We would like to thank the Neuroscience Imaging Service at Stanford University School of Medicine for providing equipment and space for this project. This research was supported by intramural funding of the Stanford Cancer Institute and the Gynecologic Oncology Division Stanford as well as GINOP-2.3.2-15-2016-00026, GINOP-2.3.3-15-2016-00030, NN129371, ANN135107 from the National Research, Development and Innovation Office, Hungary.

1. Plasmid construction
<ol>
	<li>For generating the eGFP-mCherry1 fusion probe, use an N1 mammalian cell expression vector (see Table of Materials) with mCherry110 inserted using the restriction sites AgeI and BsrGI.</li>
	<li>Use the following oligonucleotides to amplify eGFP11 without a stop codon as SalI-BamHI fragment: N-terminal primer 5&#39;-AAT TAA CAG TCG ACG ATG GTG AGC AAG GGC GAG G 3&#39; and C-terminal primer 5&#39;-AAT ATA TGG ATC CCG CTT GTA CAG CTC GTC CAT GC 3&#39;.</li>
	<li>Insert this SalI-BamHI fragment into the multiple cloning site of the N1 vector to introduce RNPPV linker (five amino-acid) linker between the green and red fluorescent protein. 
	NOTE: This linker yields a mean FRET efficiency for the GFP-Cherry donor-acceptor pair of about 0.25 -0.3 (Figure 1A). The choice of stiff12 and helical13 linkers of varying lengths to scale measured FRET efficiencies has been discussed elsewhere but is not required for our purpose of the fusion protein. Going forward for simplicity we will call the fluorescent proteins &#39;GFP&#39; and &#39;Cherry&#39;.</li>
</ol>
2. Cell culture and transfection
<ol>
	<li>Use any cell line, e.g., NRK cells, for FRET experiments in media, e.g., Dulbecco&#39;s modified Eagle&#39;s media (DMEM),&#160;without phenol red, to reduce background fluorescence. For the same reason, the usage of phenol red free trypsin is advised.</li>
	<li>Once cells are 80% confluent, detach cells with 1 mL of 0.05% trypsin-EDTA, count the number of cells in suspension using a Neubauer chamber and seed about 10,000 cells per well of an 8-well chambered cover glass; alternatively, from a confluent cell culture grown in T25 flask, use 1 drop of cell suspension from a 2 mL pipette or 3 drops from a 5-mL cell suspension from a confluent culture grown in a T 12.5-cell culture flask.</li>
	<li>Grow cells in 8-well chambers (0.8 cm2/ well) with #1.0 cover glass for fluorescence live-cell microscopy at standard cell culture conditions (37&#176;C and 5% CO2).</li>
	<li>24 h after plating transfect the cells using an appropriate commercially available transfection media (see Table of Materials), with GFP, Cherry, GFP/ Cherry mix (1:1 mix, i.e., 0.8 &#956;g and 0.8 &#956;g GFP and Cherry plasmid DNA), and the GFP-Cherry chimera.
	<ol>
		<li>For transfection, use 5 &#956;L of the transfection reagent in 45 &#956;L of DMEM and 1.6 &#956;g of plasmid DNA. Stir by gently flicking the microcentrifuge tube.</li>
		<li>After 15 min incubation of the mix at room temperature, add 1-2 &#956;L of the transfection reagent mixture to each well of the 8-well chamber slide. Return the chambered cover glass to the incubator.</li>
	</ol>
	</li>
	<li>Let 20 h after transfection elapse before live-cell imaging, to allow for proper fluorescent protein expression, folding and maturation, especially of the red fluorophore.</li>
</ol>
3. FRET Imaging
<ol>
	<li>Image transfected cells in a humidified and heated environmental chamber at 37 &#176;C. To buffer the cell media at physiological pH, use CO2 gas set to 5% flow, or add 20 mM HEPES to render the cell media CO2-independent.</li>
	<li>Use a confocal laser scanning microscope. Set the excitation and emission as follows to optimize the signal and minimize cross-talk.
	<ol>
		<li>Use the 488-nm line of the argon ion laser to excite GFP and the 561-nm diode pumped solid state laser (or 543-nm Helium Neon laser, depending upon available laser lines) to excite Cherry.</li>
		<li>Set the following in the software of a commercial confocal microscope. Set the Dichroic mirror to 488/ 561 by button click using the pull-down menu. Collect fluorescence using 488-nm laser light for excitation in channel 1 through an emission band of 505 - 530 nm (or 505 - 550 nm) and in channel 2 with a long pass filter &#62;585 nm and use the 561-nm laser light for excitation in channel 3 with a long pass filter &#62; 585 nm (type in wavelengths). Band pass filters e.g., 590 - 650 nm or similar can also be used which have the advantage of excluding Raman-scattering.</li>
	</ol>
	</li>
	<li>Excite with the two lasers sequentially and set the imaging mode to Switch after each line so that the excitation of the 512 x 512 pixels image alternates after each line (and not after each frame which would abrogate the ability to detect FRET due to diffusion of the labeled proteins while recording the images with different excitations; button click).</li>
