We thank C.F. Kaminski for the kind gift of the plasmid encoding the CFP-18aa-YFP construct, A. Klemmer for technical assistance, and all current and former members of the Stauber laboratory who contributed to the development of this method.

1. Preparation of buffer and reagents
<ol>
	<li>Prepare medium and isotonic, hypotonic, and hypertonic buffers and measure the osmolarity of the buffers with an osmometer to ensure that the osmolarity (Osm) is in the expected range with only an acceptable deviation.
	<ol>
		<li>Cell culture media: Prepare Dulbecco&#180;s Modified Eagle Medium (DMEM) supplemented with 10% Fetal Bovine Serum (FBS) and 1% penicillin/streptomycin</li>
		<li>Isotonic imaging buffer (~340 mOsm): Prepare the isotonic imaging buffer by mixing 150 mM NaCl, 6 mM KCl, 1 mM MgCl2, 1.5 mM CaCl2, 10 mM D(+)-Glucose and 10 mM 4-(2-Hydroxy Ethyl)-1-Piperazine Ethane Sulfonic acid (HEPES).</li>
		<li>Hypotonic imaging buffer (~250 mOsm): Prepare the hypotonic imaging buffer by mixing 105 mM NaCl, 6 mM KCl, 1 mM MgCl2, 1.5 mM CaCl2, 10 mM D(+)-Glucose and 10 mM HEPES.</li>
		<li>Hypertonic imaging buffer (~500 mOsm): Prepare the hypertonic imaging buffer by mixing 160 mM D(-)-Mannitol, 150 mM NaCl, 6 mM KCl, 1 mM MgCl2, 1.5 mM CaCl2, 10 mM D(+)-Glucose and 10 mM HEPES. 
		NOTE: Buffers may be supplemented with specific drugs depending on what will be investigated.</li>
	</ol>
	</li>
</ol>
2. Growth of adherent cells on glass bottom dishes
<ol>
	<li>Prepare at least one dish for transfection only with the protein of interest (POI) fused to either the donor or the acceptor fluorophore and at least one dish for transfection with the donor and acceptor pair (Table 1 and Figure 2). Use glass-bottom dishes to allow for fluorescence microscopy. Depending on the cell type, pretreat the surface by special cleaning or coating it with polylysine. 
	NOTE: Here, FRET donor mCerulean3 (mCer)54 and FRET acceptor mVenus (mVen)55 are used as FRET pair, but other combinations of fluorophores are also applicable. Sample expressing the POI fused to the donor or acceptor alone are used to determine the correction factors &#946; and &#947; (Table 1) necessary to correct for donor bleed through in the acceptor channel and cross excitation of the acceptor by the donor excitation wavelength (Figure 2Ac, Bf).</li>
	<li>The day before transfection, seed 1 x 105 HeLa cells in 2 mL of cell culture media on 35-mm dishes with a glass bottom suitable for fluorescence microscopy. Culture cells overnight at 37 &#176;C and 5% CO2 atmosphere in a cell culture incubator. 
	NOTE: Volumes of reagents can be adapted to the surface area of other cell culture dishes/flask with a different size. The cell number can be adapted to a density suitable for different cell lines and experimental approaches.
	<ol>
		<li>Aspirate the cell culture supernatant from the monolayer of adherent cells cultured on T75 cell culture flasks using a vacuum pump.</li>
		<li>Wash the cells by adding 10 mL of Dulbecco&#39;s Phosphate Buffered Saline (DPBS) to the cell culture flask and aspirating the solution afterward with a vacuum pump.</li>
		<li>Cover the cells completely with 1mL of 0,05% Trypsin-EDTA and incubate the cells for 2 min at 37 &#176;C and 5% CO2 atmosphere in a cell culture incubator.</li>
		<li>Add 9 mL of cell culture media and suspend the cells by pipetting.</li>
		<li>Transfer the cell suspension into a 15 mL tube and centrifuge at 700 x g for 3 min at room temperature (RT).</li>
		<li>Aspirate the supernatant with a vacuum pump and resuspend the cell pellet in 10 mL of cell culture media.</li>
		<li>Mix equal volumes of cell suspension and 0.4% trypan blue in a tube and add 10 &#181;L to a counting slide. Place the slide into the automated cell counter and start counting using the appropriate program for cell counting.</li>
		<li>To calculate the volume of cell suspension needed for each dish, divide the cell number needed per dish (1 x 105) by the cell number per milliliter obtained from the cell counter.</li>
		<li>Prepare a cell suspension in a suitable tube containing the number of cells needed per dish in the volume needed per dish (2 mL) plus the amount for one dish extra.</li>
		<li>Mix cell suspension at least 20 times by inverting the tube and pipette 2 mL of the cell suspension in each dish.</li>
		<li>To ensure a more even cell distribution, leave the dishes for 30 min at RT before placing them in the cell culture incubator.</li>
	</ol>
	</li>
</ol>
3. Cell transfection
NOTE: Here, FuGENE was used as a transfection reagent. Other transfection reagents and methods are also applicable. Optimal ratios of plasmid DNA (pDNA) to transfection reagents and time for optimal expression for each POI and cell model need to be assessed empirically. Here, 2 &#181;g of total pDNA per 35 mm dish was used. FRET donor and acceptor constructs were used in a 1:1 ratio, and the pDNA-to-transfection reagent ratio was 1:4 (Table 1).
<ol>
	<li>Before transfection, replace the cell culture supernatant with fresh 2 mL of pre-warmed media.</li>
	<li>Dilute pDNA in Opti-Minimal Essential Medium (MEM) with a final volume of 100&#160;&#181;L (=pDNA solution).</li>
	<li>Dilute the transfection reagent in Opti-MEM with a final volume of 100&#160;&#181;L (=reagent solution). 
	NOTE: For a more uniform transfection, prepare a master mix for the pDNA and reagent solution.</li>
	<li>Mix pDNA and reagent solution well.</li>
	<li>Add the pDNA solution into the reagent solution (=transfection solution).</li>
	<li>Mix the transfection solution well.</li>
	<li>Incubate the transfection solution for 15 min at RT.</li>
	<li>Add the transfection solution dropwise in a spiral motion to the dish.</li>
	<li>Mix by moving the dish 5 times horizontally and vertically on the surface of the bench.</li>
	<li>Culture cells overnight at 37 &#176;C and 5% CO2 atmosphere in a cell culture incubator.</li>
</ol>
&#160;&#160;
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Condition</td>
			<td>Construct (s)</td>
			<td>Sample</td>
			<td>Used for</td>
		</tr>
		<tr>
			<td>1</td>
			<td>LRRC8A-mCerulean</td>
			<td>Donor construct only</td>
			<td>determine correction factor &#946;</td>
		</tr>
		<tr>
			<td>2</td>
			<td>LRRC8E-mVenus</td>
			<td>Acceptor construct only</td>
			<td>determine correction factor &#947;</td>
		</tr>
		<tr>
			<td>3</td>
			<td>LRRC8A-mCerulean and LRRC8E-mVenus</td>
			<td>FRET pair</td>
			<td>SE-FRET quantification</td>
		</tr>
	</tbody>
</table>
Table 1: Example of conditions necessary for a typical SE-FRET experiment to measure LRRC8/VRAC activity of a channel composed of the LRRC8A subunit fused to the donor (mCerulean3) and LRRC8E subunit fused to the acceptor (mVenus) fluorophore.
4. Image acquisition for correction factor determination
NOTE: There is a bleed-through of the donor emission into the detected emission of the acceptor during FRET. Moreover, there is cross-excitation of the acceptor fluorophore by the donor excitation wavelength. These processes must be compensated for during the calculation of the SE-FRET. To this end, correction factors are determined in cells expressing only the FRET donor or acceptor 24 h after transfection. Here, the imaging was carried out on a Leica THUNDER Imager equipped with a Leica LED8 lamp, the filter cube CYR71010, an HC PL APO 63x/1.40 OIL objective, long pass filter for 460/80 and 553/70 and a Leica DFC9000GTC camera. Experiments were performed without environmental control but in the presence of HEPES in the imaging buffers to stabilize the pH. For long-term observation/measurements, it is advised to use an environmental control system. For analysis, SE-FRET is calculated from the captured raw images. This can be done simultaneously during acquisition or afterward. Here, the Leica LAS X software with the SE-FRET plugin was used to simplify the experimental procedure regarding the calculation of the correction factors and visualizing the SE-FRET value changes in real-time during image acquisition. For post-acquisition, correction factors and SE-FRET can be determined with other software packages (e.g., FIJI) after raw data acquisition according to the protocol provided below.
<ol>
	<li>Before image acquisition of single transfected cells to calculate the correction factors, use one sample expressing the FRET pair to set up the microscopy setting for all channels (donor excitation/donor emission DD, donor/acceptor DA, and acceptor/acceptor AA; see Table 1 and Figure 2C and Figure 3A). 
