We acknowledge Dr. Jesse Kwiek (The Ohio State University) for kindly allowing us to use his multi-mode detection platform for some preliminary experiments. Research reported in this article was supported by the National Institute of Allergy and Infectious Diseases of the National Institutes of Health under award number RO1AI107250 to Stephanie Seveau. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

1. Preparation
<ol>
	<li>Cell Plating 
	Note: Human cervical epithelial cells, HeLa and HeLa expressing Histone 2B-GFP (H2B-GFP), were used in this protocol, but this assay can be adapted to other mammalian cells19.
	<ol>
		<li>Detach adherent cells from a 75 cm2 cell culture flask by washing the cells with 2 mL of Trypsin-EDTA 0.25%. Replace the used trypsin with 2 mL of fresh trypsin-EDTA 0.25%.</li>
		<li>Incubate the cells at 37 &#730;C for 5 min until the cells have rounded and detached from the flask.</li>
		<li>Resuspend the cells in 8 mL of growth medium (DMEM containing 10% heat-inactivated fetal bovine serum, 100 U/mL penicillin, and 100 &#181;g/mL streptomycin).</li>
		<li>Determine the cell concentration using a hemocytometer and 10 &#181;L of cell suspension.</li>
		<li>Dilute the cells in growth medium to a concentration of 2.5 x 105 cells/mL.</li>
		<li>Pour the cell suspension into a sterile pipette basin and thoroughly mix the suspension using a 10 mL serological pipette.</li>
		<li>Using a 12-multichannel micropipette and 200 &#181;L tips, distribute HeLa cells (2.5 x 104 cells/100 &#181;L/well) in triplicate (or quadruplicate) in a 96-well flat, clear bottom, black polystyrene tissue culture-treated plate. 
		Note: A plating arrangement is presented as an example in Figure 1.</li>
		<li>Culture the cells for 24 h in a humidified cell culture incubator at 37 &#730;C and 5% CO2.</li>
	</ol>
	</li>
	<li>Stock Solution Preparation
	<ol>
		<li>Prepare 1 L of a 10x stock of buffer M (used to prepare M1 and M2) by adding&#160;95 g of Hanks Balanced Salt Solution,&#160;0.476 g of MgCl2 (5 mM), and&#160;23.83 g of HEPES (100 mM) to 900 mL of water. Adjust the pH to 7.4 and raise the volume to 1 L. Filter sterilize.</li>
		<li>Prepare 50 mL of a 50x (1.25 M) stock of glucose by adding 11.26 g of D-(+)-Glucose to a total of 50 mL of water. Filter sterilize the solution.</li>
		<li>Prepare 50 mL of a 100x (120 mM) stock of calcium by adding 0.666 g of CaCl2 to a total of 50 mL of water. Filter sterilize the solution.</li>
		<li>Prepare 50 mL of a 10x (50 mM) stock of ethylene glycol-bis(2-aminoethylether)-N,N,N&#8217;,N&#8217;,tetraacetic acid (EGTA) by adding 0.951 g of EGTA to 40 mL of water. Increase the pH to 8 using NaOH to dissolve the EGTA, then raise the volume to 50 mL. Filter sterilize the solution.</li>
		<li>For a single 96-well plate, prepare 50 mL of Medium 1 (M1, contains Ca2+), 50 mL of Medium 2 (M2, Ca2+-free), and 15 mL of Medium 2 supplemented with EGTA, accordingly:
		<ol>
			<li>For M1, add 5 mL of 10x Buffer M, 0.5 mL of 100x CaCl2, and 1 mL of 50x glucose to 43.5 mL of water.</li>
			<li>For M2, add 5 mL of 10x Buffer M and 1 mL of 50x glucose to 44 mL of water.</li>
			<li>For M2/EGTA, add 1.5 mL of 10x Buffer M and 1.5 mL of 10x EGTA to 12 mL of water. 
			Note: All solutions containing propidium iodide (PI) should be prepared directly prior to adding to the cells.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Plate Reader/Imaging Cytometer Settings 
	Note: Use a multi-mode plate reader equipped with two detection units: a spectrofluorometer and an imaging cytometer. Limit the fluorescence exposure to avoid photobleaching the fluorophores.
	<ol>
		<li>Pre-warm the plate reader to 37 &#176;C before performing the assay.</li>
		<li>Set up the parameters for the kinetic assay accordingly within the Settings mode:
		<ol>
			<li>Choose Monochromator, FL (fluorescence), and Kinetic for the optical configuration, read modes, and read type, respectively.</li>
			<li>Under Wavelength Settings, select a 9 and 15 nm excitation and emission bandpass, respectively. For assays using propidium iodide (PI), set the excitation and emission wavelengths to 535 and 617 nm, respectively.</li>
			<li>Under Plate Type, select 96 Wells for the plate format and a pre-set plate configuration corresponding to a black-wall clear bottom plate.</li>
			<li>Under Read Area, highlight the wells that will be analyzed throughout the kinetic.</li>
			<li>Under PMT and Optics, preset the flashes per read to 6 and check the box for Read from Bottom.</li>
			<li>Under Timing, insert 00:30:00 in the Total Run Time box for a 30 min kinetic assay, and insert 00:05:00 for the Interval. 
			Note: For each time point and one wavelength, the reading time of a full 96-well plate is 30 s.</li>
			<li>Confirm the specified settings in the Settings Information to the right and select OK. Press Read to initiate the kinetic run.</li>
		</ol>
		</li>
		<li>Set up the imaging parameters accordingly within the Settings mode:
		<ol>
			<li>Choose Minimax, Imaging, and Endpoint for the optical configuration, read modes, and read type, respectively.</li>
			<li>Under Wavelengths, select transmitted light, and either or both the fluorescence boxes corresponding to excitation and emission wavelengths of 456/541 nm (GFP) and 625/713 nm (PI).</li>
			<li>Use the same options for the Plate Type and Read Area as defined in steps 1.3.2.3 and 1.3.2.4.</li>
			<li>Under Well Area Setting, select the number of sites within a well to be imaged. 
			Note: 12 sites correspond to a full-well image.</li>
			<li>Under the Image Acquisition Settings, select the exposure times for transmitted light,&#160;541 (GFP), and 713 (PI). For GFP, image the entire well with an exposure time of 20 ms/image. For transmitted light (TL) and PI fluorescence, acquire a single image of the center of each well with exposure times of 8 and 20 ms, respectively.</li>
			<li>Confirm the specified settings in the Settings Information to the right and select OK. The acquisition time for imaging the entire surface of each well (12 images/well) of a 96-well plate and for one wavelength is ~15 min. Press Read to initiate imaging. 
			Note: The acquisition time of a single image/well of a 96-well plate requires ~2.5 min/plate for one wavelength. The parameters described above correspond to the specific equipment in our laboratory. Spectrofluorometric measurements: A xenon flash lamp displaying 1.0 nm increment excitation wavelengths (250-850 nm) with an adjustable 9 or 15 nm bandpass, a photomultiplier tube detector with a &#62; 6 log dynamic range and an adjustable 15 or 25 nm emission bandpass. Imaging cytometer: An illumination light source capable of white light, 460 nm and 625 nm excitation wavelengths with a 20 nm bandpass, emission filters centered at 541 nm (108 nm bandpass) and 713 nm (123 nm bandpass), respectively, and a 4X objective coupled to a 1.25 megapixel 12-bit charge-coupled device camera.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Assay
