The authors would like to thank Emilie Josse for the critical review of the manuscript. This work was supported by the foundation &#34;La Ligue contre le Cancer&#34;, the &#34;Fondation ARC pour la Recherche sur le Cancer&#34;, and the &#34;Centre National de la Recherche Scienti&#64257;que&#34; (CNRS).

1. Preparation of Large Unilamellar Vesicles (LUVs)
<ol>
	<li>Prepare LUVs for their use as cell membrane mimics for fluorescence leakage assay.</li>
	<li>Mix with a Hamilton glass syringe phosphatidylcholine (DOPC, 786.11 g/mol), sphingomyelin (SM, 760.22 g/mol) and Cholesterol (Chol, 386.65 g/mol) at the molar ratio 4:4:2. The lipid solution is obtained from a stock solution of each lipid solubilized in a methanol/chloroform (3/1; volume/volume) solvent at 25 mg/mL in a 25 mL glass round-bottom flask. Based on 4 &#181;mol&#160;of DOPC, 4 &#181;mol&#160;of SM, and 2 &#181;mol&#160;of Chol, the lipid solution is obtained from stock solution by mixing 126 &#181;L, 117 &#181;L, and 31 &#181;L, respectively. 
	CAUTION: Methanol is a toxic and inflammable solvent and chloroform is toxic and carcinogenic. Both should be handled with the appropriate protection under a hood.</li>
	<li>Evaporate methanol/chloroform using a rotary evaporator under vacuum during 45-60 min at 60 &#176;C until formation of a dried lipid film.</li>
	<li>Prepare two stock HEPES buffer solutions. Prepare HEPES buffer 1 by mixing 20 mM HEPES (238.3 g/mol) and 75 mM NaCl (58.44 g/mol) and adjust pH to 7.4. Prepare HEPES buffer 2 by mixing 20 mM HEPES and 145 mM NaCl and adjust pH to 7.4. HEPES buffers can be stored at 4 &#176;C for 1 month. 
	NOTE: It is recommended to check the osmolarity of the buffers using an osmometer.</li>
	<li>Prepare lipid hydration solution by dissolving membrane impermeable fluorescent dye-quencher couple, 8-aminonaphthalene-1, 3, 6-trisulfonic acid, disodium salt at 12.5 mM (ANTS, 427.33 g/mol) and p-xylene-bispyridinium bromide at 45 mM (DPX, 422.16 g/mol) in HEPES buffer solution. Mixing ANTS with DPX leads to a yellow-colored solution. To achieve the concentrations of 12.5 mM of ANTS and 45 mM of DPX, dissolve 21.4 mg and 76 mg, respectively in 4 mL of HEPES buffer 1. 
	NOTE: Lipid hydration solution can be stored for 2 weeks at 4 &#176;C by wrapping the tube with aluminum foil.</li>
	<li>Reconstitute multilamellar vesicles (MLV) by resuspending the dried lipid film with 1 mL of the lipid hydration solution and by vortexing until dissolution of the dried lipid film. Ensure that the solution is completely solubilized as small lipid aggregates will negatively impact the preceding steps. Also, check the wall of the glass round-bottom flask to ensure that there is no remaining lipid film. 
	NOTE: The solution will appear opalescent and light yellow after the solubilization.</li>
	<li>Subject the vesicles to five freeze/thaw cycles to obtain unilamellar vesicles. Perform each cycle by putting the glass round-bottom flask for 30 s in liquid nitrogen for freezing step, then leaving it in a water bath for 2 min for thawing step. 
	NOTE: The temperature of the bath water should be 5-10&#176;C higher than the melting temperature of the lipids.</li>
	<li>Prepare lipid extruder by inserting two filter supports preliminary humidified with HEPES buffer in each polytetrafluoroethylene (PTFE) extruder piece placed in the metal extruder canister.</li>
	<li>Put a HEPES humidified polycarbonate membrane (0.1 &#181;m pore size, 25 mm diameter) on the top of one filter support.</li>
	<li>Assemble the two metal extruder canisters and screw them.</li>
	<li>Place the assembled extruder in the holder and introduce a 1 mL syringe in the appropriate hole at the extremity of each polytetrafluoroethylene extruder piece. Extrusion corresponds to the passage of the liquid tested from one syringe to the other through the polycarbonate membrane.</li>
	<li>Test the extruder with 1 mL of HEPES buffer loaded in one of the 1 mL syringe to ensure that there are no leaks or problems.</li>
	<li>Replace the 1 mL HEPES buffer with the MLV sample.</li>
	<li>Perform extrusion by passaging the MLV sample from one syringe to the other through the polycarbonate membrane at least 21 times to obtain uniform LUVs of same size. 
	&#8203;NOTE: Extrusion should be performed at a temperature higher than the melting temperature of the lipid mixture.</li>
</ol>
2. Purification of LUVs
<ol>
	<li>Prepare a column purification to remove non-encapsulated ANTS and DPX excess.</li>
	<li>Introduce cross-linked dextran gel (G-50) resuspended in aqueous medium with 0.01% NaN3 (65 g/mol) in a liquid chromatography column (Luer Lock, Non-jacketed, 1.0 cm x 20 cm, bed volume 16 mL) up to 1 cm below the top of the colorless part of the column.</li>
	<li>Open the tap and let the liquid flow to settle the cross-linked dextran gel.</li>
	<li>Wash the column by eluting with 20 mL of HEPES buffer 2 and discard the output flow of the column.</li>
	<li>Close the tap once the dead volume of solvent above the column is minimized (&#60;100 &#181;L) but sufficient to avoid any drying of the cross-linked dextran gel.</li>
	<li>Place the freshly extruded LUVs (yellow) on the column and let them enter into the cross-linked dextran gel.</li>
	<li>Continuously add HEPES buffer 2 to the column to perform the LUV purification.</li>
	<li>Elute approximately 2 mL of HEPES buffer 2 (do not forget to regularly fill the top of the column to avoid drying the cross-linked dextran gel): the free yellow ANTS and DPX solution migrates slower than the liposomes.</li>
	<li>Start collecting purified LUVs in tubes (1.5 mL).</li>
	<li>Observe the drops of eluent from the column and when they become opalescent, they contain liposomes. Change the tube to recover the LUV-containing fraction.</li>
	<li>Elute until the drops are no longer opalescent (~1 mL). Afterwards, elute another 0.5 mL in a separate fraction and then stop eluting. 
	NOTE: Standards are now available in a wide range of molecular weights, as kits or individual molecular weights to calibrate the elution volume of the LUVs.</li>
	<li>Wrap the tubes with the LUVs in aluminum foil to avoid bleaching of the fluorescence dye.</li>
	<li>Wash the column with 20 mL HEPES buffer 2.</li>
	<li>The LUVs can then be stored for a week at 4 &#176;C. 