	<li>Set up a mini-time series of three images by button clicks to detect if significant photobleaching occurs, and potentially reduce the laser power. Photobleaching of less than 1% is optimal. High laser intensity can also lead to absorption saturation reducing the apparent FRET efficiency14. Laser power, up to 10-20 &#956;W measured at the objective lens are safe to use.</li>
	<li>First, image cells expressing the GFP-Cherry fusion construct. Set the parameters that define the time-integrated laser intensity per pixel in a confocal image, i.e., the pixel dwell time in microseconds, the acousto-optical tunable filter (AOTF) transmission in percent, and the zoom.
	<ol>
		<li>Image cells using a 63x oil objective and Zoom set to 3x. This provides sufficient magnification and resolution to image cells in its entirety. Aim for a pixel size of 70-80 nm.</li>
		<li>Set Pixel dwell time to 2-4 &#956;s and AOTF transmission for the 488-nm and 561-nm laser such that images have a good signal-to-noise ratio without bleaching and no pixels showing fluorescence intensity saturation. It is advantageous to adjust the laser power of 488 and 561 such that signal levels in channel 1 and channel 3 are similar.</li>
		<li>Set the Photomultiplier (averaging mode) gain to 600-800.</li>
	</ol>
	</li>
	<li>Image with these settings 15-20 cells expressing the GFP-Cherry fusion protein. 15-20 cells provide good statistics while keeping the total time of a FRET measurement session limited to a few hours to help ensure stability of the microscopic set-up.</li>
	<li>Image with the same settings cells expressing GFP, Cherry, GFP and Cherry and non-transfected cells. Search for expressing cells in the green channel or red channel, respectively.</li>
	<li>Then, image 15-20 cells co-expressing proteins of interest coupled to GFP and Cherry, respectively. Searching for expressing cells, avoid long exposure of the cells in order to not bleach the fluorescent proteins. Cherry has a lower photostability than GFP, and bleaching Cherry, the acceptor compromises FRET analysis. 
	NOTE: Absorption and emission spectra of GFP and Cherry is shown in Supplementary Figure 1. After measuring for 5-6 h, it is advisable to repeat imaging a few cells expressing the GFP-Cherry chimera at the end of the imaging session to document that the set-up remained stable and detected FRET efficiencies of the GFP-Cherry fusion protein did not significantly change during the course of an imaging session.</li>
</ol>
4. Image analysis for detecting absolute FRET efficiencies using donor quenching and sensitized emission
NOTE: Here, a practical step-by-step guide as to how to determine FRET efficiency with the use of the attached spreadsheet (Supplementary File 2) is provided. Theory and derivation of the presented equations can be found in detail in previous publications4,15,16,17. With the described settings, the following fluorescence intensities are collected.
<ol>
	<li>Measure the donor signal I1&#160;in channel 1, the donor channel, with 488-nm excitation and an emission band of 505-530 nm. 
	<img alt="Equation 2" src="/files/ftp_upload/62241/62241eq02.jpg" /> 
	where ID is the unquenched donor signal in channel 1 that would be measured in the absence of an acceptor, is the mean FRET efficiency, and B1 the average background signal in channel 1.</li>
	<li>Measure the acceptor signal I3 in channel 3, the acceptor channel, with 561-nm excitation and emission at &#62;585 nm. 
	<img alt="Equation 3" src="/files/ftp_upload/62241/62241eq03.jpg" /> 
	where IA is the acceptor signal and B3&#160;the background in channel 3.</li>
	<li>Measure the FRET signal in channel 2, the transfer channel, with 488-nm excitation and emission at &#62;585 nm. 
	<img alt="Equation 4" src="/files/ftp_upload/62241/62241eq04.jpg" /> 
	Where, the signal in channel 2 is a sum of four different components: (i) ID(1 - E)S1 is the spectral spill over from the quenched donor signal into the &#62;585 detection channel (with the cross-talk factor S1), (ii)&#160;IAS2 is the acceptor signal from the direct excitation by 488-nm light (with the cross-talk factor S2), (iii) IDE&#945; is the sensitized emission of the acceptor by FRET from the excited donor molecule (&#945; will be detailed further in 4.8. - 4.10.), and (iv) B2 is the background signal.</li>
	<li>Measure average background intensities in channels 1, 2, 3 in non-transfected or mock-transfected cells; either is fine with negligible difference. For all cell measurements, use the free-hand tool to delineate regions of interests and avoid perinuclear vesicles with increased autofluorescence. It is important to avoid significant autofluorescence from these perinuclear vesicles.
	<ol>
		<li>Enter the measurements into the columns X, Y, and Z of the provided spreadsheet. Average background intensities in the 3 channels are entered into A2, B2, and C2 of the excel spreadsheet (Supplementary File 2).</li>
	</ol>
	</li>
	<li>Measure average intensities in channels 1, 2, 3 of cells expressing GFP or Cherry alone, and enter the measurements into columns C, D, E, and N, O, P. Respective background intensities are subtracted (in F, G, H, and Q, R, S).</li>