	NOTE: For SE-FRET measurements with mCerulean3- and mVenus-tagged VRAC subunits transfected as described above, the following parameters were used: 8x8 pixel binning, 100ms exposure time, and 10% LED lamp intensity.</li>
	<li>Take the sample expressing the donor construct only (Figure 2A), aspirate the cell culture media, and wash the cells three times with 2 mL of isotonic buffer.</li>
	<li>Add 3 mL of isotonic buffer and place the sample on the microscope stage.</li>
	<li>Find a field of view (FOV) with at least one cell expressing the donor construct.</li>
	<li>Image all the channels (DD, DA, and AA see Table 2 and Figure 2A).</li>
	<li>Draw a region of interest (ROI) around the cell/cells and measure the mean intensity of DA (= IDA; Figure 2Ac) and DD (= IDD; Figure 2Aa).</li>
	<li>For background subtraction, draw an ROI in the DA and DD channels where only the background signal is found and measure the mean intensity (IBDA and IBDD).</li>
	<li>Subtract the mean intensity of the background ROI from the mean intensity measured in the cell ROI of the corresponding channel (<img alt="Equation 1" src="/files/ftp_upload/66953/66953eq01.jpg" style="margin: auto;" /> and <img alt="Equation 2" src="/files/ftp_upload/66953/66953eq02.jpg" style="margin: auto;" />).</li>
	<li>Repeat steps 4.2- 4.5 with the sample expressing only the acceptor construct (Figure 2B).</li>
	<li>Draw an ROI around the cell(s) and measure the mean intensity of DA (= IDA; Figure 2Bf) and AA (= IAA; Figure 2Be).</li>
	<li>For background subtraction, draw an ROI in the DA and AA channels where only the background signal is found and measure the mean intensities (IBDA and IBAA).</li>
	<li>Subtract the mean intensity of the background ROI from the mean intensity measured in the cell ROI of the corresponding channel (<img alt="Equation 1" src="/files/ftp_upload/66953/66953eq01.jpg" style="margin: auto;" /> and <img alt="Equation 3" src="/files/ftp_upload/66953/66953eq03.jpg" style="margin: auto;" />).</li>
	<li>Use the values determined for IDA*, IDD*, and IAA* to calculate the correction factors <img alt="Equation 4" src="/files/ftp_upload/66953/66953eq04.jpg" style="margin: auto;" /> and <img alt="Equation 5" src="/files/ftp_upload/66953/66953eq05.jpg" style="margin: auto;" />, 
	&#8203;Whereby the correction factor &#946; is used to compensate for the bleed-through of the donor emission into the DA channel. The correction factor &#947; is used to compensate for the cross-excitation of the acceptor fluorophore by the donor excitation wavelength in the DA channel.</li>
</ol>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Excitation of</td>
			<td>Emission of</td>
			<td>Channel name</td>
			<td>LED line</td>
			<td>Filter cube</td>
			<td>Long-pass filter</td>
		</tr>
		<tr>
			<td>Donor</td>
			<td>Donor</td>
			<td>DD</td>
			<td>440 nm</td>
			<td>CYR71010</td>
			<td>460/80 nm</td>
		</tr>
		<tr>
			<td>Donor</td>
			<td>Acceptor</td>
			<td>DA</td>
			<td>440 nm</td>
			<td>CYR71010</td>
			<td>535/70 nm</td>
		</tr>
		<tr>
			<td>Acceptor</td>
			<td>Acceptor</td>
			<td>AA</td>
			<td>510 nm</td>
			<td>CYR71010</td>
			<td>535/70 nm</td>
		</tr>
	</tbody>
</table>
Table 2: Summary of channels required for SE-FRET experiments.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66953/66953fig02.jpg" /> 
Figure 2: Representative fluorescence images of samples used to calculate the correction factors &#946; and &#947; necessary to determine the VRAC activity of a channel composed of LRRC8A subunit fused to the donor mCerulean3 (mCer) and LRRC8E subunit fused to the acceptor mVenus (mVen) fluorophore by SE-FRET measurements. (A,B) Detection of the donor/donor DD, acceptor/acceptor AA, and donor/acceptor DA channel in HeLa cells only expressing the donor (A) LRRC8A-mCer or the acceptor (B) LRRC8E-mVen. (C) Detection of DD, AA, and DA channels in HeLa cells co-transfected with the donor and acceptor pair LRRC8A-mCer and LRRC8E-mVen. Panels a-i show images taken in the donor detection channel (excitation of the donor and detection of the donor signal; DD; a, d and g), the acceptor detection channel (excitation of the acceptor and detection of the acceptor signal; AA; b, e, and h ) and the FRET signal detection channel (excitation of the donor and detection of the acceptor signal; DA; c, f, and i). Panel j is the overlay of panels g and h. DD channel is shown in green and AA channel in magenta. Scale bar = 10 &#181;m. <a href="https://www.jove.com/files/ftp_upload/66953/66953fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
5. Time-lapse imaging for SE-FRET quantification
<ol>
	<li>Take the sample expressing the donor and acceptor construct (Table 1, Figure 2C and Figure 3A), aspirate cell culture media, and wash the cells three times with 2 mL of isotonic buffer 24 h after transfection.</li>
	<li>Add 3 mL of isotonic buffer and place the sample on the microscopy stage.</li>
	<li>For later aspiration of the isotonic buffer, fixate and adjust a hose cannula so the cannula tip reaches the bottom of the dish.</li>
	<li>For adding buffers, fixate and adjust tubings so that the buffer driven by gravity flow can drop into the dish. 
	NOTE: Here, a valve controlled gravity perfusion system with four magnetic valves to control the buffer flow was used, but other methods are also applicable.</li>
	<li>Find a FOV with at least one cell expressing the donor and acceptor construct simultaneously.</li>
	<li>Set up a time-lapse experiment for the channels DD, DA, and AA with an interval of 10 s and a duration to cover all conditions of the stimulation sequence, e.g., 12 cycles of baseline followed by 15 cycles per condition (Figure 3). 
	NOTE: The interval and the number of cycles can be adopted according to the experimental needs but should always include a baseline measurement for normalization for better visualization and comparison of the data.</li>
	<li>After baseline measurement, wash the sample in the buffer for the first condition.
	<ol>
		<li>Aspirate the isotonic buffer via the hose cannula, applying a vacuum with a syringe.</li>
		<li>Add 3 mL of the buffer of the next condition by gravity flow.</li>
		<li>Aspirate the buffer and add again 3 mL of the same buffer.</li>
		<li>Repeat step 5.7.3 once more.</li>
	</ol>
	</li>
	<li>After measurement of the first condition, wash the sample in the buffer for the next condition (steps 5.7.1-5.7.4).</li>
	<li>Repeat steps 5.5-5.8 until all conditions are captured.</li>
	<li>For SE-FRET quantification, draw an ROI around the cell/cells and measure the mean intensity in the DD = (IDD), DA (IDA), and AA (IAA) channel for all images (time points) in the time series (Figure 3A, B).</li>
	<li>For background subtraction for the FRET signal, draw an ROI in the DA channel where only the background signal is found and measure the mean intensity (IBDA).</li>
	<li>Subtract the mean intensity of the background ROI from the mean intensity measured in the cell ROI of the DA channel (<img alt="Equation 1" src="/files/ftp_upload/66953/66953eq01.jpg" style="margin: auto;" />).</li>
	<li>Use the values determined to calculate the mean SE-FRET values for each ROI and all time points <img alt="Equation 6" src="/files/ftp_upload/66953/66953eq06.jpg" style="margin: auto;" />.</li>
	<li>For better comparability of different conditions and visualization of the SE-FRET values, use the mean of the baseline values of each ROI to normalize all time points of the corresponding ROI.</li>
	<li>Plot the normalized SE-FRET values over time (Figure 3B).</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66953/66953fig03.jpg" /> 
Figure 3: Representative fluorescence images and SE-FRET quantification. (A) Representative fluorescence images and the apparent SE-FRET of the first time point of a time laps experiment to measure the VRAC activity by SE-FRET of a channel composed of LRRC8A and LRRC8E subunits depending on the tonicity. Scale bar = 10 &#181;m. The same cells are shown in Figure 2C. Panels a-i show the detection of the DD, AA, and DA channels and the calculated apparent SE-FRET. White outlines represent the ROIs (cells i-iii in panel d) used to measure the mean signal intensities in DD, AA and DA and the apparent SE-FRET image. (B) Quantification of the SE-FRET values over time. Sequence of conditions was 12 cycles of isotonic imaging buffer (baseline) followed by 15 cycles of hypotonic and 15 cycles of hypertonic imaging buffer. The raw mean SE-FRET value of each ROI (cells i-iii) and time point was normalized to the mean of the baseline (isotonic) value for the corresponding ROI. <a href="https://www.jove.com/files/ftp_upload/66953/66953fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