	&#8203;NOTE: As LUV&#160;stability might depend on LUV&#160;concentration and composition, as well as on ionic strength, the size of the LUVs should be controlled using a dynamic light scattering (DLS) instrument (see section 4. Characterization of LUV Size and Homogeneity) before each test.</li>
</ol>
3. Quantifying the concentration of LUVs
<ol>
	<li>Estimate LUV concentration by a phospholipid quantification kit, which enables the evaluation of choline concentration19. This assay might be applied when phospholipids with choline containing polar head is substantial (&#62;50% of the LUVs).</li>
	<li>Prepare the color reagent by dissolving 18 mg of chromogen substrate in 3 mL of buffer provided.</li>
	<li>Load a polystyrene cuvette, 10 x 10 x 45 mm, with 3 mL of color reagent.</li>
	<li>Use the pure color reagent as blank condition (Blank). Add 20 &#181;L of LUV&#160;sample (Test) or 20 &#181;L of standard solution of known choline concentration (Standard).</li>
	<li>Mix well and incubate for 5 min at 37 &#176;C all conditions (Blank, Test, and Standard).</li>
	<li>Measure the absorbance (optical density, OD) of the test sample and standard solution with the blank solution as the control at 600 nm with a spectrophotometer.</li>
	<li>Check the OD values which enable to estimate the lipid concentration of the LUVs, C[LUV], in choline equivalent compared to the standard of known concentration.</li>
	<li>Perform the calculation using the following equation: 
	C[LUV] (mol / l) = (OD Sample / OD Standard) x C[Standard] (mol / l) 
	&#8203;NOTE: The phospholipid quantification kit provided a Choline Chloride (139.6 mg/l) standard solution at 54 mg/dL corresponding to molar concentration of C[Standard] = 3.87 mmol/L. OD Sample and OD Standard are the absorbances measured at 600 nm for the LUV and Choline solutions, respectively.</li>
</ol>
4. Characterization of LUV size and homogeneity
<ol>
	<li>Perform a measurement using a DLS instrument in order to determine the LUV size (in nm) and polydispersity index (PdI).</li>
	<li>Program the appropriated &#34;standard operation procedure&#34; (SOP) by indicating the viscosity of the solvent/buffer and the used cuvette.</li>
	<li>Place 500 &#181;L of the LUV solution in a polystyrene semi-micro cuvette.</li>
	<li>Insert the polystyrene semi-micro cuvette in a DLS instrument.</li>
	<li>At room temperature, measure the size distribution in terms of mean size (Z-average) of the particle distribution and of homogeneity (polydispersity index, PdI).</li>
	<li>All the results are obtained from two independent measurements performed each in three repetitive cycles. 
	&#8203;NOTE: Standard values for LUVs will be a mean size of 137 &#177; 7 nm with a PdI of 0.149 &#177; 0.041.</li>
</ol>
5. Preparing peptide solutions
<ol>
	<li>Prepare a stock solution of the peptide, which should be analyzed for the leakage assay.</li>
	<li>Dissolve peptide powder (&#62;95% purity) in pure water (e.g., 1 mg peptide in 500 &#181;L pure water). 
	NOTE: It is recommended to dilute peptides in pure water and to avoid dimethyl sulfoxide (DMSO) solubilization, which could induce artifacts (e.g., membrane permeabilization20).</li>
	<li>Vortex the peptide solution for 5 s.</li>
	<li>Sonicate the peptide solution in a water sonication bath for 5 min and then centrifuge for 5 min at 12,225 x g. Collect the supernatant for concentration determination.</li>
	<li>Measure the absorbance at 280 nm of three independent peptide dilutions and then calculate peptide concentration using its molar extinction coefficient &#949; (depending on tryptophan and tyrosine content in the peptide sequence) and Beer-Lambert rule. 
	NOTE: If the peptide contains tryptophan and tyrosine, the molar extinction coefficient &#949; is computed on the basis of Tryptophan &#949; = 5,690 M-1cm-1 and Tyrosine &#949; = 1,280 M-1cm-1. If the peptide sequence contains no tryptophan or tyrosine, other colorimetric assay could be performed to measure the concentration (e.g., BCA or Bradford).</li>
	<li>Dilute the peptide solution in pure water to a final solution of 100 &#181;M and store at 4 &#176;C. 
	&#8203;NOTE: In pure water, no peptide degradation occurs during the 4 &#176;C storage. However, peptide concentration should be measured every 2 weeks to ensure that no water evaporation occurs.</li>
</ol>
6. Fluorescence leakage assay
<ol>
	<li>Fluorescence leakage assay is measured on a spectrofluorometer at room temperature. Excitation and emission wavelength are fixed at Ex = 360 nm &#177; 3 nm and Em = 530 nm &#177; 5 nm, respectively.</li>
	<li>Dilute LUVs in 1 mL HEPES buffer 2 to a final concentration of 100 &#181;M in a quartz fluorescence cuvette. Add a magnetic stirrer to homogenize the solution during experiment.</li>
	<li>Measure the LUVs alone during the first 100 s, between t = 0 s and t = 99 s in order to access the background fluorescence. 
	NOTE: LUVs alone could also be measured during the whole experiment (15 min) in order to access background fluorescence and potential leaks.</li>
	<li>Thereafter, measure leakage as an increase in fluorescence intensity upon addition of aliquots of peptide solution for the next 900 s (15 min). This protocol is carried out for each concentration of peptide tested from 0.1 &#181;M to 2.5 &#181;M.</li>
	<li>Finally, 100% fluorescence was achieved by solubilizing the LUVs by addition of 1 &#181;L of Triton X-100 (0.1%, v/v), resulting in the completely unquenched probe between t = 1,000 s and t = 1,100 s.</li>
</ol>
7. Quantification of the leakage
<ol>
	<li>Suppress values obtained after t = 1,090 s in order to keep the same number of points for each tested condition.</li>
	<li>Calculate the minimal fluorescence, Fmin, by making the average of 50 points between t = 0 s and t = 49 s (LUVs alone).</li>
	<li>Calculate the maximal fluorescence, Fmax, by making the average of 50 points between t = 1,041 s and t = 1,090 s (LUVs with Triton X-100).</li>
	<li>Calculate the leakage percentage (%Leak) at each time point (t = x), according to the following equation: 
	%Leak(t=x) = (F(t=x) - Fmin) / (Fmax - Fmin) x 100</li>
	<li>Calculate the average and standard deviation for values obtained with different LUV&#160;preparation (n &#8805; 2) for the same condition.</li>
	<li>Plot the leakage percentage, %Leak(t=x), in function of time (s).</li>
</ol>