	<li>In order to calculate E, the FRET efficiency, determine the cross-talk factors S1 and S2. The spectral cross-talk factor S1 is calculated from cells expressing only GFP 
	<img alt="Equation 6" src="/files/ftp_upload/62241/62241eq6v5.png" /> 
	in column I. Enter the mean value for S1 into cell D2 on the excel spreadsheet.</li>
	<li>Calculate the spectral cross-talk factor S2 from cells expressing only Cherry 
	<img alt="Equation 7" src="/files/ftp_upload/62241/62241eq07.jpg" /> 
	in column T. Enter the mean for S2 into cell E2 on the excel spreadsheet.</li>
	<li>Ensure that the &#945;&#160;factor relates the signal from any given number of excited GFP molecules in channel 1 to the signal of an equal number of excited Cherry molecules in channel 2, and is defined by 
	<img alt="Equation 8" src="/files/ftp_upload/62241/62241eq08.jpg" />&#160;&#160;&#160;<img alt="Equation 13" src="/files/ftp_upload/62241/62241eq13v5.png" /> 
	where QA and QD are the fluorescence quantum yields of Cherry and GFP; &#951;A and&#160;&#951;D the detection efficiencies of acceptor and donor fluorescence in channels 2 and 1, respectively. 
	NOTE: The &#945;&#160;factor could be determined from two samples expressing known absolute amounts of GFP and Cherry. It is, however, impossible to know the exact amount of GFP and Cherry expressed in a cell. Therefore, we calculated the factor by using cells that express the GFP-Cherry fusion protein. Here, while the absolute amount is still unknown, the ratio of donor and acceptor molecules is known to be one.</li>
	<li>Measure average intensities in channels 1, 2, 3 of cells expressing the GFP-Cherry fusion protein, and enter the measurements into the columns AE, AF, AG. Background intensities are subtracted (in AH, AI, AJ).</li>
	<li>Calculate the &#945;&#160;factor (AJ column) from the fluorescence intensities in channel 1, 2 and 3 of the GFP-Cherry fusion protein as follows: 
	<img alt="Equation 10" src="/files/ftp_upload/62241/62241eq10v6.png" /></li>
	<li>Background-corrected intensities measured in channel 1 (I1 - B1), 2 (I2 - B2) and 3 (I3 - B3), respectively, are measured using the GFP-Cherry chimera. The spectral cross talk factor S1 was determined using cells expressing GFP only (see 4.7.). &#949;D and &#949;A are the extinction coefficients of GFP, the donor, and Cherry, the acceptor, at 488 nm, and can be determined from the literature (&#949;GFP = 53,000 M-1cm-1)18 and the absorption curve of Cherry (&#949;Cherry &#8776; 5560 M-1cm1). The ratio&#160;&#160;<img alt="Equation 11" src="/files/ftp_upload/62241/62241eq11.jpg" />&#160;has been entered in cell G2 of the excel spreadsheet. Enter the mean value for the &#945; factor into J2.</li>
	<li>Use the determined &#945; factor for the calculation of the FRET efficiency, E, as follows (column AK): 
	<img alt="Equation 12" src="/files/ftp_upload/62241/62241eq12v6.png" /></li>
	<li>Alternatively, determine FRET efficiency, E, for the negative controls, i.e., the co-expression of GFP and Cherry and the expression of GFP alone by adding the measurements of channels 1, 2, and 3 in the excel sheet in column AD, AE, and AF under the GFP-Cherry fusion protein measurements. Determine FRET efficiencies between the GFP and Cherry labeled proteins of interest in the same way.</li>
	<li>Determine the unquenched donor intensity, ID as (I1 - B1)/(1 -&#160;E), and the acceptor intensity as IA = I3 - B3; these values are proportional to the expression levels of the tagged proteins.</li>
	<li>Determine the corrected acceptor-to-donor intensity ratio (Q) of the GFP-Cherry fusion protein as follows (column AL): 
	<img alt="Equation 13" src="/files/ftp_upload/62241/62241eq13v5.png" /></li>
	<li>For other co-transfected cells, calculate the acceptor-to-donor molecular ratio NA/ND as follows: 
	<img alt="Equation 14" src="/files/ftp_upload/62241/62241eq14.jpg" /> 
	NOTE: The rationale to determine the NA/ND&#160;ratio and plot the mean cellular FRET efficiency E versus NA/ND, is that one donor molecule can transfer energy to multiple acceptors while an acceptor molecule can only receive energy from one donor at a given time. Even if only one acceptor can interact with a donor because of the stoichiometry of the interaction, an increase of acceptor concentration is expected to increase the fraction of donors in complex with the acceptor because of the law of mass action. Thus, for a fixed (or narrow range of) donor expression, the FRET efficiency should rise with increasing NA/ND. When plotting the FRET efficiency E versus&#160;NA/ND for the co-expression of GFP and Cherry, i.e. the negative probe, however, an increase in NA/ND&#160;should not result in an increase in FRET efficiency (at least at sufficiently low acceptor concentrations where random FRET due to the vicinity of acceptor dyes to donor dyes does not occur).</li>
</ol>