FRET-Based Method to Monitor LRRC8 Channel Activity Without Electrophysiology

The authors have no conflicts of interest to disclose.

FRET microscopy is a well-established, widely used technique to study the interaction between proteins. Hence, FRET-based methods can be applied in many laboratories of variable expertise. Conformational rearrangements during gating have been monitored for a broad range of ion channels using FRET-based assays (for examples, see references34,50,51,52,53,60,61,62,63,64,65,66,67), in some cases combined with electrophysiology in patch-clamp fluorometry68,69,70,71. FRET can be used to study the structure-function relationships of these ion channels or to monitor their activity independent of ion transport. The method presented here can have clear advantages over electrophysiology as it allows monitoring of the activity of LRRC8/VRAC channels in situ.
Critical steps in the protocol include plating the cells to reach optimal confluency for transfection and imaging, which ideally facilitates easy cell distinction for later analysis. Effective co-transfection of the different subunits is crucial for correct subcellular localization; for example, an excess of the non-LRRC8A subunit will lead to enhanced endoplasmic reticulum (ER) localization4. Therefore, plasmid ratios may need to be adjusted. Depending on the system, newly generated FRET pairs should be verified, e.g., by acceptor bleaching. Binning and exposure time must be balanced against each other to enable optimal temporal and spatial resolution for the research question. Binning enables shorter exposure times and hence reduces potential bleaching of the FRET sensor while decreasing spatial resolution. Therefore, if the experimental setup requires, e.g., subcellular discrimination of LRRC8/VRAC activity, binning should be avoided. The research question equally determines the number and interval of cycles in a time-lapse series. The interval is only relevant if the kinetics of the FRET changes (and hence LRRC8/VRAC activation/inactivation) are required; otherwise, simple &#34;before-and-after&#34; recordings can also be performed. The length of the experiment depends on the physiological process. Ideally, LRRC8/VRAC activity upon stimuli should be monitored until SE-FRET has stabilized. These factors can be determined in pilot experiments. Correction factors to calculate the real SE-FRET signal must be determined for all conditions. Incorrectly determined correction factors may lead to an over- or underestimation of the SE-FRET intensities. Lastly, after establishing a stable baseline, the time interval between images has to be short enough to capture the physiological process of interest.
The method bears some limitations. One of them is that changes in inter-LRRC8 FRET intensities while reflecting movements of the LRRDs, do not necessarily correspond to ion or osmolyte transport through the pore. This is clear from the FRET changes observed with LRRC8A homomers45 despite their minimal currents4,32,72. Pore blockers of LRRC8/VRAC channels may not affect the FRET signal, rendering this method unsuitable for the search for specific channel modulators. Moreover, the expression levels of the overexpressed LRRC8 proteins could affect the physiological processes that are observed, especially as the C-terminally tagged LRRC8 proteins display basal activity14.
An aspect that can be considered a limitation or an advantage depending on the particular research question is that in this method, only the ectopically expressed LRRC8 subunits are selectively measured. So, background levels of endogenous proteins hardly interfere with the measurements. On the other hand, the overexpressed proteins may not behave like the endogenous LRRC8 channels with potentially different subunit composition and stoichiometry. For example, various stimuli such as oxidation may have opposing modulatory effects on differently composed LRRC8 channels36. By altering the ratios between co-expressed subunits, their stoichiometry, and overall ion conductance can be adjusted14,73, but their native composition, with likely often more than two paralogs within one complex21, is not clear and may vary between cell types74,75,76. Furthermore, the fusion of fluorescent proteins to the cytosolic C-termini of LRRC8 proteins was shown to increase basal LRRC8/VRAC channel activity in Xenopus oocytes14, likely because the large tags modulate the conformation of the LRRDs, which may govern channel opening14,44,45. Therefore, the size of the fluorescent proteins, the linker, and their orientation may not only affect FRET efficiency but also channel activity. However, importantly, VRAC channels of LRRC8 proteins fused with fluorescent proteins remained responsive to hypotonic stimulation14, enabling their use as FRET sensors45.
Advantages of this non-invasive method to monitor LRRC8/VRAC channel activity by light microscopy compared to other methods comprise: (i) It allows observing LRRC8/VRAC within cells or compartments typically inaccessible for electrophysiology. This includes intracellular organelles on which LRRC8 complexes can be found or targeted to45,77,78. (ii) The cytosolic composition remains unaltered by the method, whereas during whole-cell patch-clamp measurements, the cytosol is largely replaced by pipette solution, which may affect signaling pathways as observed with phorbol-12-myristate-13-acetate (PMA)-induced LRRC8/VRAC activation45. (iii) It offers the possibility to observe LRRC8/VRAC activation with subcellular resolution, such as distinguishing activity at the leading and trailing edges during cell migration, where -restricted to confined spaces- VRAC has been implicated79,80. (iv) It enables continuous monitoring of LRRC8/VRAC activity during extended physiological processes such as myocyte differentiation56.
While there are limitations and challenges with this method, it holds promise for further exploration, including potential applications in animal models. In combination with other methods to study this ion and osmolyte channel family, this FRET-based assay may contribute significantly to unraveling activation mechanisms and exploring the diverse physiological functions of LRRC8 channels in their native environments.