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C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Determining+the+effects+of+membrane-interacting+peptides+on+membrane+integrity.">Determining the effects of membrane-interacting peptides on membrane integrity.</a> Cell-Penetrating Peptides. 1324, 89-106 (2015).</li><li>B&#225;r&#225;ny-Wallje, E., Gaur, J., Lundberg, P., Langel, U., Gr&#228;slund, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Differential+membrane+perturbation+caused+by+the+cell+penetrating+peptide+Tp10+depending+on+attached+cargo.">Differential membrane perturbation caused by the cell penetrating peptide Tp10 depending on attached cargo.</a> FEBS Letters. 581 (13), 2389-2393 (2007).</li><li>van Rooijen, B. D., Claessens, M. M. A. E., Subramaniam, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Membrane+permeabilization+by+oligomeric+&#945;-Synuclein:+in+search+of+the+mechanism.">Membrane permeabilization by oligomeric &#945;-Synuclein: in search of the mechanism.</a> PloS One. 5 (12), 14292 (2010).</li><li>Hassane, F. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Insights+into+the+cellular+trafficking+of+splice+redirecting+oligonucleotides+complexed+with+chemically+modified+cell-penetrating+peptides.">Insights into the cellular trafficking of splice redirecting oligonucleotides complexed with chemically modified cell-penetrating peptides.</a> Journal of Controlled Release: Official Journal of the Controlled Release Society. 153 (2), 163-172 (2011).</li><li>Asciolla, J. J., Renault, T. T., Chipuk, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Examining+BCL-2+family+function+with+large+unilamellar+vesicles.">Examining BCL-2 family function with large unilamellar vesicles.</a> Journal of Visualized Experiments: JoVE. (68), e4291 (2012).</li><li>Grewer, C., Gameiro, A., Mager, T., Fendler, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electrophysiological+characterization+of+membrane+transport+proteins.">Electrophysiological characterization of membrane transport proteins.</a> Annual Review of Biophysics. 42 (1), 95-120 (2013).</li></ol>

The authors have no conflicts of interest.