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The authors have nothing to disclose.

The presented protocol details the use of the genetically coupled one-to-one fluorescent protein calibration probe for quantifying FRET using the detection of sensitized emission of the acceptor and quenching of the donor molecule by confocal microscopy. This method can be applied to assess protein interactions in the physiological context of the living cell in different subcellular compartments. Spatial resolution can be further improved by applying the presented algorithm to calculate FRET efficiencies in each pixel of...

An erratum was issued for:&#160;<a href="https://www.jove.com/t/62241">Assessing Protein Interactions in Live-Cells with FRET-Sensitized Emission</a>. The Authors section was updated from:
Gy&#246;rgy V&#225;mosi1 
Sarah Miller2 
Molika Sinha2 
Gabor Mocs&#225;r1 
Malte Renz2 
1Department of Biophysics and Cell Biology, Faculty of Medicine, University of Debrecen 
2Gynecologic Oncology Division, Stanford University School of Medicine
to:
Gy&#246;rgy V&#225;mosi1 
Sarah Miller2 
Molika Sinha2 
Maria Kristha Fernandez2 
Gabor Mocs&#225;r1 
Malte Renz2 
1Department of Biophysics and Cell Biology, Faculty of Medicine, University of Debrecen 
2Gynecologic Oncology Division, Stanford University School of Medicine

An erratum was issued for:&#160;<a href="https://www.jove.com/t/62241">Assessing Protein Interactions in Live-Cells with FRET-Sensitized Emission</a>. The Authors section was updated.

