declared.

There are several steps that are critical to success. The most prominent among them is the relative levels of expression between the two chimeric receptors. To circumvent this issue, one may make stable cell lines expressing both proteins of interest, or identify optimal ratios of vector DNA to permit equivalent expression. Similarly, due to non-uniform transfection across a dish, protein levels will seldom be equivalent among cells. Therefore, attention must be paid to distinguish 'high' expressors from 'low'. Those that display 'average' levels of protein typically yield reliable and reproducible FRET efficiencies. One method our lab has pursued to relieve this issue is the use of adeno and lenti-viruses to 'even' gene expression. Furthermore, an alternative method to acceptor photo-bleaching for determining FRET efficiencies is sensitized emission. Although, despite the potential of sensitized emission to monitor single cells in real-time, we have found acceptor photo-bleaching to be more sensitive and more reliable.
Finally, using FRET to monitor protein interactions can be challenging, and requires careful selection of linker lengths for membrane bound receptors. Furthermore, addition of C/YFP often greatly influences protein expression levels and results in aggregation in the endoplasmic reticulum and golgi apparatus. However, for transient, or unstable, interactions, FRET is the ideal methodology to utilize.

<table><tbody><tr><td>DMEM</td><td>Invitrogen</td><td>11960-069</td><td></td></tr><tr><td>Penicillin- Streptomycin</td><td>Invitrogen</td><td>15070-063</td><td></td></tr><tr><td>Fetal Bovine Serum</td><td>Hyclone</td><td>N/A</td><td></td></tr><tr><td>Opti-MEM</td><td>Invitrogen</td><td>11058-021</td><td></td></tr><tr><td>FUGENE 6</td><td>Roche Group</td><td>11814443001</td><td></td></tr><tr><td>Coverslip Dishes</td><td>MatTek Corp.</td><td>P35G014C</td><td></td></tr><tr><td>pcDNA3.1(+)</td><td>Invitrogen</td><td>V790&amp;#8208;20</td><td></td></tr><tr><td>Mach 1 Competent Cells</td><td>Invitrogen</td><td>C862003</td><td></td></tr></tbody></table>

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.

Watch this Scientific Journal Video about Imaging Protein-protein Interactions in vivo at JoVE.com

Imaging Protein-protein Interactions in vivo

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

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, ...

imaging-protein-protein-interactions-in-vivo

Nanyang Technological University

Research

JoVE Journal

Biology

21.3K Views.  Virginia Commonwealth University. This protocol describes how to image protein-protein interactions using a FRET-based proximity assay.

Video: Imaging Protein-protein Interactions in vivo

Visualization of Protein-protein Interaction in Nuclear and Cytoplasmic Fractions by Co-immunoprecipitation and In Situ Proximity Ligation Assay

Protein-protein interactions are involved in thousands of cellular processes and occur in distinct spatial context. Traditionally, co-immunoprecipitation is a popular technique to detect protein-protein interactions. Subsequent Western blot analysis is the most common method to visualize co-immunoprecipitated proteins. Recently, the proximity ligation assay has become a powerful tool to visualize protein-protein interactions in situ and provides the possibility to quantify protein-protein interactions by this method. Similar to conventional immunocytochemistry, the proximity ligation assay technique is also based on the accessibility of primary antibodies to the antigens, but in contrast, proximity ligation assay detects protein-protein interactions with a unique technique involving rolling-circle PCR, while conventional immunocytochemistry only shows co-localization of proteins.
Nuclear factor 90 (NF90) and RNA-binding motif protein 3 (RBM3) have been previously demonstrated as interacting partners. They are predominantly localized in the nucleus, but also migrate into the cytoplasm and regulate signaling pathways in the cytoplasmic compartment. Here, we compared NF90-RBM3 interaction in both the nucleus and the cytoplasm by co-immunoprecipitation and proximity ligation assay. In addition, we discussed the advantages and limitations of these two techniques in visualizing protein-protein interactions in respect to spatial distribution and the properties of protein-protein interactions.