Note: At the time of the assay, cells must be 70-90% confluent. During the wash steps, the medium should be removed from and applied to the side-wall of the well (not directly above the cells). Maintain the temperature of LLO at &#60; 4 &#730;C to prevent its aggregation until step 3.1.5.
<ol>
	<li>Prepare a stock of 30 &#181;M PI in M1 and a stock of 30 &#181;M PI in M2 pre-warmed at 37 &#730;C.</li>
	<li>Gently wash the cells in plate 1 using a 12-multichannel micropipette and 200 &#181;L tips, as follows:
	<ol>
		<li>For repair-permissive conditions, remove the growth medium and wash the cells twice with 200 &#181;L/well M1 pre-warmed at 37 &#730;C. Replace the medium with 100 &#181;L/well of warm M1 containing 30 &#181;M PI.</li>
		<li>For repair restrictive-conditions, remove the growth medium and wash the cells once with 200 &#181;L/well warm M2 containing 5 mM EGTA to chelate Ca2+, followed by one wash with 200 &#181;L/well M2. Replace the medium with 100 &#181;L/well warm M2 containing 30 &#181;M PI.</li>
		<li>After the growth medium has been washed and replaced with medium containing propidium iodide, directly move to step 2.1.3.</li>
	</ol>
	</li>
	<li>Image plate 1 under transmitted light, GFP, and PI as detailed under 1.3.3 (pre-kinetic). This step takes 15-20 min.</li>
	<li>During the 15 min period in step 2.1.3, prepare plate 2 using a 12-multichannel micropipette and 200 &#181;L tips as follows:
	<ol>
		<li>Place a 96-well round bottom polypropylene microplate on ice. Configure the plate using an experimental design corresponding to plate 1 (Figure 1).</li>
		<li>For repair-permissive conditions, add 100 &#181;L/well of ice-cold M1 containing 60 &#181;M PI, followed by the addition of 100 &#181;L/well of ice-cold M1 containing 4x LLO or not for the control.</li>
		<li>For repair-restrictive conditions, add 100 &#181;L/well of ice-cold M2 containing 60 &#181;M PI, followed by the addition of 100 &#181;L/well of ice-cold M2 containing 4x LLO or not for the control.</li>
	</ol>
	</li>
	<li>After imaging plate 1 (step 2.1.3), immediately place it on ice, using aluminum foil to separate the plate from direct contact with ice. Allow plate 1 to cool down for 5 min.</li>
	<li>Using a 12-multichannel micropipette and 200 &#181;L tips, transfer 100 &#181;L from each well into plate 2 (step 2.1.4) to the corresponding wells in the plate 1. To properly distribute the toxin in the media of plate 1, insert the tips below the&#160;meniscus and gently eject the volume without introducing bubbles. 
	Note: Do not pipette up and down, as this may inadvertently detach the cells.</li>
	<li>Leave the plate for an additional 1 min to allow the toxin to bind to the&#160;cells and immediately transfer plate 1 to the plate reader for the kinetic assay using the spectrofluorometer mode (step 1.3.2).</li>
	<li>At the end of the kinetic assay, immediately image plate 1 (post-kinetic) using step 1.3.3.</li>
</ol>
3. Analysis: Cell Enumeration
<ol>
	<li>Determine the cell count based on the nuclear fluorescence using the microplate cell enumeration software.
	<ol>
		<li>Within Settings, select Re-analysis, and under the category section within the Image Analysis Settings select Discreet Object Analysis using 541 as the wavelength for finding objects.</li>
		<li>Within the Find Objects option, using the Draw on Images finding method, select Nuclei under the settings tab, and press Apply.</li>
		<li>Press OK and Read to initiate the cell counting algorithm.</li>
	</ol>
	</li>
	<li>Alternatively, if no such tool is available, use an image analysis software such as ImageJ to enumerate cells.
	<ol>
		<li>In ImageJ, open the image file as a stack.</li>
		<li>Convert the stack to 8-bit greyscale images by clicking Image in the menu bar, hover over Type, and select 8-bit.</li>
		<li>Subtract the background: Click Image in the menu bar, hover over Adjust, and select Brightness/Contrast. Adjust the minimum value to remove the background noise and select Apply.</li>
		<li>Threshold to create binary images: Click Image in the menu bar, hover over Adjust, and select Threshold. Select Dark background, adjust the minimum and maximum threshold values, and click Apply.</li>
		<li>In the case of overlapping nuclei, a Watershed tool can be used to segment nuclei. Click Process in the menu, hover over Binary and select Watershed. 
		Note: This will automatically separate connected nuclei.</li>
		<li>Analyze the masked images by applying user-specified criteria (size and circularity) to refine the identification of nuclei and exclude cell debris.
		<ol>
			<li>Click on Analyze in the menu and then Analyze particles. Set the desired size (pixel^2) and circularity (a value of 1 is a perfect circle) ranges that are sufficient to include individual cells/nuclei.</li>
			<li>In the Show dropdown box, select the option(s) desired, check Summarize, and click OK to obtain cell counts.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
4. Analysis: Kinetic Curves
<ol>
	<li>Transfer the kinetic data from the plate reader software to an analytical data software.</li>
	<li>For each experimental condition, average the fluorescence intensities of the replicates at each timepoint, along with the corresponding standard deviation and standard error of the mean for each experimental condition.</li>
	<li>For each experimental condition, trace the corresponding kinetic curve: PI intensity (y-axis) versus time (x-axis).</li>
	<li>To calculate the resealing efficiency of a given treatment condition, calculate the area under the curve (AUC) of the +LLO in M1 (AUC(M1)) and +LLO in M2 (AUC(M2)). Use the approach suggested below to assess the efficiency (E) of resealing: 
	<img alt="Equation" src="/files/ftp_upload/58351/58351eq1v2.jpg" /></li>
	<li>Perform a comparison between control and test treatment by determining the efficiency ratio (REff) indicated below: 
	<img alt="Equation 2" src="/files/ftp_upload/58351/58351eq2.jpg" /> 
	REFF = 1, test treatment has no effect on repair 
	REFF &#60; 1, test treatment inhibits repair 
	REFF &#62; 1, test treatment improves repair</li>
	<li>Calculate the area under the curve using the following equation: 
	<img alt="Equation 3" src="/files/ftp_upload/58351/58351eq3.jpg" />, where k is the total number of follow-ups.</li>
</ol>