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

Microscopy-based analysis of F&#246;rster Resonance Energy Transfer (FRET) permits assessment of interactions between proteins in live cells. It provides spatial and temporal information, including information on where in the cell and in which subcellular compartment the interaction takes place and if this interaction changes over time.
Theodor F&#246;rster laid the theoretical foundation of FRET in 19481. FRET is a radiationless transfer of energy from an excited donor to an acceptor molecule and depends upon the distance of the molecules and the relative orientation of their transition dipoles as well as the overlap between the donor emission and acceptor absorption spectra. The rate of energy transfer is inversely proportional to the sixth power of the donor-acceptor distance. Thus, FRET can be used to measure molecular proximity in the range of 1-10 nm.
FRET competes with other de-excitation processes of the donor molecule and results in the so-called donor-quenching and sensitized emission of the acceptor. Donor-quenching is a reduction of the number of emitted donor photons, while sensitized emission is an increase in emitted acceptor photons. Many microscopic FRET analyses use fluorescence intensity measurements, including acceptor photobleaching2, donor photobleaching2, or FRET-sensitized photobleaching of the acceptor3.
Here, a step-by-step experimental protocol and mathematical algorithm are presented to quantify FRET using donor quenching and acceptor sensitized emission4,5, a method often referred to as ratiometric FRET. Many protocols on how to approximate sensitized emission have been published, few have quantified the absolute FRET efficiency6,7,8,9. The quantification of FRET efficiencies in the living cell requires determining (i) the crosstalk (spectral spill-over, or bleed-through) of the fluorescent proteins and, also (ii) the detection efficiency of the microscopic setup. While crosstalk can be assessed by imaging cells expressing only one of the fluorophores, the assessment of the relative detection efficiency of the donor and acceptor fluorescence is more complicated. It requires the knowledge of at least the ratio of the number of donor and acceptor molecules giving rise to the measured signals. The number of fluorophores expressed in live cells varies, however, from cell to cell and is unknown. The so-called &#945;&#160;factor characterizes the relative signal strengths from a single excited donor and acceptor molecule. Knowledge of the factor is a prerequisite for quantitative ratiometric FRET measurements in samples with variable acceptor-to-donor molecule ratios as encountered during live-cell imaging with fluorescent proteins. Using a 1-to-1 donor-acceptor fusion protein as a calibration probe permits the determination of the &#945;&#160;factor and also serves as a positive control. This genetically coupled probe is expressed by cells in unknown total amounts but in a fixed and known relative amount of one-to-one. The following protocol lays out how to construct the 1-to-1 probe and how to use it for quantification of FRET efficiency. A spreadsheet that includes all formulae can be found in the supplement and can be used by the readers to enter their own measurements in the respective columns as outlined below.
While the protocol uses the GFP-Cherry donor/ acceptor pair, the presented approach can be performed with any other FRET pair. The Supplementary File 1 provides details on cyan-yellow pairs.

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		<tr>
			<td>0.5% Trypsin-EDTA without phenol red (10x)</td>
			<td>Thermo Fisher Scientific</td>
			<td>15400054</td>
			<td></td>
		</tr>
		<tr>
			<td>Clontech mCherry N1 vector</td>
			<td>Addgene</td>
			<td>3553</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM without phenol red</td>
			<td>Thermo Fisher Scientific</td>
			<td>11054020</td>
			<td></td>
		</tr>
		<tr>
			<td>Fugene 6</td>
			<td>Promega</td>
			<td>E2691</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Thermo Fisher Scientific</td>
			<td>15630080</td>
			<td></td>
		</tr>
		<tr>
			<td>LabTek 8-well chambers #1.0</td>
			<td>Thermo Fisher Scientific</td>
			<td>12565470</td>
			<td></td>
		</tr>
		<tr>
			<td>L-Glutamine (200 mM)</td>
			<td>Thermo Fisher Scientific</td>
			<td>25030081</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Figure 1&#160;shows the images obtained in the donor channel, channel 1 (488, 505-530 nm), the transfer channel, channel 2 (488, &#62;585 nm), and the acceptor channel, channel 3 (561, &#62;585 nm), respectively. Representative images of cells expressing GFP only, Cherry only, co-expressing GFP and Cherry, and expressing the GFP-Cherry fusion protein. The mean cellular FRET efficiencies calculated in NRK cells expressing GFP-Cherry fusion protein (positive co...

