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...

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.

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	<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&#160;</td>
		</tr>
		<tr >
			<td >Chloroform&#160;</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 &#181;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&#160;</td>
			<td >WAKO&#160;</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 &#181;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&#160;</td>
		</tr>
		<tr >
			<td >quartz fluorescence cuvette</td>
			<td >Hellma</td>
			<td >109.004F-QS</td>
			<td></td>
		</tr>
		<tr >
			<td >rotavapor system&#160;</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 (NaN3)</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&#160;</td>
			<td >Eromedex</td>
			<td >2000-B</td>
			<td></td>
		</tr>
		<tr >
			<td >Zetaziser NanoZS&#160;</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...

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

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

Single-molecule Super-resolution Imaging of Phosphatidylinositol 4,5-bisphosphate in the Plasma Membrane with Novel Fluorescent Probes

Phosphoinositides in the cell membrane are signaling lipids with multiple cellular functions. Phosphatidylinositol 4,5-bisphosphate (PI(4,5)P2) is a determinant phosphoinositide of the plasma membrane (PM), and it is required to modulate ion channels, actin dynamics, exocytosis, endocytosis, intracellular signaling, and many other cellular processes. However, the spatial organization of PI(4,5)P2 in the PM is controversial, and its nanoscale distribution is poorly understood due to the technical limitations of research approaches. Here by utilizing single molecule localization microscopy and the Pleckstrin Homology (PH) domain based dual color fluorescent probes, we describe a novel method to visualize the nanoscale distribution of PI(4,5)P2 in the PM in fixed membrane sheets as well as live cells.

PI(4,5)P2 regulates various cellular functions, but its nanoscale organization in the cell plasma membrane is poorly understood. By labeling PI(4,5)P2 with a dual-color fluorescent probe fused to the Pleckstrin Homology domain, we describe a novel approach to study the PI(4,5)P2 spatial distribution in the plasma membrane at the nanometer scale.

Phosphoinositides in the cell membrane are signaling lipids with multiple cellular functions. Phosphatidylinositol 4,5-bisphosphate (PI(4,5)P2) is a ...

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating these studies, reincorporation of detergent-solubilized membrane proteins into liposomes is a stochastic process where protein topology is impossible to enforce. This paper offers an alternative method to these challenging techniques that utilizes a liposome-based scaffold. Protein solubility is enhanced by deletion of the transmembrane domain, and these amino acids are replaced with a tethering moiety, such as a His-tag. This tether interacts with an anchoring group (Ni2+ coordinated by nitrilotriacetic acid (NTA(Ni2+)) for His-tagged proteins), which enforces a uniform protein topology at the surface of the liposome. An example is presented wherein the interaction between Dynamin-related protein 1 (Drp1) with an integral membrane protein, Mitochondrial Fission Factor (Mff), was investigated using this scaffold liposome method. In this work, we have demonstrated the ability of Mff to efficiently recruit soluble Drp1 to the surface of liposomes, which stimulated its GTPase activity. Moreover, Drp1 was able to tubulate the Mff-decorated lipid template in the presence of specific lipids. This example demonstrates the effectiveness of scaffold liposomes using structural and functional assays and highlights the role of Mff in regulating Drp1 activity.

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

This paper describes a method for assessing the interactions and assemblies of integral membrane proteins in vitro with various partner factors in a lipid-proximal environment.

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating ...

Method to Visualize and Analyze Membrane Interacting Proteins by Transmission Electron Microscopy

Monotopic proteins exert their function when attached to a membrane surface, and such interactions depend on the specific lipid composition and on the availability of enough area to perform the function. Nanodiscs are used to provide a membrane surface of controlled size and lipid content. In the absence of bound extrinsic proteins, sodium phosphotungstate-stained nanodiscs appear as stacks of coins when viewed from the side by transmission electron microscopy (TEM). This protocol is therefore designed to intentionally promote stacking; consequently, the prevention of stacking can be interpreted as the binding of the membrane-binding protein to the nanodisc. In a further step, the TEM images of the protein-nanodisc complexes can be processed with standard single-particle methods to yield low-resolution structures as a basis for higher resolution cryoEM work. Furthermore, the nanodiscs provide samples suitable for either TEM or non-denaturing gel electrophoresis. To illustrate the method, Ca2+-induced binding of 5-lipoxygenase on nanodiscs is presented.

Many proteins perform their function when attached to membrane surfaces. The binding of extrinsic proteins on nanodisc membranes can be indirectly imaged by transmission electron microscopy. We show that the characteristic stacking (rouleau) of nanodiscs induced by the negative stain sodium phosphotungstate is prevented by the binding of extrinsic protein.

Monotopic proteins exert their function when attached to a membrane surface, and such interactions depend on the specific lipid composition and on the ...