The authors have nothing to disclose.

This assay measures the efficiency of membrane resealing at the cell population level with high-throughput capacity. It can be used to screen for cellular components or drug libraries that could affect membrane repair. The described assay used a 96-well plate format, but it can be adapted to 384-well plates for higher throughput. An advantage of this assay is its ability to obtain fluorescence measurements of adherent living cells in real time without the need for excessive cell processing such as cell detachment, fixation, or fluorescence labeling post-fixation. Multimode plate readers, such as the one used in this protocol, have sufficient sensitivity for rapid spectrofluorometric measurements at time intervals as low as 30 s for a 96-well plate. The acquisition of fluorescence images provides additional information including cell enumeration, eventual changes in cell morphology, and potential identification of distinct cell populations. The present assay does not establish the kinetics of plasma membrane resealing at the single cell level, but identifies experimental conditions (pharmacological compounds or cellular components) that can affect, positively or negatively, the process of membrane resealing at the cell population level.
Several other experimental approaches have been developed to assess the membrane resealing mechanisms. For example, mechanical disruption by microneedle puncture, bead abrasion, and laser ablation have been used to model mechanical membrane damage. The measurement of wounding/repair has involved fluorescence microscopy or flow cytometry by quantifying the entry of fluorescent probes (FM 1-43, propidium iodide, fluorescein-conjugated dextrans) or tracking fluorescent protein chimeras47,48,49,50. Each of these approaches has its own advantages; however, they are not amenable to high-throughput screening in living cells as presented in this assay.
The present assay was optimized to analyze the resealing efficiency of cells wounded by the pore-forming protein LLO, which forms large pores that permit the massive Ca2+ influx as provoked by mechanical ruptures of the plasma membrane. Although pore-forming toxins represent one form of plasma membrane damage, repair of large toxin pores and mechanical wounds were proposed to share common Ca2+-dependent pathways17,51. It is important to note that LLO interaction with cell membrane components such as cholesterol may affect cell signaling and thereby may influence membrane resealing mechanisms when compared to mechanical wounds. Our knowledge about membrane repair is still limited and further studies are required to establish if resealing of mechanical wounds differ from resealing following the formation of toxin pores. There are several advantages of using LLO. First, the initiation of membrane damage can be synchronized by raising the temperature from 4 to 37 &#176;C. Second, the soluble form of LLO (not bound to the cell membrane) irreversibly aggregates at neutral pH and 37 &#176;C, thus limiting cytotoxic effects and abrogating the need to wash the cells. Finally, the degree of damage can be adjusted by varying the concentration of the toxin. However, a limitation of this assay is the temperature switch between 37 and 4 &#176;C, which may affect the repair mechanism such as vesicular transport, endocytosis, and membrane fluidity, among other processes, which are influenced by temperature52,53,54. It is important to verify that any pharmacological inhibitor included in the assay does not interfere with the formation of LLO pores by performing a hemolysis assay in the presence (versus absence) of the drug55. Due to potential batch differences in LLO activity, it is important to prepare a stock of LLO that is large enough for an&#160;entire high-throughput screen55.
We included the use of cells expressing the nuclear-localized Histone-2B-GFP chimera as a means to enumerate cells before and after the membrane repair assay via microscopic imaging followed by automated image analysis. Importantly, equivalent cell counts across conditions and unchanging cell counts before and after the kinetic assay are crucial as PI or CNABD fluorescence intensities cannot easily&#160;be normalized. Indeed, a difference in cell count will result in differences in the degree of damage by a given concentration of LLO, which cannot be corrected for via fluorescence normalization due to variations in resealing efficiency (Figure 7). We showed that Histone-2B-GFP expression does not interfere with PI or CNABD incorporation or fluorescence emission. Conversely, PI or CNABD incorporation does not affect GFP-based cell enumeration. If other combinations of fluorophores are to be used, it would be necessary to assess potential spectral overlap between fluorochromes, as was performed in this work. Although PI has been extensively used to measure membrane damage, we show that CNABD is an excellent substitute that exhibits higher fluorescence quantum yield, resulting in a larger dynamic range suitable for characterizing resealing efficiency.
Z-factor and SSMD confirmed that this assay has the robustness necessary to perform high-throughput analyses. The calculation of the resealing efficiency is a critical and reliable tool to identify potential hits. In addition, the mixed effects model can be used as a statistical tool to evaluate hits within a single assay. The experimental plan must include a minimum of three technical replicates if the screen can be repeated several times. Quadruplicates should be used if statistical tools, such as the mixed effects model, are to be included in a single experiment, whether or not it will be repeated. It is advised however to perform the screen several times&#160;and&#160;to validate the results by performing complementary experiments.