Visualization of Protein-protein Interaction in Nuclear and Cytoplasmic Fractions by Co-immunoprecipitation and In Situ Proximity Ligation Assay

Protein-protein interactions can occur in both the nucleus and the cytoplasm of a cell. To investigate these interactions, traditional co-immunoprecipitation and modern proximity ligation assay are applied. In this study, we compare these two methods to visualize the distribution of NF90-RBM3 interactions in the nucleus and the cytoplasm.

Protein-protein interactions are involved in thousands of cellular processes and occur in distinct spatial context. Traditionally, co-immunoprecipitation ...

Biochemistry

Detection of Heterodimerization of Protein Isoforms Using an in Situ Proximity Ligation Assay

Regulated protein-protein interactions are a guiding principle for many signaling events, and the detection of such events is an important element in understanding how such pathways are organized and how they function. There are many methods to detect protein-protein interactions in cells, but relatively few can be used to detect interactions between endogenous proteins. One such method, the proximity ligation assay (PLA), has several advantages to recommend its use. Compared to other common methods of protein-protein interaction analysis, PLA has relatively high sensitivity and specificity, can be performed with minimal cell manipulation, and, in the protocol described herein, requires only two target-specific antibodies derived from different species (e.g., from mouse and rabbit) and one specialized reagent: a set of secondary antibodies that are covalently linked to specific oligonucleotides that, when brought in close proximity of one another, create an amplifiable platform for in situ PCR or rolling circle amplification. In this presentation, we show how to apply the PLA technique to visualize changes in MST1 and MST2 proximity in fixed cells. The technique described in this manuscript is particularly applicable for the analysis of cell signaling studies.

Detection of Heterodimerization of Protein Isoforms Using an in Situ Proximity Ligation Assay

Here, we show how to use a Proximity Ligation Assay (PLA) to visualize MST1/MST2 heterodimerization in fixed cells with high sensitivity.

Regulated protein-protein interactions are a guiding principle for many signaling events, and the detection of such events is an important element in ...

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

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

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

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

Quantification of Protein Interaction Network Dynamics using Multiplexed Co-Immunoprecipitation

Dynamic protein-protein interactions control cellular behavior, from motility to DNA replication to signal transduction. However, monitoring dynamic interactions among multiple proteins in a protein interaction network is technically difficult. Here, we present a protocol for Quantitative Multiplex Immunoprecipitation (QMI), which allows quantitative assessment of fold changes in protein interactions based on relative fluorescence measurements of Proteins in Shared Complexes detected by Exposed Surface epitopes (PiSCES). In QMI, protein complexes from cell lysates are immunoprecipitated onto microspheres, and then probed with a labeled antibody for a different protein in order to quantify the abundance of PiSCES. Immunoprecipitation antibodies are conjugated to different MagBead spectral regions, which allows a flow cytometer to differentiate multiple parallel immunoprecipitations and simultaneously quantify the amount of probe antibody associated with each. QMI does not require genetic tagging and can be performed using minimal biomaterial compared to other immunoprecipitation methods. QMI can be adapted for any defined group of interacting proteins, and has thus far been used to characterize signaling networks in T cells and neuronal glutamate synapses. Results have led to new hypothesis generation with potential diagnostic and therapeutic applications. This protocol includes instructions to perform QMI, from the initial antibody panel selection through to running assays and analyzing data. The initial assembly of a QMI assay involves screening antibodies to generate a panel, and empirically determining an appropriate lysis buffer. The subsequent reagent preparation includes covalently coupling immunoprecipitation antibodies to MagBeads, and biotinylating probe antibodies so they can be labeled by a streptavidin-conjugated fluorophore. To run the assay, lysate is mixed with MagBeads overnight, and then beads are divided and incubated with different probe antibodies, and then a fluorophore label, and read by flow cytometry. Two statistical tests are performed to identify PiSCES that differ significantly between experimental conditions, and results are visualized using heatmaps or node-edge diagrams.

Quantitative Multiplex Immunoprecipitation (QMI) uses flow cytometry for sensitive detection of differences in the abundance of targeted protein-protein interactions between two samples. QMI can be performed using a small amount of biomaterial, does not require genetically engineered tags, and can be adapted for any previously defined protein interaction network.