The presented fluorescence leakage assay is a simple and fast method to address membrane destabilization by cell-penetrating peptide. Easy to do, it also enables an indirect comparison between different membrane-interacting peptides or other membrane-interacting molecules. Concerning critical steps of the protocol, as this assay provides relative values between the baseline (LUVs alone) and maximal fluorescence release (Triton condition), we usually evaluate the concentration of LUVs using the phospholipid quantification kit, which only estimates choline contribution of the LUVs. However, it is also possible to include a more accurate measurement of LUV&#160;concentration by determining the total phosphorus content by acid digestion as described by Rouser and colleagues27 or by Bartlett28 or by colorimetric method using ammonium ferrothiocyanate complexing phospholipids29. In our hands, it does not have an impact on the relative quantification of leakage and on the comparison between membrane-interacting peptides.
With regard to the use of LUVs, one should also notice the importance to control the state of the baseline of LUV alone in order to check whether they are usable and if they are already showing leaks (characterized by a continuous increase of fluorescence). In the same way, it is important to measure the maximal fluorescence release after each peptide incubation to ensure that no false-negative results were recorded due to an undesired quenching process (e.g., peptide/dye quenching).
In general, according to DLS characterization, LUVs are quite stable for 1 week and should not display a too high fluorescence background. In this context, purification of ANTS/DPX loaded liposomes by gel filtration is also a key point since enabling formation of LUVs without having residual fluorescence that might contribute to the background or produce fluorescence overestimation. Additionally, it is strongly recommended to calibrate the protocol by including a specific positive control that can always give the same leakage percentage. This constitutes an internal control through the different measurements, which might reinforce characterization of the LUVs for each test and increase statistics.
Although the fluorescence leakage assay might provide a fast comparison in membrane-destabilization by CPPs, it is however limited in the interpretation of peptide-membrane interactions since some peptides are able to interact with lipid bilayer without inducing any membrane disturbance or leakage. For these peptides, it is highly recommended to use additional experiments with specific fluorescence markers enabling FRET30.
It should be also noted that other fluorescence dye/quencher pairs (e.g., Tb3+/DPA31) could be used. Both the methods have their inconveniences: Terbium (III) is not very soluble in water and cannot be used in the presence of phosphate, whereas the ANTS quenching is not a linear function of DPX concentration. Furthermore, other self-quenching materials such as 70 mM Calcein solution32 or dextran-PTS33 could be handled depending on the membrane defects provoked by the analyzed peptide or compound.
Finally, the main advantage of this fluorescence leakage assay is the ability to test a multitude of potential membrane interacting molecules as well as different membrane compositions, such as endosomal membrane mimics [e.g., dioleoylphosphatidylcholine (DOPC)/dioleoyl-phospha-tidylethanolamine (DOPE)/phosphatidylinositol from soybean (PI)/bis(monooleoylglycero) phosphate (LBPA) at a molar ratio of 5:2:1:2]34, mitochondrial membrane mimics [e.g., 46.5% DOPC, 28.5% phosphatidylethanolamine (PE), 9% phosphatidylinositol (PI), 9% phosphatidylserine (PS), and 7% cardiolipin (CL)35 or any other desired lipid bilayer composition. However, one will first have to ensure stability of the vesicles (no LUV fusion, aggregation, or precipitation, no lipidic falling out of suspension, no negative membrane curvature, etc.) having a constant mean size close to 100 nm during experiment and storage.
In addition, the advantage of this method compared to extracted membranes (red blood cells, mitochondria, etc.) is the use of purified well-characterized lipids in the absence of proteins. The control of the lipid bilayer composition (plasma, endosomal, or mitochondrial membrane) also enables the insertion of specific membrane proteins (proton pump). Moreover, the ability to control both LUVs internal and external environments allows a clear interpretation of membrane disturbance/leakage. Compared to black-lipid membrane experiments36, which can also visualize membrane permeabilization events, this leakage protocol provides a more simple screening method of membrane-active peptides.
In conclusion, this simple method favors a rapid identification of strong peptide/membrane interactions leading to membrane destabilization. It may be applied to investigate the mechanism of internalization of CPPs and in a more general approach of &#34;membrane-active peptides&#34; such as fusiogenic or antimicrobial peptides.
In general, the characterization of the main route used by CPPs to reach the cytoplasm is very complex and requires several distinct approaches, from biophysics to cellular biology. For example, the investigation of WRAP internalization revealed a balance between different mechanisms to enter the cells (endocytosis versus direct translocation) and the fluorescence leakage assay, associated to other methods, has contributed to support a direct penetration process7.

After their discovery in the nineties, cell-penetrating peptides (CPPs) were developed to promote an efficient cellular delivery of cargoes through the plasma membrane1,2. CPPs are usually short peptides, generally 8 to 30 amino acids, having a wide variety of origins. They were first defined as &#34;direct-translocating&#34; carriers, meaning they were able to cross the plasma membrane and to transfer a cargo into cells independently of any endocytotic pathway neither energy requirement nor receptor involvement. However, further investigations revealed that these first observations mainly came from fluorescence overestimation due to the experimental artefact and/or to fixation protocols using methanol3. Nowadays, it is widely accepted that CPP uptake takes place by both endocytosis and energy-independent translocation4,5,6,7 depending on different parameters such as the nature of cargo, the used link between CPP and cargo, the studied cell line, etc.
CPPs can be used as transfection agents according to two strategies, either involving a chemical link (covalent strategy) or electrostatic/hydrophobic interactions (non-covalent strategy) between the CPP and its cargo8,9,10,11. Although both strategies have shown their efficiency in the cell transfer of several cargoes, the understanding of the mechanism of internalization by CPPs is still under controversy and the balance between endocytosis pathways or direct penetration is still difficult to measure12,13. Although a set of experimental tools and strategies are available to clearly address the involvement of endocytic processes, the direct translocation seems, however, more difficult to characterize since it implies more discrete interactions with plasma membrane components. Biological membranes are usually composed of numerous components, from phospholipids to membrane proteins, which might vary according to the cellular type and/or the environment (stress conditions, cell division, etc.). This diversity of composition, and consequently the absence of a universal cellular membrane model does not enable studies in a single way. However, to circumvent these limitations step-by-step approaches were developed with artificial membrane or membrane extracts. From small unilamellar vesicles to monolayer approaches, every model was clearly pertinent to answer specific questions14,15. Among them, large unilamellar vesicles (LUVs) constitute an appropriate membrane mimicking model to study peptide/membrane interactions as being a key point in the internalization process.
In this context, the following protocol describes the investigation of the effects of peptides and peptide/membrane interactions on LUVs integrity through the monitoring of both an anionic fluorescent dye and its corresponding poly-cationic quencher encapsulated in liposomes. This tool is used to study CPP/membrane interactions in order to understand whether they are able to perform a direct membrane translocation. Although usually applied to compare different membrane-interacting peptides, this LUV fluorescence leakage assay could also be used for investigating both CPPs-cargo conjugates (covalent strategy) and CPP:cargo complexes (non-covalent strategy).
The present protocol will hence be first exemplified with the recently developed tryptophan (W)- and arginine (R)-rich Amphipathic Peptides (WRAP)16. WRAP is able to form peptide-based nanoparticles to rapidly and efficiently deliver small interfering RNA (siRNA) in several cell lines16. The fluorescence leakage properties of WRAP peptide alone or siRNA-loaded WRAP-based nanoparticles were monitored to characterize their mechanism of cellular internalization. We showed that their mechanism of internalization mainly involved direct translocation7. In a second example, the WRAP peptide was covalently conjugated to the protein/protein interfering peptide iCAL36 (WRAP-iCAL36)17 and its ability to destabilize membranes was compared in a fluorescence leakage assay to iCAL36 coupled to Penetratin18 (Penetratin-iCAL36), another CPP.
Finally, the advantages and limitations of the method will be discussed both from a technological point of view and with respect to biological relevance.