Ion channels comprised of heteromers of leucine-rich repeat containing 8 (LRRC8) family proteins are found throughout vertebrate cells, participating in a wide range of physiological functions1,2. These LRRC8 channels, first identified as volume-regulated anion channels (VRACs) or volume-sensitive outwardly rectifying channels (VSOR), play a crucial role in cellular regulatory volume decrease3,4. They facilitate the expulsion of chloride ions and organic osmolytes, which is followed by water efflux in response to osmotic swelling. Beyond their role in osmotic stress response, their role in cellular volume regulation has been linked to cell proliferation and migration, apoptosis, spermiogenesis, and epithelial integrity5,6,7. Alteration of the membrane potential upon LRRC8/VRAC activation has been shown to contribute to myotube differentiation8 and insulin secretion by pancreatic &#946;-cells9,10,11. Furthermore, LRRC8 channels conduct a variety of organic osmolytes such as purinergic signaling molecules ATP and cGAMP or the excitatory amino acid glutamate, placing these channels in cell-cell communication in the immune system or glia-neuron interaction12,13,14,15,16,17,18,19,20,21,22. Even xenobiotics, such as the dye fluorescein, the antibiotic blasticidin S or the anticancer drug cisplatin, are conducted by LRRC8 channels23,24,25.
There are numerous reports on the signal transduction leading to LRRC8/VRAC activation26,27,28. However, the mechanism remains unclear, and the literature presents a broad range of potential mechanisms that could depend on the specific physiological process. These include changes in cytosolic ion strength, interaction with the cytoskeleton, membrane composition, G proteins, the redox state, and phosphorylation cascades2,27,29,30,31.
LRRC8/VRAC channels contain LRRC8A as an essential subunit3,4 that must heteromerize with at least one of its paralogues LRRC8B-E to form physiologically functional channels4,14,32. The subunit composition determines biophysical properties of the channel, such as rectification and depolarization-dependent inactivation4,29,32,33,34, substrate specificity15,17,20,21,24,35, and some activation pathways36,37. Cryo-electron microscopy (cryo-EM) structures show that LRRC8A homomers, as well as heteromers, assemble as hexamers38,39,40, while LRRC8A/LRRC8C chimeras that form functional channels are heptamers41. The N-terminal part of all LRRC8 proteins comprises four transmembrane helices, and the C-terminal part contains a domain with leucine-rich repeats (LRRD). The available LRRC8 complex structures provide evidence that the LRRDs, which stretch into the cytosol3,4,23, may undergo conformational rearrangements during channel gating34,42,43. This notion is corroborated by the finding that C-terminal fusion of fluorescent proteins results in basal channel activity14 and that binding of nanobodies to the domains can modulate channel activity44. Moreover, conformational alterations of the C-termini were shown by intra-complex F&#246;rster resonance energy transfer (FRET)45.
The most common method to study ion channel activity is electrophysiological measurements46, which were extensively applied in the investigation of VRACs before their molecular identification47. However, there are various additional ways to monitor VRAC activity indirectly, including the measurement of its conducted substrates -halide ions or organic osmolytes- or its effect on cell volume48. In fact, the identification of LRRC8 proteins as VRAC relied on an assay based on the quenching of a halide-sensitive fluorescent protein49 by iodide entering the cell through activated VRACs3,4. Another method to monitor LRRC8/VRAC channel activity makes use of the movement of the cytosolic domains which can be observed, as in other ion channels50,51,52,53, by changes in FRET45. To this end, fluorescent proteins that serve as FRET pairs, such as cyan-fluorescent protein (CFP)/mCerulean3 as donor and yellow-fluorescent protein (YFP)/mVenus as acceptor, were fused to the C-termini of the LRRC8 proteins (Figure 1). Intra-complex FRET between LRRC8 subunits was shown by acceptor photobleaching experiments45. Avoiding the destructive photobleaching method, FRET changes over time were monitored by sensitized-emission FRET (SE-FRET), where basically the sensitized emission of the acceptor upon excitation of the donor due to the overlap of the emission spectrum of the donor with the excitation spectrum of the acceptor is measured. Application of extracellular hypotonicity, a stimulus for LRRC8/VRAC activation, resulted in a reversible reduction in SE-FRET intensity45. Importantly, simultaneous whole-cell patch-clamp measurements and FRET monitoring during hypotonic treatment showed that this reduction in FRET indeed mirrored LRRC8/VRAC activation45. This method, which avoids disrupting the plasma membrane or altering the intracellular environment by pipette solution, offers an alternative for monitoring LRRC8/VRAC activity. It is particularly useful in physiological settings where maintaining the native cytosol is crucial, subcellular resolution is necessary, or prolonged observation of channel activity is required.
Here, we present a protocol to study LRRC8/VRAC with such a FRET-based read-out. The protocol depicts how to handle and transfect cells, acquire sample and control images, analyze the data, and calculate sensitized emission FRET (SE-FRET) values.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66953/66953fig01v2.jpg" /> 
Figure 1: Schematic of the LRRC8 FRET pair system. mCerulean3 is shown in cyan, and mVenus is shown in yellow. Following VRAC opening, the distance (and/or the spatial orientation) between the fluorophores changes, resulting in a reduced energy transfer between donor (Don) and acceptor (Acc) and, in turn, lowering the observed FRET.&#160;Created with BioRender.com.&#160;<a href="https://www.jove.com/files/ftp_upload/66953/66953fig01largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table><tbody><tr><td>0.05% Trypsin-EDTA</td><td>gibco</td><td>25300-054</td><td></td></tr><tr><td>Camera DFC9000GTC</td><td>Leica</td><td>11547007</td><td></td></tr><tr><td>CFP-18aa-YFP</td><td>N/A</td><td>N/A</td><td>Elder et al. 2009 PMCID: PMC2706461; Gift from C.F. Kaminski (University of Cambridge, UK)</td></tr><tr><td>Cycloheximide (CHX)</td><td>Sigma-Aldrich</td><td>66-81-9</td><td></td></tr><tr><td>D(-)-Mannitol</td><td>Carl Roth</td><td>4175.1</td><td></td></tr><tr><td>D(+)-Glucose</td><td>Carl Roth</td><td>HN06.1</td><td></td></tr><tr><td>DMEM (Dulbeccos Modified Eagle Medium)</td><td>PAN-Biotech</td><td>P04-03590</td><td></td></tr><tr><td>DPBS (Dulbecco&#039;s Phosphate Buffer Saline)</td><td>PAN-Biotech</td><td>P04-36500</td><td></td></tr><tr><td>Emission filter wheel (460/80, 535/70, 590/50, 642/80, 100%)</td><td>Leica</td><td>11525480</td><td></td></tr><tr><td>FBS (Fetal Bovine Serum)</td><td>PAN-Biotech</td><td>P30-3302</td><td></td></tr><tr><td>Filter cube CYR71010</td><td>Leica</td><td>11525416</td><td></td></tr><tr><td>FuGENE</td><td>Promega</td><td>E2691</td><td></td></tr><tr><td>Glas Bottom Culture Dishes 35 mm</td><td>MatTek</td><td>P35G-0-10-C</td><td></td></tr><tr><td>HeLa cells</td><td>Leibniz Forschungsinstitut DSMZ</td><td>ACC 57</td><td>Mammalian cervix carcinoma/ Obtained from Leibniz Forschungsinstitut DSMZ</td></tr><tr><td>HEPES</td><td>Carl Roth</td><td>9105.4</td><td></td></tr><tr><td>ibidi &amp;micro;-Disch 35 mm</td><td>ibidi</td><td>81156</td><td></td></tr><tr><td>KCl (Potassium chloride)</td><td>Carl Roth</td><td>6781.1</td><td></td></tr><tr><td>LAS X FRET Wizard</td><td>Leica</td><td>11640862</td><td></td></tr><tr><td>Light source LED8</td><td>Leica</td><td>11504256</td><td></td></tr><tr><td>LRRC8A-mCerulean3</td><td>N/A</td><td>N/A</td><td>K&amp;ouml;nig et al. 2019</td></tr><tr><td>LRRC8E-mVenus</td><td>N/A</td><td>N/A</td><td>K&amp;ouml;nig et al. 2019</td></tr><tr><td>Luna-II Automated Cell Counter</td><td>logos biosystems</td><td>L40002</td><td></td></tr><tr><td>Luna-II Cell Counter Slides</td><td>logos biosystems</td><td>L12001</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt; (Magnesium chloride)</td><td>Carl Roth</td><td>KK36.1</td><td></td></tr><tr><td>Microscope THUNDER Imager live cell</td><td>Leica</td><td>11525681</td><td></td></tr><tr><td>NaCl (Sodium chloride)</td><td>Carl Roth</td><td>9263</td><td></td></tr><tr><td>Objective HC PL APO 63x/1.40 OIL</td><td>Leica</td><td>11506349</td><td></td></tr><tr><td>Opti-Minimal Essential Medium (MEM)</td><td>gibco</td><td>11058</td><td></td></tr><tr><td>Osmometer OM807</td><td>Vogel</td><td>V04807</td><td></td></tr><tr><td>Penicillin Streptomycin (Pen Step)</td><td>gibco</td><td>15070-063</td><td></td></tr><tr><td>Trypan blue solution (0,4%)</td><td>Sigma</td><td>T8154</td><td></td></tr><tr><td>Tumor necrosis factor (TNF)-a</td><td>Sigma-Aldrich</td><td>94948-59-1</td><td></td></tr><tr><td>Valve Controlled Gravity Perfusion System</td><td>ALA Scientific Instruments</td><td>VC3-4xG</td><td></td></tr></tbody></table>