Watch this Scientific Journal Video about Assessing Protein Interactions in Live-Cells with FRET-Sensitized Emission at JoVE.com

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

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 ...

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Research

JoVE Journal

Biology

3.0K Views.  University of Debrecen. Förster 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.

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

Staining of Proteins in Gels with Coomassie G-250 without Organic Solvent and Acetic Acid

In classical protein staining protocols using Coomassie Brilliant Blue (CBB), solutions with high contents of toxic and flammable organic solvents (Methanol, Ethanol or 2-Propanol) and acetic acid are used for fixation, staining and destaining of proteins in a gel after SDS-PAGE. To speed up the procedure, heating the staining solution in the microwave oven for a short time is frequently used. This usually results in evaporation of toxic or hazardous Methanol, Ethanol or 2-Propanol and a strong smell of acetic acid in the lab which should be avoided due to safety considerations. In a protocol originally published in two patent applications by E.M. Wondrak (US2001046709 (A1), US6319720 (B1)), an alternative composition of the staining solution is described in which no organic solvent or acid is used. The CBB is dissolved in bidistilled water (60-80mg of CBB G-250 per liter) and 35 mM HCl is added as the only other compound in the staining solution. The CBB staning of the gel is done after SDS-PAGE and thorough washing of the gel in bidistilled water. By heating the gel during the washing and staining steps, the process can be finished faster and no toxic or hazardous compunds are evaporating. The staining of proteins occurs already within 1 minute after heating the gel in staining solution and is fully developed after 15-30 min with a slightly blue background that is destained completely by prolonged washing of the stained gel in bidistilled water, without affecting the stained protein bands.

A short protocol for protein staining with Coomassie Brilliant Blue (CBB) G-250 in polyacrylamide gels is described without using organic solvents or acetic acid as in the classical staining procedures with CBB.

In classical protein staining protocols using Coomassie Brilliant Blue (CBB), solutions with high contents of toxic and flammable organic solvents ...

Using the GELFREE 8100 Fractionation System for Molecular Weight-Based Fractionation with Liquid Phase Recovery

The GELFREE 8100 Fractionation System is a novel protein fractionation system designed to maximize protein recovery during molecular weight based fractionation. The system is comprised of single-use, 8-sample capacity cartridges and a benchtop GELFREE Fractionation Instrument. During separation, a constant voltage is applied between the anode and cathode reservoirs, and each protein mixture is electrophoretically driven from a loading chamber into a specially designed gel column gel. Proteins are concentrated into a tight band in a stacking gel, and separated based on their respective electrophoretic mobilities in a resolving gel. As proteins elute from the column, they are trapped and concentrated in liquid phase in the collection chamber, free of the gel. The instrument is then paused at specific time intervals, and fractions are collected using a pipette. This process is repeated until all desired fractions have been collected. If fewer than 8 samples are run on a cartridge, any unused chambers can be used in subsequent separations.
This novel technology facilitates the quick and simple separation of up to 8 complex protein mixtures simultaneously, and offers several advantages when compared to previously available fractionation methods. This system is capable of fractionating up to 1mg of total protein per channel, for a total of 8mg per cartridge. Intact proteins over a broad mass range are separated on the basis of molecular weight, retaining important physiochemical properties of the analyte. The liquid phase entrapment provides for high recovery while eliminating the need for band or spot cutting, making the fractionation process highly reproducible1.

The accompanying video describes the use of the GELFREE 8100 Fractionation System, which partitions complex protein samples on the basis of molecular weight and recovers the fractions in liquid phase. The video describes how the technology works, how it is used, and provides resultant data, with polyacrylamide gel electrophoresis analysis of fractionated bovine liver homogenate.