Mammalian cells are subject to mechanical, osmotic, and biochemical stress, resulting in the loss of plasma membrane integrity. Without rapid and efficient resealing, damaged cells would quickly succumb to programmed or necrotic death. Since the 1960s, efforts to understand the plasma membrane resealing process have been motivated by the devastating consequences associated with its dysfunctions. Indeed, diseases such as Limb-Girdle Muscular Dystrophy, diabetes, and Chediak-Higashi Syndrome have been linked to deficient plasma membrane repair due to mutations in the gene encoding dysferlin, production of advanced glycation end products, and defects in the lysosomal trafficking regulator CHS1, respectively1,2,3,4,5,6. However, to date, our understanding of membrane resealing is still limited7. Initial studies have demonstrated that membrane resealing is initiated by the influx of extracellular Ca2+ through the damaged plasma membrane8,9,10. Since then, several non-mutually exclusive Ca2+-dependent mechanisms have been proposed to reseal cells. The patch hypothesis proposes that in proximity to the wound, intracellular vesicles fuse with each other and the damaged plasma membrane to act as a patch11,12,13,14. A second model proposes that calcium-dependent exocytosis of lysosomes at the wound site releases the lysosomal enzyme acid sphingomyelinase, which converts sphingomyelin to ceramide in the outer leaflet of the plasma membrane. This sudden change in lipid composition results in ceramide-driven endocytosis of the damaged region15,16,17. Lastly, the third proposed mechanism involves a role for the endosomal sorting complex required for transport (ESCRT) to promote the formation of outward-facing vesicles that bud off from the plasma membrane18. Only a limited set of proteins was identified in these models, and their machinery must be further elucidated.&#160;
Here we describe a high-throughput assay that measures the plasma membrane resealing efficiency in adherent mammalian cells subjected to damage mediated by recombinant listeriolysin O (LLO)19. LLO is a pore-forming toxin (PFT) secreted by the facultative intracellular pathogen Listeria monocytogenes20,21,22 and belongs to the MACPF/CDC (membrane attack complex, perforin, and cholesterol-dependent cytolysin) superfamily. MACPF are mammalian pore-forming proteins involved in immune defenses, whereas CDCs are bacterial toxins mainly produced by Gram-positive pathogens that damage host cells to promote their pathogenic lifestyles23. CDCs are synthesized as water-soluble monomers or dimers that bind to cholesterol present in the plasma membrane and oligomerize into a prepore complex of up to 50 subunits. The prepore complex then rearranges to insert &#946;-strands across the lipid bilayer, forming a &#946;-barrel pore that spans 30-50 nm in diameter24,25,26,27.&#160; These large pores permit fluxes of ions and small cellular components in and out of the cell; though, some studies have proposed that pores of smaller sizes are also formed28,29,30. Among the CDCs, LLO displays unique properties including irreversible pH- and temperature-dependent aggregation, which is conducive to high-throughput analyses31,32. LLO can be added to the cell culture medium at 4 &#730;C, a temperature permissive to its binding to cells, but not to the formation of the pore complex. Initiation of pore formation can then be synchronized by raising the temperature to 37 &#730;C, allowing for the efficient diffusion of toxin molecules in the plane of the membrane to form oligomers and for the conformational remodeling involved in pore generation. Therefore, following the switch in temperature, the kinetic of cell damage will depend on the amount of toxin bound to the plasma membrane. Importantly, soluble LLO (not bound to the plasma membrane) rapidly and irreversibly aggregates when the temperature reaches 37 &#730;C, which alleviates the need to wash away unbound toxin molecules and limits the extent of membrane damage over time. Lastly, because LLO binds to cholesterol and forms pores in cholesterol-rich membranes, this assay is amenable to a wide range of mammalian cells. It is important to keep in mind that LLO affects host cell signaling mainly via pore formation, with a few exceptions in which pore-independent cell signaling may occur33,34,35,36,37,38,39. Therefore, it cannot be excluded that LLO signaling activities may influence the process of membrane repair.&#160;
This assay directly assesses the extent of cell wounding by measuring the incorporation of a cell impermeant fluorochrome (e.g., propidium iodide) that passively enters wounded cells and becomes highly fluorescent once it associates with nucleic acids. Hence, the fluorochrome can be maintained in the cell culture medium throughout the experiment, allowing real-time analyses of cell wounding. The fluorescence intensity of the nucleic acid-binding dye will increase with the concentration of toxin and, for a given concentration of toxin, will increase over time until all pores are formed, and cells are fully repaired or until saturation is reached. The influx of extracellular Ca2+ through membrane pores is a sine qua non event for resealing. Therefore, the resealing efficiency can be indirectly evidenced by comparing cell wounding in culture medium containing Ca2+ (repair permissive condition) to wounding in a Ca2+-free medium (repair restrictive condition). Because the fluorescence intensity of the nucleic acid-binding dye is directly proportional to the cell concentration in each well, it is important to seed cells at the same concentration in all wells. It is also important to enumerate cells in each well before and after the assay to ensure that cell detachment does not occur, as floating, aggregated cells can obscure fluorescence readings which may complicate data interpretation. To enumerate cells, cells expressing nuclear-localized histone 2B-GFP (H2B-GFP) were used in this assay. Temperature-controlled, multi-mode, microplate readers combine rapid, high-throughput measurements (using a 96 or 384-well plate format) of fluorescence intensities with microscopy imaging of living cells at 37 &#176;C. The latter can be used to enumerate cell number and observe the eventual formation of distinct cell populations.
Ultimately, this assay provides users the ability to expand their knowledge of the complexity of membrane repair mechanisms by screening for host molecules or exogenously added compounds that may control membrane repair. The following protocol describes the experimental steps to measure the resealing efficiency of cells exposed to LLO and evaluate the effects of a given drug or cellular treatment on resealing efficiency.&#160;