<table><tbody><tr><td>25 mL glass round-bottom flask</td><td>Pyrex</td><td></td><td></td></tr><tr><td>8-aminonaphthalene-1, 3, 6-trisulfonic acid, disodium salt (ANTS)</td><td>Invitrogen</td><td>A350</td><td>Protect from light&amp;nbsp;</td></tr><tr><td>Chloroform&amp;nbsp;</td><td>Sigma-Aldrich</td><td>288306</td><td></td></tr><tr><td>Cholesterol</td><td>Sigma-Aldrich</td><td>C8667</td><td></td></tr><tr><td>DOPC (dioleoylphosphatidylcholine)</td><td>Avanti Polar</td><td>850375P</td><td>Protect from air</td></tr><tr><td>Extruder</td><td>Avanti Polar</td><td>610000</td><td></td></tr><tr><td>Fluorimeter</td><td>PTI Serlabo</td><td></td><td></td></tr><tr><td>50 &amp;micro;L glass syringe</td><td>Hamilton</td><td>705N</td><td></td></tr><tr><td>HEPES</td><td>Sigma-Aldrich</td><td>H3375</td><td></td></tr><tr><td>LabAssay Phospholipid&amp;nbsp;</td><td>WAKO&amp;nbsp;</td><td>296-63801</td><td></td></tr><tr><td>liquid chromatography column</td><td>Sigma-Aldrich</td><td></td><td></td></tr><tr><td>Methanol</td><td>Carlo Erba</td><td>414902</td><td></td></tr><tr><td>Nuclepore polycarbonate membrane (0.1 &amp;micro;m pore size, 25 mm diameter)</td><td>Whatman</td><td>800309</td><td></td></tr><tr><td>polystyrene cuvette, 10 x 10 x 45 mm</td><td>Grener Bio-One</td><td>614101</td><td></td></tr><tr><td>polystyrene semi-micro cuvette, DLS</td><td>Fisher Scientific</td><td>FB55924</td><td></td></tr><tr><td>p-xylene-bispyridinium bromide (DPX)</td><td>Invitrogen</td><td>X1525</td><td>Protect from light&amp;nbsp;</td></tr><tr><td>quartz fluorescence cuvette</td><td>Hellma</td><td>109.004F-QS</td><td></td></tr><tr><td>rotavapor system&amp;nbsp;</td><td>Heidolph</td><td>Z334898</td><td></td></tr><tr><td>Sephadex G-50 resin</td><td>Amersham</td><td>17-0042-01</td><td></td></tr><tr><td>Sodium azide (NaN&lt;sub&gt;3&lt;/sub&gt;)</td><td>Sigma-Aldrich</td><td>S2002</td><td></td></tr><tr><td>Sodium chlorid (NaCl)</td><td>Sigma-Aldrich</td><td>S5886</td><td></td></tr><tr><td>Sonicator bath USC300T</td><td>VWR</td><td>142-6001</td><td></td></tr><tr><td>Sphingomyelin</td><td>Avanti Polar</td><td>860062P</td><td>Protect from air</td></tr><tr><td>Triton X-100&amp;nbsp;</td><td>Eromedex</td><td>2000-B</td><td></td></tr><tr><td>Zetaziser NanoZS&amp;nbsp;</td><td>Malvern</td><td>ZEN3500</td><td></td></tr></tbody></table>