Dynamic protein-protein interactions control cellular behavior, from motility to DNA replication to signal transduction. However, monitoring dynamic ...

In-vivo Detection of Protein-protein Interactions on Micro-patterned Surfaces

Unraveling the interaction network of molecules in-vivo is key to understanding the mechanisms that regulate cell function and metabolism. A multitude of methodological options for addressing molecular interactions in cells have been developed, but most of these methods suffer from being rather indirect and therefore hardly quantitative. On the contrary, a few high-end quantitative approaches were introduced, which however are difficult to extend to high throughput. To combine high throughput capabilities with the possibility to extract quantitative information, we recently developed a new concept for identifying protein-protein interactions (Schwarzenbacher et al., 2008). Here, we describe a detailed protocol for the design and the construction of this system which allows for analyzing interactions between a fluorophore-labeled protein ("prey") and a membrane protein ("bait") in-vivo. Cells are plated on micropatterned surfaces functionalized with antibodies against the bait exoplasmic domain. Bait-prey interactions are assayed via the redistribution of the fluorescent prey. The method is characterized by high sensitivity down to the level of single molecules, the capability to detect weak interactions, and high throughput capability, making it applicable as screening tool.

In-vivo Detection of Protein-protein Interactions on Micro-patterned Surfaces

This video shows experiments with subsequent analysis of protein-protein interactions by the use of micro-patterned surfaces. The approach offers the possibility to detect protein interactions in living cells and combines high throughput capabilities with the possibility to extract quantitative information.

Unraveling the interaction network of molecules in-vivo is key to understanding the mechanisms that regulate cell function and metabolism. A multitude of ...

Protein Membrane Overlay Assay: A Protocol to Test Interaction Between Soluble and Insoluble Proteins in vitro

Validating interactions between different proteins is vital for investigation of their biological functions on the molecular level. There are several methods, both in vitro and in vivo, to evaluate protein binding, and at least two methods that complement the shortcomings of each other should be conducted to obtain reliable insights. 
For an in vivo assay, the bimolecular fluorescence complementation (BiFC) assay represents the most popular and least invasive approach that enables to detect protein-protein interaction within living cells, as well as identify the intracellular localization of the interacting proteins 1,2. In this assay, non-fluorescent N- and C-terminal halves of GFP or its variants are fused to tested proteins, and when the two fusion proteins are brought together due to the tested proteins&#x2019; interactions, the fluorescent signal is reconstituted3-6. Because its signal is readily detectable by epifluorescence or confocal microscopy, BiFC has emerged as a powerful tool of choice among cell biologists for studying about protein-protein interactions in living cells 3. This assay, however, can sometimes produce false positive results. For example, the fluorescent signal can be reconstituted by two GFP fragments arranged as far as 7 nm from each other due to close packing in a small subcellular compartment, rather that due to specific interactions7.
Due to these limitations, the results obtained from live cell imaging technologies should be confirmed by an independent approach based on a different principle for detecting protein interactions. Co-immunoprecipitation (Co-IP) or glutathione transferase (GST) pull-down assays represent such alternative methods that are commonly used to analyze protein-protein interactions in vitro. However, iIn these assays, however, the tested proteins must be readily soluble in the buffer that supportsused for the binding reaction. Therefore, specific interactions involving an insoluble protein cannot be assessed by these techniques.
 Here, we illustrate the protocol for the protein membrane overlay binding assay, which circumvents this difficulty. In this technique, interaction between soluble and insoluble proteins can be reliably tested because one of the proteins is immobilized on a membrane matrix. This method, in combination with in vivo experiments, such as BiFC, provides a reliable approach to investigate and characterize interactions faithfully between soluble and insoluble proteins. In this article, binding between Tobacco mosaic virus (TMV) movement protein (MP), which exerts multiple functions during viral cell-to-cell transport8-14, and a recently identified plant cellular interactor, tobacco ankyrin repeat-containing protein (ANK) 15, is demonstrated using this technique.