<table><tbody><tr><td>SpectraMax i3x Multi-Mode Microplate Reader</td><td>Molecular Devices</td><td>i3x</td><td></td></tr><tr><td>MiniMax 300 Imaging cytometer</td><td>Molecular Devices</td><td>5024062</td><td></td></tr><tr><td>TO-PRO-3</td><td>ThermoFisher Scientific</td><td>T3605</td><td></td></tr><tr><td>Propidium Iodide</td><td>ThermoFisher Scientific</td><td>P3566</td><td></td></tr><tr><td>HeLa</td><td>ATCC</td><td>CCL2</td><td></td></tr><tr><td>HeLa H2B-GFP</td><td>Millipore</td><td>SCC117</td><td></td></tr><tr><td>Trypsin-EDTA 0.25%</td><td>ThermoFisher Scientific</td><td>25200056</td><td></td></tr><tr><td>96-well Corning flat bottom black polystyrene tissue culture treated plate</td><td>Corning</td><td>3603</td><td></td></tr><tr><td>Hanks&#039; balanced Salts</td><td>Sigma-Aldrich</td><td>H4891</td><td></td></tr><tr><td>EGTA</td><td>ISC BioExpress</td><td>0732-100G</td><td></td></tr><tr><td>HEPES</td><td>Fisher Scientific</td><td>BP310-500</td><td></td></tr><tr><td>D-(+)-Glucose, HybriMax</td><td>Sigma-Aldrich</td><td>G5146-1KG</td><td></td></tr></tbody></table>

high-throughput measurement of plasma membrane resealing efficiency in mammalian cells

In their physiological environment, mammalian cells are often subjected to mechanical and biochemical stresses that result in plasma membrane damage. In response to these damages, complex molecular machineries rapidly reseal the plasma membrane to restore its barrier function and maintain cell survival. Despite 60 years of research in this field, we still lack a thorough understanding of the cell resealing machinery. With the goal of identifying cellular components that control plasma membrane resealing or drugs that can improve resealing, we have developed a fluorescence-based high-throughput assay that measures the plasma membrane resealing efficiency in mammalian cells cultured in microplates. As a model system for plasma membrane damage, cells are exposed to the bacterial pore-forming toxin listeriolysin O (LLO), which forms large 30-50 nm diameter proteinaceous pores in cholesterol-containing membranes. The use of a temperature-controlled multi-mode microplate reader allows for rapid and sensitive spectrofluorometric measurements in combination with brightfield and fluorescence microscopy imaging of living cells. Kinetic analysis of the fluorescence intensity emitted by a membrane impermeant nucleic acid-binding fluorochrome reflects the extent of membrane wounding and resealing at the cell population level, allowing for the calculation of the cell resealing efficiency. Fluorescence microscopy imaging allows for the enumeration of cells, which constitutively express a fluorescent chimera of the nuclear protein histone 2B, in each well of the microplate to account for potential variations in their number and allows for eventual identification of distinct cell populations. This high-throughput assay is a powerful tool expected to expand our understanding of membrane repair mechanisms via screening for host genes or exogenously added compounds that control plasma membrane resealing.

Watch this Scientific Journal Video about High-throughput Measurement of Plasma Membrane Resealing Efficiency in Mammalian Cells at JoVE.com

High-throughput Measurement of Plasma Membrane Resealing Efficiency in Mammalian Cells

Here we describe a high-throughput fluorescence-based assay that measures the plasma membrane resealing efficiency through fluorometric and imaging analyses in living cells. This assay can be used for screening drugs or target genes that regulate plasma membrane resealing in mammalian cells.