monitoring leucine-rich repeat containing 8 channel (lrrc8/vrac) activity using sensitized-emission f&#246;rster resonance energy transfer (se-fret)

Members of the LRRC8 protein family form heteromeric ion and osmolyte channels with roles in numerous physiological processes. As volume-regulated anion channels (VRACs)/volume-sensitive outwardly rectifying channels (VSORs), they are activated upon osmotic cell swelling and mediate the extrusion of chloride and organic osmolytes, leading to the efflux of water and hence cell shrinkage. Beyond their role in osmotic volume regulation, VRACs have been implicated in cellular processes such as differentiation, migration, and apoptosis. Through their effect on membrane potential and their transport of various signaling molecules, leucine-rich repeat containing 8 (LRRC8) channels play roles in neuron-glia communication, insulin secretion, and immune response. The activation mechanism has remained elusive. LRRC8 channels, like other ion channels, are typically studied using electrophysiological methods. Here, we describe a method to detect LRRC8 channel activation by measuring intra-complex sensitized-emission F&#246;rster resonance energy transfer (SE-FRET) between fluorescent proteins fused to the C-terminal leucine-rich repeat domains of LRRC8 subunits. This method offers the possibility to study channel activation in situ without exchange of the cytosolic environment and during processes such as cell differentiation and apoptosis.