Protein Membrane Overlay Assay: A Protocol to Test Interaction Between Soluble and Insoluble Proteins in vitro

Testing protein-protein interaction is indispensable for dissection of protein functionality. Here, we introduce an in vitro protein-protein binding assay to probe a membrane-immobilized protein with a soluble protein. This assay provides a reliable method to test interaction between an insoluble protein and a protein in solution.

Validating interactions between different proteins is vital for investigation of their biological functions on the molecular level.  There are several ...

In Vivo Two-Color 2-Photon Imaging of Genetically-Tagged Reporter Cells in the Skin

Fibroblasts are mesenchymal cells that change their morphology upon activation, ultimately influencing the microenvironment of the tissue they are located in. Although traditional imaging techniques are useful in identifying protein interactions and morphology in fixed tissue, they are not able to give insight as to how quickly cells are able to bind and uptake proteins, and once activated how their morphology changes in vivo. In the present study, we ask 2 major questions: 1) what is the time-course of fibroblast activation via toll-like receptor-4 (TLR4) and lipopolysaccharide (LPS) interaction and 2) how do these cells behave once activated? Using 2-photon imaging, we have developed a novel technique to assess the ability of LPS-FITC to bind to its cognate receptor, TLR4, expressed on peripheral fibroblasts in the genetic reporter mouse line; FSP1cre; tdTomatolox-stop-lox&#160;in vivo. This unique approach allows researchers to create in-depth, time-lapse videos and/or pictures of proteins interacting with live cells that allows one to have an increased level of granularity in understanding how proteins can alter cellular behavior.

Morphological changes occur in immune responsive fibroblast cells following activation and promote alterations in cellular recruitment. Utilizing 2-photon imaging in conjunction with a genetically engineered Fibroblast-specific protein 1 (FSP1)-cre; tdTomato floxed-stop-floxed (TB/TB) mouse line and green fluorescently tagged lipopolysaccharide-FITC, we can illustrate highly specific uptake of lipopolysaccharide in dermal fibroblasts and morphological changes in vivo.

Fibroblasts are mesenchymal cells that change their morphology upon activation, ultimately influencing the microenvironment of the tissue they are located ...

Bioengineering

4D Imaging of Protein Aggregation in Live Cells

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental conditions and an intracellular environment that is tightly packed, sticky, and hazardous to protein stability1. The ability to dynamically balance protein production, folding and degradation demands highly-specialized quality control machinery, whose absolute necessity is observed best when it malfunctions. Diseases such as ALS, Alzheimer's, Parkinson's, and certain forms of Cystic Fibrosis have a direct link to protein folding quality control components2, and therefore future therapeutic development requires a basic understanding of underlying processes. Our experimental challenge is to understand how cells integrate damage signals and mount responses that are tailored to diverse circumstances.
The primary reason why protein misfolding represents an existential threat to the cell is the propensity of incorrectly folded proteins to aggregate, thus causing a global perturbation of the crowded and delicate intracellular folding environment1. The folding health, or "proteostasis," of the cellular proteome is maintained, even under the duress of aging, stress and oxidative damage, by the coordinated action of different mechanistic units in an elaborate quality control system3,4. A specialized machinery of molecular chaperones can bind non-native polypeptides and promote their folding into the native state1, target them for degradation by the ubiquitin-proteasome system5, or direct them to protective aggregation inclusions6-9.
In eukaryotes, the cytosolic aggregation quality control load is partitioned between two compartments8-10: the juxtanuclear quality control compartment (JUNQ) and the insoluble protein deposit (IPOD) (Figure 1 - model). Proteins that are ubiquitinated by the protein folding quality control machinery are delivered to the JUNQ, where they are processed for degradation by the proteasome. Misfolded proteins that are not ubiquitinated are diverted to the IPOD, where they are actively aggregated in a protective compartment.
Up until this point, the methodological paradigm of live-cell fluorescence microscopy has largely been to label proteins and track their locations in the cell at specific time-points and usually in two dimensions. As new technologies have begun to grant experimenters unprecedented access to the submicron scale in living cells, the dynamic architecture of the cytosol has come into view as a challenging new frontier for experimental characterization. We present a method for rapidly monitoring the 3D spatial distributions of multiple fluorescently labeled proteins in the yeast cytosol over time. 3D timelapse (4D imaging) is not merely a technical challenge; rather, it also facilitates a dramatic shift in the conceptual framework used to analyze cellular structure.
We utilize a cytosolic folding sensor protein in live yeast to visualize distinct fates for misfolded proteins in cellular aggregation quality control, using rapid 4D fluorescent imaging. The temperature sensitive mutant of the Ubc9 protein10-12 (Ubc9ts) is extremely effective both as a sensor of cellular proteostasis, and a physiological model for tracking aggregation quality control. As with most ts proteins, Ubc9ts is fully folded and functional at permissive temperatures due to active cellular chaperones. Above 30 &deg;C, or when the cell faces misfolding stress, Ubc9ts misfolds and follows the fate of a native globular protein that has been misfolded due to mutation, heat denaturation, or oxidative damage. By fusing it to GFP or other fluorophores, it can be tracked in 3D as it forms Stress Foci, or is directed to JUNQ or IPOD.