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 principle of the fluorescence leakage assay is shown in the Figure 1. In detail, large unilamellar vesicles (LUVs) encapsulating a fluorescent dye and a quencher (no fluorescence signal) are treated with the biomolecule of interest. Due to the interaction of the peptide with lipid membranes, which could imply membrane permeability, reorganization or even rupture, the fluorescence dye and the quencher are released from the LUVs. Subsequent dilutions in the buffer results in an increased fluorescence signal.
Although this scheme displays a test with free peptides, the advantage of the system lies in the ability to also test cargo-conjugated peptides, peptide-based nanoparticles or other biopolymers, which are suspected to destabilize lipid membranes. Although a preliminary optimization of the protocol especially with regards to the molecules tested is required, this fluorescence leakage assay might be extended to a huge variety of membrane-interacting components. In the present protocol, we show some results obtained with the CPPs and their complexed (non-covalent strategy) or conjugated (covalent strategy) forms. The following examples imply WRAP alone as well as siRNA-loaded WRAP-based nanoparticles and two different peptide-conjugates (WRAP-iCAL3617 and Penetratin-iCAL36).
With regard to non-covalent strategy, fluorescence leakage assay with peptides and with their corresponding siRNA-loaded nanoparticles in the presence of LUVs are displayed in Figure 2. The vesicles are composed of a mixture of DOPC/SM/Chol (4:4:2) reflecting the plasma membrane as described in Konate et al.21 and are usually used to directly evaluate the possibility of lipid bilayer interaction and/or transduction properties of free WRAP and WRAP-based nanoparticles7. In the absence of peptides, no leakage is observed (baseline during the first 100 s). Addition of free WRAP on the LUVs induces a significant increase of fluorescence revealing an important LUV leakage and ANTS release. After 15 min, a leakage of 67.8% &#177; 0.4% compared to the Triton condition (positive control) is obtained at the used concentration (2.5 &#181;M) of WRAP peptide. It should be noted here that several different concentrations have also been tested and revealed a dose-dependent fluorescence increase, corresponding to a dose-dependent LUV leakage7. In contrast, when WRAP is assembled at the same concentration with siRNA to form peptide-based nanoparticles, the leakage is 1.5-fold weaker (40.5% &#177; 0.5%) compared to the free peptide (Figure 2). Similar leakage values have been reported for the RICK peptide (60%) or the RICK:siRNA nanoparticles (28%)22. The difference in values between free peptide compared to nanoparticles might be explained by the fact that, when engaged in the nanoparticles, a substantial part of the peptide is involved in direct interactions with the siRNA, reducing the peptide availability for interactions with lipids.
Concerning the covalent strategy, fluorescence leakage assays with CPP-conjugates in the presence of LUVs are shown in Figure 3. With the same LUV composition [DOPC/SM/Chol (4:4:2)], two conjugated peptides are applied: WRAP-iCAL36 and Penetratin-iCAL36. As previously noticed, no leakage is observed in the absence of peptides. 15 min after injection of 2.5 &#181;M of Penetratin-iCAL36, no significant fluorescence increase is detected (2.3% &#177; 0.7%), whereas an addition of 2.5 &#181;M of WRAP-iCAL36 induces a net leakage characterized by very strong fluorescence signal (85.8% &#177; 11.1%)17. These observations indicate that for some peptides, or conjugates, no fluorescence leakage might occur, suggesting no peptide/membrane interactions or no lipid bilayer disturbance. This is in accordance with previously published results showing that Penetratin as well as Tat were not able to destabilize membranes23,24,25. It should be highlighted that the membrane destabilization properties of a CPP could change depending on the coupled cargo26.
Furthermore, although WRAP-iCAL36 caused a strong leakage, additional studies do not reveal specific cellular internalization, indicating that these conjugates remained inside the lipids bilayer of the plasma membrane17. In contrast to the WRAP nanoparticle, we suppose that the WRAP-cargo conjugate is able to destabilize the LUV membrane by sticking between the lipid chains or by forming pores.
These results indicate that the fluorescence leakage assay might reveal the ability of some CPPs to develop peptide/membrane interactions, which could lead to a more or less pronounced membrane permeability. Moreover, these interactions might occur whatever the strategy of cargoes delivery (nanoparticles versus conjugates). Conversely, this method does not discriminate whether a CPP, which does not induce any fluorescence leakage, might still interact with the bilayer or biological membrane of the lipids. This kind of behavior requires additional approaches such as zeta-potential measurements, FRET between peptide and membrane, or tryptophan fluorescence experiments to mention only a few examples.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62028/62028fig01.jpg" /> 
Figure 1: Principle of the fluorescence leakage assay. LUVs were loaded with a fluorescent dye (ANTS) in white and its corresponding quencher (DPX) in grey. In the absence of peptides (in red), no fluorescence signal was observed because ANTS fluorescence was quenched by DPX. Addition of peptides on the LUVs induced membrane permeability and the subsequent release of both ANTS and DPX resulting in a significant increase in ANTS fluorescence (yellow). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62028/62028fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62028/62028fig02.jpg" /> 
Figure 2: Fluorescence leakage assay with free WRAP and WRAP:siRNA nanoparticles. Peptide alone and siRNA-loaded nanoparticles were applied on LUVs at the peptide concentration of 2.5 &#181;M. WRAP-based nanoparticles were formulated at a peptide:siRNA molar ratio of 20 (R 20) as described in Konate et al.7,16. Black arrows show injections of peptide (nanoparticle) and Triton X-100, respectively. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62028/62028fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62028/62028fig03.jpg" /> 
Figure 3: Fluorescence leakage assay with WRAP-iCAL36 and Penetratin-iCAL36 conjugates. Conjugates were applied on LUVs at the concentration of 2.5 &#181;M. Black arrows show injections of peptide and Triton X-100, respectively.&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62028/62028fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Fluorescent Leakage Assay to Investigate Membrane Destabilization by Cell-Penetrating Peptide at JoVE.com

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

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 ...

fluorescent-leakage-assay-to-investigate-membrane-destabilization

Nanyang Technological University

Research

JoVE Journal

Biochemistry

6.1K Views.  INSERM U1046 - CNRS UMR 9214 - University of Montpellier. 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.

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

Construction of Cyclic Cell-Penetrating Peptides for Enhanced Penetration of Biological Barriers

Cancer has been a grand challenge in global health. However, the complex tumor microenvironment generally limits the access of therapeutics to deeper tumor cells, leading to tumor recurrence. To conquer the limited penetration of biological barriers, cell-penetrating peptides (CPPs) have been discovered with excellent membrane translocation ability and have emerged as useful molecular transporters for delivering various cargoes into cells. However, conventional linear CPPs generally show compromised proteolytic stability, which limits their permeability across biological barriers. Thus, the development of novel molecular transporters that can penetrate biological barriers and exhibit enhanced proteolytic stability is highly desired to promote drug delivery efficiency in biomedical applications. We have previously synthesized a panel of short cyclic CPPs with aromatic crosslinks, which exhibited superior permeability in cancer cells and tissues compared to their linear counterparts. Here, a concise protocol is described for the synthesis of the fluorescently labeled cyclic polyarginine R8 peptide and its linear counterpart, as well as key steps for investigating their cell permeability.