With this FRET-based method, LRRC8/VRAC activity can be monitored during osmotic stimulation, and the reduction in SE-FRET correlates with the degree of extracellular hypotonicity45. Representative results for hypotonicity-induced channel activation are also shown here (Figure 3 and Figure 4). In addition, LRRC8/VRAC activation by different isosmotic stimuli, such as manipulation of diacylglycerol signaling45 or during myocyte activation56, can be observed.
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/66953/66953fig04.jpg" /> 
Figure 4: SE-FRET traces. (A) Apparent SE-FRET traces from 5 independent experiments. Data represent mean &#177; SD of N = 2 to N = 7 cells per field of view (FOV). (B) Mean &#177; SD of all cells (N = 31 cells). <a href="https://www.jove.com/files/ftp_upload/66953/66953fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
As LRRC8/VRAC channels are also involved in apoptotic volume decrease24,57, observing channel activity upon induction of apoptosis would be another application for the method described here. Accordingly, the protocol to monitor SE-FRET in HeLa cells expressing LRRC8A-mCerulean3 and LRRC8E-mVenus was carried out while applying death receptor-mediated apoptosis-inducing drugs. Tumor necrosis factor (TNF)-&#945; and cycloheximide (CHX) were previously shown to evoke VRAC currents within a few minutes58. After the addition of 2 ng/mL TNF-&#945; and 1 &#181;g/mL CHX in isotonic buffer, there was a robust decrease in SE-FRET (Figure 5). Replacing the buffer by a hypertonic medium, even though containing TNF-&#945; and CHX, SE-FRET values recovered close to baseline (Figure 5A), corresponding to diminishing VRAC currents in hypertonic bath solution during treatment with apoptosis-inducing Fas ligand58. Treatment of the cells with DMSO, the solvent for TNF-&#945; and CHX, did not result in SE-FRET reduction. TNF-&#945; + CHX did not affect SE-FRET of CFP-18aa-YFP, an EYFP and ECFP tandem construct as FRET control59, demonstrating the specificity for LRRC8/VRAC (Figure 5B).
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/66953/66953fig05v2.jpg" /> 
Figure 5: Isosmotic VRAC activation by death receptor-mediated apoptosis. (A) Normalized SE-FRET values from LRRC8A-mCer/LRRC8E-mVen-expressing HeLa cells (n = 8 dishes, 23 cells) over time. After 15 cycles in isotonic imaging buffer (baseline), the bath solution was replaced by isotonic buffer supplemented with 2 ng/mL TNF-&#945; and 1 &#181;g/mL cycloheximide (CHX) for 30 cycles followed by 20 cycles hypertonic imaging buffer with TNF-&#945; and CHX. The raw mean SE-FRET value of each ROI and time point was normalized to the mean of the baseline (isotonic) value for the respective ROI. (B) Quantification of normalized SE-FRET values of LRRC8A/E-expressing HeLa cells as in A with isotonic solution containing DMSO as vehicle control for CHX (n = 5 dishes, 12 cells) or isotonic buffer containing apoptosis inducers as in A (n = 8 dishes, 23 cells), or of HeLa cells expressing CFP-18aa-YFP with apoptosis inducers (n = 3 dishes, 9 cells). Data represent the mean of the last 10 time points in the respective buffer of individual cells (symbols) and mean of all cells &#177; SD; ** p &#60; 0.01 ordinary one-way ANOVA followed by Tukey&#180;s multiple comparison post-hoc test. <a href="https://www.jove.com/files/ftp_upload/66953/66953fig05largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Monitoring Leucine-Rich Repeat Containing 8 Channel (LRRC8/VRAC) Activity Using Sensitized-Emission F&#246;rster Resonance Energy Transfer (SE-FRET)

Electrophysiology is the gold standard for investigating ion channel activity. However, there are plenty of alternative approaches, including optical methods. Here, we describe a method to monitor the activity of the leucine-rich repeat containing 8 channel (LRRC8)-formed anion channels using an inter-subunit F&#246;rster resonance energy transfer (FRET)-based method.

Members of the LRRC8 protein family form heteromeric ion and osmolyte channels with roles in numerous physiological processes. As volume-regulated anion ...

monitoring-leucine-rich-repeat-containing-8-channel-lrrc8vrac

Universidad Científica del Sur

Research

JoVE Journal

Biochemistry

387 Views.  MSH Medical School Hamburg. Electrophysiology is the gold standard for investigating ion channel activity. However, there are plenty of alternative approaches, including optical methods. Here, we describe a method to monitor the activity of the leucine-rich repeat containing 8 channel (LRRC8)-formed anion channels using an inter-subunit Förster resonance energy transfer (FRET)-based method.

Video: Monitoring Leucine-Rich Repeat Containing 8 Channel (LRRC8/VRAC) Activity Using Sensitized-Emission Förster Resonance Energy Transfer (SE-FRET)

Quantitative FRET (F&#246;rster Resonance Energy Transfer) Analysis for SENP1 Protease Kinetics Determination

Reversible posttranslational modifications of proteins with ubiquitin or ubiquitin-like proteins (Ubls) are widely used to dynamically regulate protein activity and have diverse roles in many biological processes. For example, SUMO covalently modifies a large number or proteins with important roles in many cellular processes, including cell-cycle regulation, cell survival and death, DNA damage response, and stress response 1-5. SENP, as SUMO-specific protease, functions as an endopeptidase in the maturation of SUMO precursors or as an isopeptidase to remove SUMO from its target proteins and refresh the SUMOylation cycle 1,3,6,7.
The catalytic efficiency or specificity of an enzyme is best characterized by the ratio of the kinetic constants, kcat/KM. In several studies, the kinetic parameters of SUMO-SENP pairs have been determined by various methods, including polyacrylamide gel-based western-blot, radioactive-labeled substrate, fluorescent compound or protein labeled substrate 8-13. However, the polyacrylamide-gel-based techniques, which used the "native" proteins but are laborious and technically demanding, that do not readily lend themselves to detailed quantitative analysis. The obtained kcat/KM from studies using tetrapeptides or proteins with an ACC (7-amino-4-carbamoylmetylcoumarin) or AMC (7-amino-4-methylcoumarin) fluorophore were either up to two orders of magnitude lower than the natural substrates or cannot clearly differentiate the iso- and endopeptidase activities of SENPs.
Recently, FRET-based protease assays were used to study the deubiquitinating enzymes (DUBs) or SENPs with the FRET pair of cyan fluorescent protein (CFP) and yellow fluorescent protein (YFP) 9,10,14,15. The ratio of acceptor emission to donor emission was used as the quantitative parameter for FRET signal monitor for protease activity determination. However, this method ignored signal cross-contaminations at the acceptor and donor emission wavelengths by acceptor and donor self-fluorescence and thus was not accurate.
We developed a novel highly sensitive and quantitative FRET-based protease assay for determining the kinetic parameters of pre-SUMO1 maturation by SENP1. An engineered FRET pair CyPet and YPet with significantly improved FRET efficiency and fluorescence quantum yield, were used to generate the CyPet-(pre-SUMO1)-YPet substrate 16. We differentiated and quantified absolute fluorescence signals contributed by the donor and acceptor and FRET at the acceptor and emission wavelengths, respectively. The value of kcat/KM was obtained as (3.2 &plusmn; 0.55) x107 M-1s-1 of SENP1 toward pre-SUMO1, which is in agreement with general enzymatic kinetic parameters. Therefore, this methodology is valid and can be used as a general approach to characterize other proteases as well.

Quantitative FRET F&#246;rster Resonance Energy Transfer Analysis for SENP1 Protease Kinetics Determination

A novel method involving quantitative analysis of FRET (F&ouml;rster Resonance Energy Transfer) signals is described for studying enzyme kinetics. KM and kcat were obtained for the hydrolysis of the catalytic domain of SENP1 (SUMO/Sentrin specific protease 1) to pre-SUMO1 (Small Ubiquitin-like MOdifier). The general principles of this quantitative-FRET-based protease kinetic study can be applied to other proteases.

Reversible posttranslational modifications of proteins with ubiquitin or ubiquitin-like proteins (Ubls) are widely used to dynamically regulate protein ...