Easy Manipulation of Architectures in Protein-based Hydrogels for Cell Culture Applications

Hydrogels are recognized as promising materials for cell culture applications due to their ability to provide highly hydrated cell environments. The field of 3D templates is rising due to the potential resemblance of those materials to the natural extracellular matrix. Protein-based hydrogels are particularly promising because they can easily be functionalized and can achieve defined structures with adjustable physicochemical properties. However, the production of macroporous 3D templates for cell culture applications using natural materials is often limited by their weaker mechanical properties compared to those of synthetic materials. Here, different methods were evaluated to produce macroporous bovine serum albumin (BSA)-based hydrogel systems, with adjustable pore sizes in the range of 10 to 70 &#181;m in radius. Furthermore, a method to generate channels in this protein-based material that are several hundred microns long was established. The different methods to produce pores, as well as the influence of pore size on material properties such as swelling ratio, pH, temperature stability, and enzymatic degradation behavior, were analyzed. Pore sizes were investigated in the native, swollen state of the hydrogels using confocal laser scanning microscopy. The feasibility for cell culture applications was evaluated using a cell-adhesive RGD peptide modification of the protein system and two model cell lines: human breast cancer cells (A549) and adenocarcinomic human alveolar basal epithelial cells (MCF7).

Different methods to manipulate three-dimensional architecture in protein-based hydrogels are evaluated here with respect to material properties. The macroporous networks are functionalized with a cell-adhesive peptide, and their feasibility in cell culture is evaluated using two different model cell lines.

Hydrogels are recognized as promising materials for cell culture applications due to their ability to provide highly hydrated cell environments. The field ...

DNA Polymerase Activity Assay Using Near-infrared Fluorescent Labeled DNA Visualized by Acrylamide Gel Electrophoresis

For any enzyme, robust, quantitative methods are required for characterization of both native and engineered enzymes. For DNA polymerases, DNA synthesis can be characterized using an in vitro DNA synthesis assay followed by polyacrylamide gel electrophoresis. The goal of this assay is to quantify synthesis of both natural DNA and modified DNA (M-DNA). These approaches are particularly useful for resolving oligonucleotides with single nucleotide resolution, enabling observation of individual steps during enzymatic oligonucleotide synthesis. These methods have been applied to the evaluation of an array of biochemical and biophysical properties such as the measurement of steady-state rate constants of individual steps of DNA synthesis, the error rate of DNA synthesis, and DNA binding affinity. By using modified components including, but not limited to, modified nucleoside triphosphates (NTP), M-DNA, and/or mutant DNA polymerases, the relative utility of substrate-DNA polymerase pairs can be effectively evaluated. Here, we detail the assay itself, including the changes that must be made to accommodate nontraditional primer DNA labeling strategies such as near-infrared fluorescently labeled DNA. Additionally, we have detailed crucial technical steps for acrylamide gel pouring and running, which can often be technically challenging.

This protocol describes the characterization of DNA polymerase synthesis of modified DNA through observation of changes to near-infrared fluorescently labeled DNA using gel electrophoresis and gel imaging. Acrylamide gels are used for high resolution imaging of the separation of short nucleic acids, which migrate at different rates depending on size.

For any enzyme, robust, quantitative methods are required for characterization of both native and engineered enzymes. For DNA polymerases, DNA synthesis ...

Biotinylated Cell-penetrating Peptides to Study Intracellular Protein-protein Interactions

Here we present a protocol to study intracellular protein-protein interactions that is based on the widely used biotin-avidin pull-down system. The modification presented includes the combination of this technique with cell-penetrating sequences. We propose to design cell-penetrating baits that can be incubated with living cells instead of cell lysates and therefore the interactions found will reflect those that occur within the intracellular context. Connexin43 (Cx43), a protein that forms gap junction channels and hemichannels is down-regulated in high-grade gliomas. The Cx43 region comprising amino acids 266-283 is responsible for the inhibition of the oncogenic activity of c-Src in glioma cells. Here we use TAT as the cell-penetrating sequence, biotin as the pull-down tag and the region of Cx43 comprised between amino acids 266-283 as the target to find intracellular interactions in the hard-to-transfect human glioma stem cells. One of the limitations of the proposed method is that the molecule used as bait could fail to fold properly and, consequently, the interactions found could not be associated with the effect. However, this method can be especially interesting for the interactions involved in signal transduction pathways because they are usually carried out by intrinsically disordered regions and, therefore, they do not require an ordered folding. In addition, one of the advantages of the proposed method is that the relevance of each residue on the interaction can be easily studied. This is a modular system; therefore, other cell-penetrating sequences, other tags, and other intracellular targets can be employed. Finally, the scope of this protocol is far beyond protein-protein interaction because this system can be applied to other bioactive cargoes such as RNA sequences, nanoparticles, viruses or any molecule that can be transduced with cell-penetrating sequences and fused to pull-down tags to study their intracellular mechanism of action.