This protocol describes the synthesis of cyclic cell-penetrating peptides with aromatic cross-links and the evaluation of their permeability across biological barriers.

Cancer has been a grand challenge in global health. However, the complex tumor microenvironment generally limits the access of therapeutics to deeper ...

Chemistry

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 ...

Immunology and Infection

Measuring Peptide Translocation into Large Unilamellar Vesicles

There is an active interest in peptides that readily cross cell membranes without the assistance of cell membrane receptors1. Many of these are referred to as cell-penetrating peptides, which are frequently noted for their potential as drug delivery vectors1-3. Moreover, there is increasing interest in antimicrobial peptides that operate via non-membrane lytic mechanisms4,5, particularly those that cross bacterial membranes without causing cell lysis and kill cells by interfering with intracellular processes6,7. In fact, authors have increasingly pointed out the relationship between cell-penetrating and antimicrobial peptides1,8. A firm understanding of the process of membrane translocation and the relationship between peptide structure and its ability to translocate requires effective, reproducible assays for translocation. Several groups have proposed methods to measure translocation into large unilamellar lipid vesicles (LUVs)9-13. LUVs serve as useful models for bacterial and eukaryotic cell membranes and are frequently used in peptide fluorescent studies14,15. Here, we describe our application of the method first developed by Matsuzaki and co-workers to consider antimicrobial peptides, such as magainin and buforin II16,17. In addition to providing our protocol for this method, we also present a straightforward approach to data analysis that quantifies translocation ability using this assay. The advantages of this translocation assay compared to others are that it has the potential to provide information about the rate of membrane translocation and does not require the addition of a fluorescent label, which can alter peptide properties18, to tryptophan-containing peptides. Briefly, translocation ability into lipid vesicles is measured as a function of the Foster Resonance Energy Transfer (FRET) between native tryptophan residues and dansyl phosphatidylethanolamine when proteins are associated with the external LUV membrane (Figure 1). Cell-penetrating peptides are cleaved as they encounter uninhibited trypsin encapsulated with the LUVs, leading to disassociation from the LUV membrane and a drop in FRET signal. The drop in FRET signal observed for a translocating peptide is significantly greater than that observed for the same peptide when the LUVs contain both trypsin and trypsin inhibitor, or when a peptide that does not spontaneously cross lipid membranes is exposed to trypsin-containing LUVs. This change in fluorescence provides a direct quantification of peptide translocation over time.

This protocol details a method for the quantitative measure of peptide translocation into large unilamellar lipid vesicles. This method also provides information about the rate of membrane translocation and can be used to identify peptides that efficiently and spontaneously cross lipid bilayers.

There is an active interest in peptides that readily cross cell membranes without the assistance of cell membrane receptors1.  Many of these are referred ...

Biology

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and the inherent instability of many membrane proteins once solubilized in detergents. A protocol is described that combines ligation independent cloning of membrane proteins as GFP fusions with expression in Escherichia coli detected by GFP fluorescence. This enables the construction and expression screening of multiple membrane protein/variants to identify candidates suitable for further investment of time and effort. The GFP reporter is used in a primary screen of expression by visualizing GFP fluorescence following SDS polyacrylamide gel electrophoresis (SDS-PAGE). Membrane proteins that show both a high expression level with minimum degradation as indicated by the absence of free GFP, are selected for a secondary screen. These constructs are scaled and a total membrane fraction prepared and solubilized in four different detergents. Following ultracentrifugation to remove detergent-insoluble material, lysates are analyzed by fluorescence detection size exclusion chromatography (FSEC). Monitoring the size exclusion profile by GFP fluorescence provides information about the mono-dispersity and integrity of the membrane proteins in different detergents. Protein: detergent combinations that elute with a symmetrical peak with little or no free GFP and minimum aggregation are candidates for subsequent purification. Using the above methodology, the heterologous expression in E. coli of SED (shape, elongation, division, and sporulation) proteins from 47 different species of bacteria was analyzed. These proteins typically have ten transmembrane domains and are essential for cell division. The results show that the production of the SEDs orthologues in E. coli was highly variable with respect to the expression levels and integrity of the GFP fusion proteins. The experiment identified a subset for further investigation.

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

A streamlined approach to screening for the expression of recombinant membrane proteins in Escherichia coli based on fusion to green fluorescent protein is presented.

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and ...

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 ...

Assays for the Degradation of Misfolded Proteins in Cells

Protein misfolding and aggregation are associated with various neurodegenerative diseases. Cellular mechanisms that recognize and degrade misfolded proteins may serve as potential therapeutic targets. To distinguish degradation of misfolding-prone proteins from other mechanisms that regulate their levels, one important method is to measure protein half-life in cells. However, this can be challenging because misfolding-prone proteins may exist in different forms, including the native form and misfolded forms of distinct characteristics. Here we describe assays to examine the half-life of misfolded proteins in mammalian cells using a highly aggregation-prone protein, Ataxin-1 with an extended polyglutamine (polyQ) stretch, and a conformationally unstable luciferase mutant as models. Cycloheximide chase is combined with cell fractionation to examine the turnover rate of misfolding-prone proteins in various cellular fractions. We further depict a fluorescence-based assay using an enhanced green fluorescence protein (EGFP)-fusion of the luciferase mutant, which can be adapted for high throughput screening on a microplate-reader.

This report describes protocols for measuring degradation rates of misfolded proteins by either western blot or fluorescence-based assays. The methods can be applied to analysis of other misfolded proteins and for high throughput screening.