In their physiological environment, mammalian cells are often subjected to mechanical and biochemical stresses that result in plasma membrane damage. In ...

high-throughput-measurement-plasma-membrane-resealing-efficiency

Nanyang Technological University

Research

JoVE Journal

Immunology and Infection

7.7K Views.  The Ohio State University. Deze test is ontwikkeld om belangrijke vragen op het gebied van plasmamembraanbiologie aan te pakken en vast te stellen hoe efficiënt beschadigde cellen hun plasmamembraan kunnen herssepen. Deze techniek kan worden gebruikt om de efficiëntie van plasmamembraanherseten in een hoge doorvoercapaciteit te meten en om specifieke eiwitten en overeenkomstige trajecten te identificeren die plasmamembraan hersluiten. Begin met het toevoegen van 20 milliliter van een 2,5 keer 10 aan de vijfde HeLa ce...

Video: High-throughput Measurement of Plasma Membrane Resealing Efficiency in Mammalian Cells

A Mimic of the Tumor Microenvironment: A Simple Method for Generating Enriched Cell Populations and Investigating Intercellular Communication

Understanding the early heterotypic interactions between cancer cells and the surrounding non-cancerous stroma is important in elucidating the events leading to stromal activation and establishment of the tumor microenvironment (TME). Several in vitro and in vivo models of the TME have been developed; however, in general these models do not readily permit isolation of individual cell populations, under non-perturbing conditions, for further study. To circumvent this difficulty, we have employed an in vitro TME model using a cell growth substrate consisting of a permeable microporous membrane insert that permits simple generation of highly enriched cell populations grown intimately, yet separately, on either side of the insert&#39;s membrane for extended co-culture times. Through use of this model, we are capable of generating greatly enriched cancer-associated fibroblast (CAF) populations from normal diploid human fibroblasts following co-culture (120 hr) with highly metastatic human breast carcinoma cells, without the use of fluorescent tagging and/or cell sorting. Additionally, by modulating the pore-size of the insert, we can control for the mode of intercellular communication (e.g., gap-junction communication, secreted factors) between the two heterotypic cell populations, which permits investigation of the mechanisms underlying the development of the TME, including the role of gap-junction permeability. This model serves as a valuable tool in enhancing our understanding of the initial events leading to cancer-stroma initiation, the early evolution of the TME, and the modulating effect of the stroma on the responses of cancer cells to therapeutic agents.

We adapted a permeable microporous membrane insert to mimic the tumor microenvironment (TME). The model consists of a mixed cell culture, allows simplified generation of highly enriched individual cell populations without using fluorescent tagging or cell sorting, and permits studying intercellular communication within the TME under normal or stress conditions.

Understanding the early heterotypic interactions between cancer cells and the surrounding non-cancerous stroma is important in elucidating the events ...

Cancer Research

Measuring Nucleotide Binding to Intact, Functional Membrane Proteins in Real Time

We have developed a method to measure binding of adenine nucleotides to intact, functional transmembrane receptors in a cellular or membrane environment. This method combines expression of proteins tagged with the fluorescent non-canonical amino acid ANAP, and FRET between ANAP and fluorescent (trinitrophenyl) nucleotide derivatives. We present examples of nucleotide binding to ANAP-tagged KATP ion channels measured in unroofed plasma membranes and excised, inside-out membrane patches under voltage clamp. The latter allows for simultaneous measurements of ligand binding and channel current, a direct readout of protein function. Data treatment and analysis are discussed extensively, along with potential pitfalls and artefacts. This method provides rich mechanistic insights into the ligand-dependent gating of KATP channels and can readily be adapted to the study of other nucleotide-regulated proteins or any receptor for which a suitable fluorescent ligand can be identified.

This protocol presents a method for measuring adenine nucleotide binding to receptors in real time in a cellular environment. Binding is measured as F&#246;rster resonance energy transfer (FRET) between trinitrophenyl nucleotide derivatives and protein labeled with a non-canonical, fluorescent amino acid.

We have developed a method to measure binding of adenine nucleotides to intact, functional transmembrane receptors in a cellular or membrane environment. ...

Biochemistry

Fluorescent Leakage Assay to Investigate Membrane Destabilization by Cell-Penetrating Peptide

Cell-penetrating peptides (CPPs) are defined as carriers that are able to cross the plasma membrane and to transfer a cargo into cells. One of the main common features required for this activity resulted from the interactions of CPPs with the plasma membrane (lipids) and more particularly with components of the extracellular matrix of the membrane itself (heparan sulphate). Indeed, independent of the direct translocation or the endocytosis-dependent internalization, lipid bilayers are involved in the internalization process both at the level of the plasma membrane and at the level of intracellular traffic (endosomal vesicles). In this article, we present a detailed protocol describing the different steps of a large unilamellar vesicles (LUVs) formulation, purification, characterization, and application in fluorescence leakage assay in order to detect possible CPP-membrane destabilization/interaction and to address their role in the internalization mechanism. LUVs with a lipid composition reflecting the plasma membrane content are generated in order to encapsulate both a fluorescent dye and a quencher. The addition of peptides in the extravesicular medium and the induction of peptide-membrane interactions on the LUVs might thus induce in a dose-dependent manner a significant increase in fluorescence revealing a leakage. Examples are provided here with the recently developed tryptophan (W)- and arginine (R)-rich Amphipathic Peptides (WRAPs), which showed a rapid and efficient siRNA delivery in various cell lines. Finally, the nature of these interactions and the affinity for lipids are discussed to understand and to improve the membrane translocation and/or the endosomal escape.

The fluorescence leakage assay is a simple method that enables the investigation of peptide/membrane interactions in order to understand their involvement in several biological processes and especially the ability of cell-penetrating peptides to disturb phospholipids bilayers during a direct cellular translocation process.

Cell-penetrating peptides (CPPs) are defined as carriers that are able to cross the plasma membrane and to transfer a cargo into cells. One of the main ...