Cellular viability depends on timely and efficient management of protein misfolding. Here we describe a method for visualizing the different potential fates of a misfolded protein: refolding, degradation, or sequestration in inclusions. We demonstrate the use of a folding sensor, Ubc9ts, for monitoring proteostasis and aggregation quality control in live cells using 4D microscopy.

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental ...

Luciferase Complementation Imaging Assay in Nicotiana benthamiana Leaves for Transiently Determining Protein-protein Interaction Dynamics

Protein-protein interactions are fundamental mechanisms for relaying signal transduction in most cellular processes; therefore, identification of novel protein-protein interaction pairs and monitoring protein interaction dynamics are of particular interest for revealing how plants respond to environmental factors and/or developmental signals. A plethora of approaches have been developed to examine protein-protein interactions, either in vitro or in vivo. Among them, the recently established luciferase complementation imaging (LCI) assay is the simplest and fastest method for demonstrating in vivo protein-protein interactions. In this assay, protein A or protein B is fused with the amino-terminal or carboxyl-terminal half of luciferase, respectively. When protein A interacts with protein B, the two halves of luciferase will be reconstituted to form a functional and active luciferase enzyme. Luciferase activity can be recorded with a luminometer or CCD-camera. Compared with other approaches, the LCI assay shows protein-protein interactions both qualitatively and quantitatively. Agrobacterium infiltration in Nicotiana benthamiana leaves is a widely used system for transient protein expression. With the combination of LCI and transient expression, these approaches show that the physical interaction between COP1 and SPA1 was gradually reduced after jasmonate treatment.

Luciferase Complementation Imaging Assay in Nicotiana benthamiana Leaves for Transiently Determining Protein-protein Interaction Dynamics

This manuscript describes an easy and rapid experimental procedure for determining protein-protein interactions based on the measurement of luciferase activity.

Protein-protein interactions are fundamental mechanisms for relaying signal transduction in most cellular processes; therefore, identification of novel ...

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 ...

Imaging Protein-protein Interactions in vivo

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Visualization of Protein-protein Interaction in Nuclear and Cytoplasmic Fractions by Co-immunoprecipitation and In Situ Proximity Ligation Assay

Detection of Heterodimerization of Protein Isoforms Using an in Situ Proximity Ligation Assay

Evaluation of Protein–Protein Interactions using an On-Membrane Digestion Technique

Quantification of Protein Interaction Network Dynamics using Multiplexed Co-Immunoprecipitation

In-vivo Detection of Protein-protein Interactions on Micro-patterned Surfaces

Protein Membrane Overlay Assay: A Protocol to Test Interaction Between Soluble and Insoluble Proteins in vitro

In Vivo Two-Color 2-Photon Imaging of Genetically-Tagged Reporter Cells in the Skin

4D Imaging of Protein Aggregation in Live Cells

Luciferase Complementation Imaging Assay in Nicotiana benthamiana Leaves for Transiently Determining Protein-protein Interaction Dynamics

Single-Molecule Measurement of Protein Interaction Dynamics Within Biomolecular Condensates