Bioengineering

Mapping Metabolism: Monitoring Lactate Dehydrogenase Activity Directly in Tissue

Mapping enzymatic activity in space and time is critical for understanding the molecular basis of cell behavior in normal tissue and disease. In situ metabolic activity assays can provide information about the spatial distribution of metabolic activity within a tissue. We provide here a detailed protocol for monitoring the activity of the enzyme lactate dehydrogenase directly in tissue samples. Lactate dehydrogenase is an important determinant of whether consumed glucose will be converted to energy via aerobic or anaerobic glycolysis. A solution containing lactate and NAD is provided to a frozen tissue section. Cells with high lactate dehydrogenase activity will convert the provided lactate to pyruvate, while simultaneously converting provided nicotinamide adenine dinucleotide (NAD) to NADH and a proton, which can be detected based on the reduction of nitrotetrazolium blue to formazan, which is visualized as a blue precipitate. We describe a detailed protocol for monitoring lactate dehydrogenase activity in mouse skin. Applying this protocol, we found that lactate dehydrogenase activity is high in the quiescent hair follicle stem cells within the skin. Applying the protocol to cultured mouse embryonic stem cells revealed higher staining in cultured embryonic stem cells than mouse embryonic fibroblasts. Analysis of freshly isolated mouse aorta revealed staining in smooth muscle cells perpendicular to the aorta. The methodology provided can be used to spatially map the activity of enzymes that generate a proton in frozen or fresh tissue.

We describe a protocol for mapping the spatial distribution of enzymatic activity for enzymes that generate nicotinatmide adenine dinucleotide phosphate (NAD(P)H) + H+ directly in tissue samples.

Mapping enzymatic activity in space and time is critical for understanding the molecular basis of cell behavior in normal tissue and disease. In situ ...

Luminescence Resonance Energy Transfer to Study Conformational Changes in Membrane Proteins Expressed in Mammalian Cells

Luminescence Resonance Energy Transfer, or LRET, is a powerful technique used to measure distances between two sites in proteins within the distance range of 10-100 &#197;. By measuring the distances under various ligated conditions, conformational changes of the protein can be easily assessed. With LRET, a lanthanide, most often chelated terbium, is used as the donor fluorophore, affording advantages such as a longer donor-only emission lifetime, the flexibility to use multiple acceptor fluorophores, and the opportunity to detect sensitized acceptor emission as an easy way to measure energy transfer without the risk of also detecting donor-only signal. Here, we describe a method to use LRET on membrane proteins expressed and assayed on the surface of intact mammalian cells. We introduce a protease cleavage site between the LRET fluorophore pair. After obtaining the original LRET signal, cleavage at that site removes the specific LRET signal from the protein of interest allowing us to quantitatively subtract the background signal that remains after cleavage. This method allows for more physiologically relevant measurements to be made without the need for purification of protein.

We describe here an improved Luminescence Resonance Energy Transfer (LRET) method where we introduce a protease cleavage site between the donor and acceptor fluorophore sites. This modification allows us to obtain specific LRET signals arising from membrane proteins of interest, allowing for the study of membrane proteins without protein purification.

Luminescence Resonance Energy Transfer, or LRET, is a powerful technique used to measure distances between two sites in proteins within the distance range ...

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 ...

Sensitive Detection of Proteopathic Seeding Activity with FRET Flow Cytometry

Increasing evidence supports transcellular propagation of toxic protein aggregates, or proteopathic seeds, as a mechanism for the initiation and progression of pathology in several neurodegenerative diseases, including Alzheimer's disease and the related tauopathies. The potentially critical role of tau seeds in disease progression strongly supports the need for a sensitive assay that readily detects seeding activity in biological samples. 
By combining the specificity of fluorescence resonance energy transfer (FRET), the sensitivity of flow cytometry, and the stability of a monoclonal cell line, an ultra-sensitive seeding assay has been engineered and is compatible with seed detection from recombinant or biological samples, including human and mouse brain homogenates. The assay employs monoclonal HEK 293T cells that stably express the aggregation-prone repeat domain (RD) of tau harboring the disease-associated P301S mutation fused to either CFP or YFP, which produce a FRET signal upon protein aggregation. The uptake of proteopathic tau seeds (but not other proteins) into the biosensor cells stimulates aggregation of RD-CFP and RD-YFP, and flow cytometry sensitively and quantitatively monitors this aggregation-induced FRET. The assay detects femtomolar concentrations (monomer equivalent) of recombinant tau seeds, has a dynamic range spanning three orders of magnitude, and is compatible with brain homogenates from tauopathy transgenic mice and human tauopathy subjects. With slight modifications, the assay can also detect seeding activity of other proteopathic seeds, such as &#945;-synuclein, and is also compatible with primary neuronal cultures. The ease, sensitivity, and broad applicability of FRET flow cytometry makes it useful to study a wide range of protein aggregation disorders.

Cell-to-cell transfer of protein aggregates, or proteopathic seeds, may underlie the progression of pathology in neurodegenerative diseases. Here, a novel FRET flow cytometry assay is described that enables specific and sensitive detection of seeding activity from recombinant or biological samples.

Increasing evidence supports transcellular propagation of toxic protein aggregates, or proteopathic seeds, as a mechanism for the initiation and ...

CRAF-14-3-3 Interaction Analysis in Live Cells Using a BRET-Based Approach

Bioluminescence Resonance Energy Transfer (BRET)-Based Assay for Measuring Interactions of CRAF with 14-3-3 Proteins in Live Cells

CRAF is a primary effector of RAS GTPases and plays a critical role in the tumorigenesis of several KRAS-driven cancers. In addition, CRAF is a hotspot for germline mutations, which are shown to cause the developmental RASopathy, Noonan syndrome. All RAF kinases contain multiple phosphorylation-dependent binding sites for 14-3-3 regulatory proteins. The differential binding of 14-3-3 to these sites plays essential roles in the formation of active RAF dimers at the plasma membrane under signaling conditions and in maintaining RAF autoinhibition under quiescent conditions. Understanding how these interactions are regulated and how they can be modulated is critical for identifying new therapeutic approaches that target RAF function. Here, I describe a bioluminescence resonance energy transfer (BRET)-based assay for measuring the interactions of CRAF with 14-3-3 proteins in live cells. Specifically, this assay measures the interactions of CRAF fused to a Nano luciferase donor and 14-3-3 fused to a Halo tag acceptor, where the interaction of RAF and 14-3-3 results in donor-to-acceptor energy transfer and the generation of the BRET signal. The protocol further shows that this signal can be disrupted by mutations shown to prevent 14-3-3 binding to each of its high-affinity RAF docking sites. This protocol describes the procedures for seeding, transfecting, and replating the cells, along with detailed instructions for reading BRET emissions, performing data analysis, and confirming protein expression levels. In addition, example assay results, along with optimization and troubleshooting steps, are provided.

Author Spotlight: Integrating BRET-Based Assays and Rare Mutation Analysis to Decipher RAF Kinase Regulation in Live Cells

This protocol describes a BRET-based assay for measuring the interactions of the CRAF kinase with 14-3-3 proteins in live cells. The protocol outlines steps for preparing the cells, reading BRET emissions, and data analysis. An example result with identification of appropriate controls and troubleshooting for assay optimization is also presented.

CRAF is a primary effector of RAS GTPases and plays a critical role in the tumorigenesis of several KRAS-driven cancers. In addition, CRAF is a hotspot ...

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 ...

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 ...