The GELFREE 8100 Fractionation System is a novel protein fractionation system designed to maximize protein recovery during molecular weight based ...

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 ...

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, ...

iCLIP - Transcriptome-wide Mapping of Protein-RNA Interactions with Individual Nucleotide Resolution

The unique composition and spatial arrangement of RNA-binding proteins (RBPs) on a transcript guide the diverse aspects of post-transcriptional regulation1. Therefore, an essential step towards understanding transcript regulation at the molecular level is to gain positional information on the binding sites of RBPs2.
Protein-RNA interactions can be studied using biochemical methods, but these approaches do not address RNA binding in its native cellular context. Initial attempts to study protein-RNA complexes in their cellular environment employed affinity purification or immunoprecipitation combined with differential display or microarray analysis (RIP-CHIP)3-5. These approaches were prone to identifying indirect or non-physiological interactions6. In order to increase the specificity and positional resolution, a strategy referred to as CLIP (UV cross-linking and immunoprecipitation) was introduced7,8. CLIP combines UV cross-linking of proteins and RNA molecules with rigorous purification schemes including denaturing polyacrylamide gel electrophoresis. In combination with high-throughput sequencing technologies, CLIP has proven as a powerful tool to study protein-RNA interactions on a genome-wide scale (referred to as HITS-CLIP or CLIP-seq)9,10. Recently, PAR-CLIP was introduced that uses photoreactive ribonucleoside analogs for cross-linking11,12.
Despite the high specificity of the obtained data, CLIP experiments often generate cDNA libraries of limited sequence complexity. This is partly due to the restricted amount of co-purified RNA and the two inefficient RNA ligation reactions required for library preparation. In addition, primer extension assays indicated that many cDNAs truncate prematurely at the crosslinked nucleotide13. Such truncated cDNAs are lost during the standard CLIP library preparation protocol. We recently developed iCLIP (individual-nucleotide resolution CLIP), which captures the truncated cDNAs by replacing one of the inefficient intermolecular RNA ligation steps with a more efficient intramolecular cDNA circularization (Figure 1)14. Importantly, sequencing the truncated cDNAs provides insights into the position of the cross-link site at nucleotide resolution. We successfully applied iCLIP to study hnRNP C particle organization on a genome-wide scale and assess its role in splicing regulation14.

The spatial arrangement of RNA-binding proteins on a transcript is a key determinant of post-transcriptional regulation. Therefore, we developed individual-nucleotide resolution UV crosslinking and immunoprecipitation (iCLIP) that allows precise genome-wide mapping of the binding sites of an RNA-binding protein.

The unique composition and spatial arrangement of RNA-binding proteins (RBPs) on a transcript guide the diverse aspects of post-transcriptional ...

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 ...

The MultiBac Protein Complex Production Platform at the EMBL

Proteomics research revealed the impressive complexity of eukaryotic proteomes in unprecedented detail. It is now a commonly accepted notion that proteins in cells mostly exist not as isolated entities but exert their biological activity in association with many other proteins, in humans ten or more, forming assembly lines in the cell for most if not all vital functions.1,2 Knowledge of the function and architecture of these multiprotein assemblies requires their provision in superior quality and sufficient quantity for detailed analysis. The paucity of many protein complexes in cells, in particular in eukaryotes, prohibits their extraction from native sources, and necessitates recombinant production. The baculovirus expression vector system (BEVS) has proven to be particularly useful for producing eukaryotic proteins, the activity of which often relies on post-translational processing that other commonly used expression systems often cannot support.3 BEVS use a recombinant baculovirus into which the gene of interest was inserted to infect insect cell cultures which in turn produce the protein of choice. MultiBac is a BEVS that has been particularly tailored for the production of eukaryotic protein complexes that contain many subunits.4 A vital prerequisite for efficient production of proteins and their complexes are robust protocols for all steps involved in an expression experiment that ideally can be implemented as standard operating procedures (SOPs) and followed also by non-specialist users with comparative ease. The MultiBac platform at the European Molecular Biology Laboratory (EMBL) uses SOPs for all steps involved in a multiprotein complex expression experiment, starting from insertion of the genes into an engineered baculoviral genome optimized for heterologous protein production properties to small-scale analysis of the protein specimens produced.5-8 The platform is installed in an open-access mode at EMBL Grenoble and has supported many scientists from academia and industry to accelerate protein complex research projects.