Measuring Plasma Membrane Protein Endocytic Rates by Reversible Biotinylation

Plasma membrane proteins are a large, diverse group of proteins comprised of receptors, ion channels, transporters and pumps.  Activity of these proteins is responsible for a variety of key cellular events, including nutrient delivery, cellular excitability, and chemical signaling.  Many plasma membrane proteins are dynamically regulated by endocytic trafficking, which modulates protein function by altering protein surface expression.  The mechanisms that facilitate protein endocytosis are complex and are not fully understood for many membrane proteins.  In order to fully understand the mechanisms that control the endocytic trafficking of a given protein, it is critical that the protein s endocytic rate be precisely measured.  For many receptors, direct endocytic rate measurements are frequently achieved utilizing labeled receptor ligands.  However, for many classes of membrane proteins, such as transporters, pumps and ion channels, there is no convenient ligand that can be used to measure the endocytic rate.  In the present report, we describe a reversible biotinylation method that we employ to measure the dopamine transporter (DAT) endocytic rate.  This method provides a straightforward approach to measuring internalization rates, and can be easily employed for trafficking studies of most membrane proteins.

Regulated endocytosis governs the cell surface expression levels of the majority of membrane proteins. Here we utilize reducible, membrane impermeant biotinylation reagents to measure the endocytic rate of the dopamine transporter (DAT), a polytopic membrane protein. The method facilitates a straightforward approach to measuring the endocytic rate of most plasma membrane proteins.

Plasma membrane proteins are a large, diverse group of proteins comprised of receptors, ion channels, transporters and pumps.  Activity of these proteins ...

Biology

Cell Membrane Repair Assay Using a Two-photon Laser Microscope

Numerous pathophysiological insults can cause damage to cell membranes and, when coupled with innate defects in cell membrane repair or integrity, can result in disease. Understanding the underlying molecular mechanisms surrounding cell membrane repair is, therefore, an important objective to the development of novel therapeutic strategies for diseases associated with dysfunctional cell membrane dynamics. Many in vitro and in vivo studies aimed at understanding cell membrane resealing in various disease contexts utilize two-photon laser ablation as a standard for determining functional outcomes following experimental treatments. In this assay, cell membranes are subjected to wounding with a two-photon laser, which causes the cell membrane to rupture and fluorescent dye to infiltrate the cell. The intensity of fluorescence within the cell can then be monitored to quantify the cell's ability to reseal itself. There are several alternative methods for assessing cell membrane response to injury, as well as great variation in the two-photon laser wounding approach itself, therefore, a single, unified model of cell wounding would beneficially serve to decrease the variation between these methodologies. In this article, we outline a simple two-photon laser wounding protocol for assessing cell membrane repair in vitro in both healthy and dysferlinopathy patient fibroblast cells transfected with or without a full-length dysferlin plasmid.

Cell membrane wounding via two-photon laser is a widely used method for assessing membrane resealing ability and can be applied to multiple cell types. Here, we describe a protocol for in vitro live-imaging of membrane resealing in dysferlinopathy patient cells following two-photon laser ablation.

Numerous pathophysiological insults can cause damage to cell membranes and, when coupled with innate defects in cell membrane repair or integrity, can ...

Measuring Transcellular Interactions through Protein Aggregation in a Heterologous Cell System

Protein interactions at cellular interfaces dictate a multitude of biological outcomes ranging from tissue development and cancer progression to synapse formation and maintenance. Many of these fundamental interactions occur in trans and are typically induced by heterophilic or homophilic interactions between cells expressing membrane anchored binding pairs. Elucidating how disease relevant mutations disrupt these fundamental protein interactions can provide insight into a myriad of cell biology fields. Many protein-protein interaction assays do not typically disambiguate between cis and trans interactions, which potentially leads to an overestimation of the extent of binding that is occurring in vivo and involve labor intensive purification of protein and/or specialized monitoring equipment. Here, we present an optimized simple protocol that allows for the observation and quantification of only trans interactions without the need for lengthy protein purifications or specialized equipment. The HEK cell aggregation assay involves the mixing of two independent populations of HEK cells, each expressing membrane-bound cognate ligands. After a short incubation period, samples are imaged and the resulting aggregates are quantified.

Here, we present an optimized protocol to rapidly and semiquantitatively measure ligand-receptor interactions in trans in a heterologous cell system using fluorescence microscopy.

Protein interactions at cellular interfaces dictate a multitude of biological outcomes ranging from tissue development and cancer progression to synapse ...

Neuroscience

Determining Membrane Protein Topology Using Fluorescence Protease Protection (FPP)

The correct topology and orientation of integral membrane proteins are essential for their proper function, yet such information has not been established for many membrane proteins. A simple technique called fluorescence protease protection (FPP) is presented, which permits the determination of membrane protein topology in living cells. This technique has numerous advantages over other methods for determining protein topology, in that it does not require the availability of multiple antibodies against various domains of the membrane protein, does not require large amounts of protein, and can be performed on living cells. The FPP method employs the spatially confined actions of proteases on the degradation of green fluorescent protein (GFP) tagged membrane proteins to determine their membrane topology and orientation. This simple approach is applicable to a wide variety of cell types, and can be used to determine membrane protein orientation in various subcellular organelles such as the mitochondria, Golgi, endoplasmic reticulum and components of the endosomal/recycling system. Membrane proteins, tagged on either the N-termini or C-termini with a GFP fusion, are expressed in a cell of interest, which is subject to selective permeabilization using the detergent digitonin. Digitonin has the ability to permeabilize the plasma membrane, while leaving intracellular organelles intact. GFP moieties exposed to the cytosol can be selectively degraded through the application of protease, whereas GFP moieties present in the lumen of organelles are protected from the protease and remain intact. The FPP assay is straightforward, and results can be obtained rapidly.