Biology

Investigating Interactions Between Histone Modifying Enzymes and Transcription Factors in vivo by Fluorescence Resonance Energy Transfer

Epigenetic regulation of gene expression is commonly affected by histone modifying enzymes (HMEs) that generate heterochromatic or euchromatic histone marks for transcriptional repression or activation, respectively. HMEs are recruited to their target chromatin by transcription factors (TFs). Thus, detecting and characterizing direct interactions between HMEs and TFs are critical for understanding their function and specificity better. These studies would be more biologically relevant if performed in vivo within living tissues. Here, a protocol is described for visualizing interactions in plant leaves between a plant histone deubiquitinase and a plant transcription factor using fluorescence resonance energy transfer (FRET), which allows the detection of complexes between protein molecules that are within &#60;10 nm from each other. Two variations of the FRET technique are presented: SE-FRET (sensitized emission) and AB-FRET (acceptor bleaching), in which the energy is transferred non-radiatively from the donor to the acceptor or emitted radiatively by the donor upon photobleaching of the acceptor. Both SE-FRET and AB-FRET approaches can be adapted easily to discover other interactions between other proteins in planta.

Investigating Interactions Between Histone Modifying Enzymes and Transcription Factors in vivo by Fluorescence Resonance Energy Transfer

Fluorescence resonance energy transfer (FRET) is an imaging technique for detecting protein interactions in living cells. Here, a FRET protocol is presented to study the association of histone-modifying enzymes with transcription factors that recruit them to the target promoters for epigenetic regulation of gene expression in plant tissues.

Epigenetic regulation of gene expression is commonly affected by histone modifying enzymes (HMEs) that generate heterochromatic or euchromatic histone ...

F&#246;rster Resonance Energy Transfer (FRET)

F&#246;rster resonance energy transfer (FRET) is a phenomenon used to investigate close-range biochemical interactions. In FRET, a donor photoluminescent molecule can non-radiatively transfer energy to an acceptor molecule if their respective emission and absorbance spectra overlap. The amount of energy transferred&#8212;and consequently the overall emission of sample&#8212;depends on the proximity of an acceptor-donor pair of photoluminescent molecules. FRET analysis is combined with other biochemistry techniques to obtain detailed information of biomolecular structures and interactions from this &#8220;spectroscopic ruler.&#8221;
This video covers the principles and concepts of FRET analysis. The procedure focuses on preparing samples for FRET and ways to present and interpret data. Finally, the applications include monitoring conformational and cellular processes by labeling parts of a cell or protein, monitoring enzyme reactions that alter protein structures, and using FRET to monitor aggregation of monomers expressed by cells.
<div class="chapter_text" id="script_0">
F&#246;rster Resonance Energy Transfer, or FRET, is a non-radiative transfer of energy between light-emitting molecules, and is often used to investigate close-range biochemical interactions. FRET only occurs when fluorescent molecules are spaced within 10 nm of each other. FRET analysis can be combined with other techniques to obtain detailed structural information. This video will introduce the underlying principles of FRET, summarize a protocol and data presentation, and discuss some biochemical applications.
</div>
<div class="chapter_text" id="script_1">
A photoluminescent molecule such as a fluorophore is excited by absorbing electromagnetic radiation at a wavelength in its absorption spectrum. As it relaxes, it emits light at a wavelength within its emission spectrum. For more information about fluorescence, see JoVE&#39;s video on fluorescence microscopy. Different fluorophores absorb and emit light at different wavelengths, which frequently overlap. If the emission spectrum of a fluorophore overlaps significantly with the absorption spectrum of another fluorophore, the &#8220;donor&#8221; will release a virtual photon, which is absorbed by the &#8220;acceptor&#8221;. When an excited donor is within 10 nm of an acceptor, energy is transferred from donor to acceptor by dipole-dipole interactions. The release of energy by emission of light from the donor correspondingly decreases. Meanwhile, the excited acceptor emits light at its emission wavelength. The FRET response is evaluated in terms of efficiency, or the percentage of energy released from the donor by FRET rather than by fluorescence or other radiative processes. The efficiency depends strongly on the distance between the donor and acceptor, which allows FRET to act as a &#39;molecular&#39; or &#39;spectroscopic&#39; ruler.
In biochemistry, FRET is often used qualitatively to observe conformational changes in molecules by monitoring fluorophores as they move in and out of FRET range of each other. Similarly, cellular functions can be studied with molecules containing a FRET pair. If the labeled molecule is cleaved by enzyme activity, FRET stops and the observed fluorescence wavelength changes.
Now that you understand the principles behind FRET, let&#39;s look at an overview of a protocol and a few ways to present and interpret the data.
</div>
<div class="chapter_text" id="script_2">
Prior to the experiment, the biomolecules of interest, typically DNA or proteins, are engineered with fluorescent tags, using molecular biology techniques.&#160;Common ways to introduce the modified genetic material into the cells include transfection and electroporation.
Then, the cells are prepared for FRET visualization on a fluorescence microscope.&#160;For instance, the molecules may be immobilized on a slide for single-molecule FRET, or samples are loaded into wells for high-throughput screening.
Then, the excitation lasers, microscope, and associated equipment are prepared. (A) FRET experiments often involve powerful lasers; (B) so appropriate PPE and safety procedures should be used.&#160;The sample is then placed in the instrument and illuminated with the excitation laser.
</div>
<div class="chapter_text" id="script_3">
For experiments monitoring cell behavior, color images showing differences or changes in emission intensity are used. Donor and acceptor emission intensities are plotted together to track FRET response over time.
FRET data can also be fitted to various functions for more complex analyses. Depending on the experiment, data may be presented in multiple ways to best represent the results, making FRET a flexible experimental tool.
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Now that you&#39;re familiar with the basics of running and analyzing a FRET experiment, let&#39;s look at some applications of FRET in biochemistry research.
FRET can be used to study conformational changes or cellular processes by labeling parts of the protein or cell predicted to move within 10 nm of each other with a FRET pair. For example, protein sensors are prepared by labeling receptors with a pair of fluorophores. The FRET response is monitored live by confocal microscopy. Variation of emission wavelength and intensity indicate conformational changes.
FRET can also be used by preparing molecules with an active FRET pair and observing changes in the response. When the substrate is cleaved, FRET is disrupted, causing an increase in donor emission and a decrease in acceptor emission. The emissions are analyzed to determine contributions by donor, acceptor, and FRET. Once the direct emission factors are calculated for the cyan and yellow fluorescent proteins, the concentration and kinetic parameters of the substrate can be determined.
Cells designed to express monomers containing either of a FRET pair function as &#39;sensors&#39; for interactions between those monomers. If aggregation of those monomers is induced, a FRET response is observed. This can be used to investigate protein aggregation triggered by &#39;seeding&#39; of misfolded proteins. Here, cells were transduced with aggregates of the protein of interest, incubated, and analyzed with flow cytometry.
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You&#39;ve just watched JoVE&#39;s video on F&#246;rster Resonance Energy Transfer, or FRET. This video contained the underlying principles of FRET, preparation and analysis of a FRET experiment, and a few biochemical applications.
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F&amp;#246;rster Resonance Energy Transfer (FRET)

F&#246;rster resonance energy transfer (FRET) is a phenomenon used to investigate close-range biochemical interactions. In FRET, a donor photoluminescent ...

Monitoring Leucine-Rich Repeat Containing 8 Channel (LRRC8/VRAC) Activity Using Sensitized-Emission Förster Resonance Energy Transfer (SE-FRET)

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