Protein complexes catalyze key cellular functions. Detailed functional and structural characterization of many essential complexes requires recombinant production. MultiBac is a baculovirus/insect cell system particularly tailored for expressing eukaryotic proteins and their complexes. MultiBac was implemented as an open-access platform, and standard operating procedures developed to maximize its utility.

Proteomics  research revealed the impressive complexity of eukaryotic proteomes in  unprecedented detail. It is now a commonly accepted notion that ...

Protein-protein Interactions Visualized by Bimolecular Fluorescence Complementation in Tobacco Protoplasts and Leaves

Many proteins interact transiently with other proteins or are integrated into multi-protein complexes to perform their biological function. Bimolecular fluorescence complementation (BiFC) is an in vivo method to monitor such interactions in plant cells. In the presented protocol the investigated candidate proteins are fused to complementary halves of fluorescent proteins and the respective constructs are introduced into plant cells via agrobacterium-mediated transformation. Subsequently, the proteins are transiently expressed in tobacco leaves and the restored fluorescent signals can be detected with a confocal laser scanning microscope in the intact cells. This allows not only visualization of the interaction itself, but also the subcellular localization of the protein complexes can be determined. For this purpose, marker genes containing a fluorescent tag can be coexpressed along with the BiFC constructs, thus visualizing cellular structures such as the endoplasmic reticulum, mitochondria, the Golgi apparatus or the plasma membrane. The fluorescent signal can be monitored either directly in epidermal leaf cells or in single protoplasts, which can be easily isolated from the transformed tobacco leaves. BiFC is ideally suited to study protein-protein interactions in their natural surroundings within the living cell. However, it has to be considered that the expression has to be driven by strong promoters and that the interaction partners are modified due to fusion of the relatively large fluorescence tags, which might interfere with the interaction mechanism. Nevertheless, BiFC is an excellent complementary approach to other commonly applied methods investigating protein-protein interactions, such as coimmunoprecipitation, in vitro pull-down assays or yeast-two-hybrid experiments.

Formation of protein complexes in vivo can be visualized by bimolecular fluorescence complementation. Interaction partners are fused to complementary parts of fluorescent tags and transiently expressed in tobacco leaves, resulting in a reconstituted fluorescent signal upon close proximity of the two proteins.

Many proteins interact transiently with other proteins or are integrated into multi-protein complexes to perform their biological function. Bimolecular ...

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 ...

Probing High-density Functional Protein Microarrays to Detect Protein-protein Interactions

High-density functional protein microarrays containing ~4,200 recombinant yeast proteins are examined for kinase protein-protein interactions using an affinity purified yeast kinase fusion protein containing a V5-epitope tag for read-out. Purified kinase is obtained through culture of a yeast strain optimized for high copy protein production harboring a plasmid containing a Kinase-V5 fusion construct under a GAL inducible promoter. The yeast is grown in restrictive media with a neutral carbon source for 6 hr followed by induction with 2% galactose. Next, the culture is harvested and kinase is purified using standard affinity chromatographic techniques to obtain a highly purified protein kinase for use in the assay. The purified kinase is diluted with kinase buffer to an appropriate range for the assay and the protein microarrays are blocked prior to hybridization with the protein microarray. After the hybridization, the arrays are probed with monoclonal V5 antibody to identify proteins bound by the kinase-V5 protein. Finally, the arrays are scanned using a standard microarray scanner, and data is extracted for downstream informatics analysis1,2 to determine a high confidence set of protein interactions for downstream validation in vivo.

Using protein microarrays containing nearly the entire S. cerevisiae proteome is probed for rapid unbiased interrogation of thousands of protein-protein interactions in parallel. This method can be utilized for protein-small molecule, posttranslational modification, and other assays in high-throughput.

High-density functional protein microarrays containing ~4,200 recombinant yeast proteins are examined for kinase protein-protein interactions using an ...