Determining Membrane Protein Topology Using Fluorescence Protease Protection FPP

Here, we present a protocol to determine the orientation and topology of integral membrane proteins in living cells. This simple protocol relies on selective protease sensitivity of chimeras between the protein of interest and GFP.

The correct topology and orientation of integral membrane proteins are essential for their proper function, yet such information has not been established ...

Determination of Plasma Membrane Partitioning for Peripherally-associated Proteins

This method provides a fast approach for the determination of plasma membrane partitioning of any fluorescently-tagged peripherally-associated protein using the profiles of fluorescence intensity across the plasma membrane. Measured fluorescence profiles are fitted by a model for membrane and cytoplasm fluorescence distribution along a line applied perpendicularly to the cell periphery. This model is constructed from the fluorescence intensity values in reference cells expressing a fluorescently-tagged marker for cytoplasm and with FM 4-64-labeled plasma membrane. The method can be applied to various cell types and organisms; however, only plasma membranes of non-neighboring cells can be evaluated. This fast microscopy-based method is suitable for experiments, where subtle and dynamic changes of plasma membrane-associated markers are expected and need to be quantified, e.g., in the analysis of mutant versions of proteins, inhibitor treatments, and signal transduction observations. The method is implemented in a multi-platform R package that is coupled with an ImageJ macro that serves as a user-friendly interface.

Here, we present a protocol to perform a quantitative analysis of the level of plasma-membrane association for fluorescently-tagged peripherally-associated protein. The method is based on the computational decomposition of membrane and cytoplasmic component of signal observed in cells labeled with plasma membrane fluorescent marker.

This method provides a fast approach for the determination of plasma membrane partitioning of any fluorescently-tagged peripherally-associated protein ...

Small-Scale Plasma Membrane Preparation for the Analysis of Candida albicans Cdr1-mGFPHis

The successful biochemical and biophysical characterization of ABC transporters depends heavily on the choice of the heterologous expression system. Over the past two decades, we have developed a yeast membrane protein expression platform that has been used to study many important fungal membrane proteins. The expression host Saccharomyces cerevisiae AD&#916;&#916; is deleted in seven major endogenous ABC transporters and it contains the transcription factor Pdr1-3 with a gain-of-function mutation that enables the constitutive overexpression of heterologous membrane protein genes stably integrated as single copies at the genomic PDR5 locus. The creation of versatile plasmid vectors and the optimization of one-step cloning strategies enables the rapid and accurate cloning, mutagenesis, and expression of heterologous ABC transporters. Here, we describe the development and use of a novel protease-cleavable mGFPHis double tag (i.e., the monomeric yeast enhanced green fluorescent protein yEGFP3 fused to a six-histidine affinity purification tag) that was designed to avoid possible interference of the tag with the protein of interest and to increase the binding efficiency of the His tag to nickel-affinity resins. The fusion of mGFPHis to the membrane protein ORF (open reading frame) enables easy quantification of the protein by inspection of polyacrylamide gels and detection of degradation products retaining the mGFPHis tag. We demonstrate how this feature facilitates detergent screening for membrane protein solubilization. A protocol for the efficient, fast, and reliable isolation of the small-scale plasma membrane preparations of the C-terminally tagged Candida albicans multidrug efflux transporter Cdr1 overexpressed in S. cerevisiae AD&#916;&#916;, is presented. This small-scale plasma membrane isolation protocol generates high-quality plasma membranes within a single working day. The plasma membrane preparations can be used to determine the enzyme activities of Cdr1 and Cdr1 mutant variants.

Small-Scale Plasma Membrane Preparation for the Analysis of Candida albicans Cdr1-mGFPHis

This article presents a small-scale plasma membrane isolation protocol for the characterization of Candida albicans ABC (ATP-binding cassette) protein Cdr1, overexpressed in Saccharomyces cerevisiae. A protease-cleavable C-terminal mGFPHis double tag with a 16-residue linker between Cdr1 and the tag was designed to facilitate the purification and detergent-screening of Cdr1.

The successful biochemical and biophysical characterization of ABC transporters depends heavily on the choice of the heterologous expression system. Over ...

A Fluorescence-Based Assay of Membrane Potential for High-Throughput Functional Study of Two Endogenous Ion Channels in Two Epithelial Cell Lines

Fluorescence-based studies are suitable for high-throughput plate reader assays of cells in culture. They have been commonly employed for drug discovery campaigns targeting recombinant ion channel proteins overexpressed in cells such as HEK-293 cells. However, there is increasing emphasis on the use of tissue-relevant cell lines for studying the effects of small molecule interventions. The following protocol describes the adaptation of a fluorescence-based membrane potential assay for the study of ion channels endogenously expressed in epithelial cell lines. The membrane potential assay details a high-throughput assay for chloride channel activity of the Cystic Fibrosis Transmembrane Conductance Regulator (CFTR) in two commonly studied epithelial cell lines, Caco-2 and Calu-3. In addition, this paper describes a novel application of this system to measure the activity of the Epithelial Sodium Channel (ENaC) in a high-throughput format in the same epithelial cell lines. Together, these fluorescence-based assays provide a robust and flexible platform for studying small molecule modulators, targeting two epithelial channels in a relevant cellular context.

This protocol describes a method for the study of electrogenic membrane proteins by measuring changes in membrane potential. This assay provides a platform for the functional readout of multiple ion channels endogenously expressed in epithelial cell lines.

Fluorescence-based studies are suitable for high-throughput plate reader assays of cells in culture. They have been commonly employed for drug discovery ...

High-throughput Measurement of Plasma Membrane Resealing Efficiency in Mammalian Cells

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