Protein misfolding and aggregation are associated with various neurodegenerative diseases. Cellular mechanisms that recognize and degrade misfolded ...

Utilizing pHluorin-tagged Receptors to Monitor Subcellular Localization and Trafficking

Understanding membrane protein trafficking, assembly, and expression requires an approach that differentiates between those residing in intracellular organelles and those localized on the plasma membrane. Traditional fluorescence-based measurements lack the capability to distinguish membrane proteins residing in different organelles. Cutting edge methodologies transcend traditional methods by coupling pH-sensitive fluorophores with total internal reflection fluorescence microscopy (TIRFM). TIRF illumination excites the sample up to approximately 150 nm from the glass-sample interface, thus decreasing background, increasing the signal to noise ratio, and enhancing resolution. The excitation volume in TIRFM encompasses the plasma membrane and nearby organelles such as the peripheral ER. Superecliptic pHluorin (SEP) is a pH sensitive version of GFP. Genetically encoding SEP into the extracellular domain of a membrane protein of interest positions the fluorophore on the luminal side of the ER and in the extracellular region of the cell. SEP is fluorescent when the pH is greater than 6, but remains in an off state at lower pH values. Therefore, receptors tagged with SEP fluoresce when residing in the endoplasmic reticulum (ER) or upon insertion in the plasma membrane (PM) but not when confined to a trafficking vesicle or other organelles such as the Golgi. The extracellular pH can be adjusted to dictate the fluorescence of receptors on the plasma membrane. The difference in fluorescence between TIRF images at neutral and acidic extracellular pH for the same cell corresponds to a relative number of receptors on the plasma membrane. This allows a simultaneous measurement of intracellular and plasma membrane resident receptors. Single vesicle insertion events can also be measured when the extracellular pH is neutral, corresponding to a low pH trafficking vesicle fusing with the plasma membrane and transitioning into a fluorescent state. This versatile technique can be exploited to study localization, expression, and trafficking of membrane proteins.

Labeling the extracellular domain of a membrane protein with a pH sensitive fluorophore, superecliptic pHluorin (SEP), allows subcellular localization, expression, and trafficking to be determined. Imaging SEP-labeled proteins with total internal reflection fluorescence microscopy (TIRFM) enables the quantification of protein levels in the peripheral ER and plasma membrane.

Understanding membrane protein trafficking, assembly, and expression requires an approach that differentiates between those residing in intracellular ...

Fluorescence Microplate-Based Cycloheximide Chase Assay: A Technique to Monitor the Degradation Kinetics of Fluorescent Nuclear Misfolded Proteins

Source: Guo, L. et. al., <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/54266">Assays for the Degradation of Misfolded Proteins in Cells.</a> J. Vis. Exp. (2016)
This video describes an in vitro fluorescence-based microplate assay to study the degradation kinetics of a fluorescently-labeled misfolded luciferase mutant protein expressed in the nuclei of transfected mammalian cells. The assay involves the treatment of cells expressing the fluorescent mutant protein with a translation inhibitor and a proteasome inhibitor to assess the proteasome-mediated degradation of the misfolded protein.

Source: Guo, L. et. al., Assays for the Degradation of Misfolded Proteins in Cells. J. Vis. Exp. (2016)
This video describes an in vitro ...

JoVE Encyclopedia of Experiments

Biological Techniques

Assay Techniques

Biochemical assays

Fluorescence Leakage Assay to Study Cell-Penetrating Peptide Interaction with Membrane

Source: Konate, K. et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/62028">Fluorescent Leakage Assay to Investigate Membrane Destabilization by Cell-Penetrating Peptide.</a> J. Vis. Exp. (2020)
This video discusses the fluorescence leakage assay to study the interactions of cell-penetrating peptides with cell membranes using cell membrane mimics &#8212; large unilamellar vesicles, or LUVs, encapsulated with a fluorescent dye and a quencher.

Source: Konate, K. et al. Fluorescent Leakage Assay to Investigate Membrane Destabilization by Cell-Penetrating Peptide. J. Vis. Exp. (2020)
This video ...

Cell-based assays

A Fluorogenic Peptide Cleavage Assay to Screen the Proteolytic Activity of Proteases

Source: Jaimes, J. A., et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/58892">A Fluorogenic Peptide Cleavage Assay to Screen for Proteolytic Activity: Applications for coronavirus spike protein activation.</a> J. Vis. Exp. (2019).
This video demonstrates an assay to screen for the proteolytic activity of proteases using fluorogenic peptides. The protease recognizes its cleavage site on the peptide, cleaving it and separating the quencher from the fluorophore, enabling its fluorescence emission. The fluorescence signal is detected and analyzed to check for the cleavage efficiency of different peptide variants.

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

Summary

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Chapters in this video

Construction of Cyclic Cell-Penetrating Peptides for Enhanced Penetration of Biological Barriers

Cell Membrane Repair Assay Using a Two-photon Laser Microscope

Measuring Peptide Translocation into Large Unilamellar Vesicles

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

Determining Membrane Protein Topology Using Fluorescence Protease Protection (FPP)

Assays for the Degradation of Misfolded Proteins in Cells

Utilizing pHluorin-tagged Receptors to Monitor Subcellular Localization and Trafficking

Fluorescence Microplate-Based Cycloheximide Chase Assay: A Technique to Monitor the Degradation Kinetics of Fluorescent Nuclear Misfolded Proteins

Fluorescence Leakage Assay to Study Cell-Penetrating Peptide Interaction with Membrane

A Fluorogenic Peptide Cleavage Assay to Screen the Proteolytic Activity of Proteases