This is a protocol to study intracellular protein-protein interactions based on the biotin-avidin pull-down system with the novelty of combining cell-penetrating sequences. The main advantage is that the target sequence is incubated with living cells instead of cell lysates and therefore the interactions will occur within the cellular context.

Here we present a protocol to study intracellular protein-protein interactions that is based on the widely used biotin-avidin pull-down system. The ...

Measuring Trans-Plasma Membrane Electron Transport by C2C12 Myotubes

Trans-plasma membrane electron transport (tPMET) plays a role in protection of cells from intracellular reductive stress as well as protection from damage by extracellular oxidants. This process of transporting electrons from intracellular reductants to extracellular oxidants is not well defined. Here we present spectrophotometric assays by C2C12 myotubes to monitor tPMET utilizing the extracellular electron acceptors: water-soluble tetrazolium salt-1 (WST-1) and 2,6-dichlorophenolindophenol (DPIP or DCIP). Through reduction of these electron acceptors, we are able to monitor this process in a real-time analysis. With the addition of enzymes such as ascorbate oxidase (AO) and superoxide dismutase (SOD) to the assays, we can determine which portion of tPMET is due to ascorbate export or superoxide production, respectively. While WST-1 was shown to produce stable results with low background, DPIP was able to be re-oxidized after the addition of AO and SOD, which was demonstrated with spectrophotometric analysis. This method demonstrates a real-time, multi-well, quick spectrophotometric assay with advantages over other methods used to monitor tPMET, such as ferricyanide (FeCN) and ferricytochrome c reduction.

The goal of this protocol is to spectrophotometrically monitor trans-plasma membrane electron transport utilizing extracellular electron acceptors and to analyze enzymatic interactions that may occur with these extracellular electron acceptors.

Trans-plasma membrane electron transport (tPMET) plays a role in protection of cells from intracellular reductive stress as well as protection from damage ...

A Fluorescence Fluctuation Spectroscopy Assay of Protein-Protein Interactions at Cell-Cell Contacts

A variety of biological processes involves cell-cell interactions, typically mediated by proteins that interact at the interface between neighboring cells. Of interest, only few assays are capable of specifically probing such interactions directly in living cells. Here, we present an assay to measure the binding of proteins expressed at the surfaces of neighboring cells, at cell-cell contacts. This assay consists of two steps: mixing of cells expressing the proteins of interest fused to different fluorescent proteins, followed by fluorescence fluctuation spectroscopy measurements at cell-cell contacts using a confocal laser scanning microscope. We demonstrate the feasibility of this assay in a biologically relevant context by measuring the interactions of the amyloid precursor-like protein 1 (APLP1) across cell-cell junctions. We provide detailed protocols on the data acquisition using fluorescence-based techniques (scanning fluorescence cross-correlation spectroscopy, cross-correlation number and brightness analysis) and the required instrument calibrations. Further, we discuss critical steps in the data analysis and how to identify and correct external, spurious signal variations, such as those due to photobleaching or cell movement.
In general, the presented assay is applicable to any homo- or heterotypic protein-protein interaction at cell-cell contacts, between cells of the same or different types and can be implemented on a commercial confocal laser scanning microscope. An important requirement is the stability of the system, which needs to be sufficient to probe diffusive dynamics of the proteins of interest over several minutes.

This protocol describes a fluorescence fluctuation spectroscopy-based approach to investigate interactions among proteins mediating cell-cell interactions, i.e. proteins localized in cell junctions, directly in living cells. We provide detailed guidelines on instrument calibration, data acquisition and analysis, including corrections to possible artefact sources.

A variety of biological processes involves cell-cell interactions, typically mediated by proteins that interact at the interface between neighboring ...

A Fluorogenic Peptide Cleavage Assay to Screen for Proteolytic Activity: Applications for coronavirus spike protein activation

Enveloped viruses such as coronaviruses or influenza virus require proteolytic cleavage of their fusion protein to be able to infect the host cell. Often viruses exhibit cell and tissue tropism and are adapted to specific cell or tissue proteases. Moreover, these viruses can introduce mutations or insertions into their genome during replication that may affect the cleavage, and thus can contribute to adaptations to a new host. Here, we present a fluorogenic peptide cleavage assay that allows a rapid screening of peptides mimicking the cleavage site of viral fusion proteins. The technique is very flexible and can be used to investigate the proteolytic activity of a single protease on many different substrates, and in addition, it also allows exploration of the activity of multiple proteases on one or more peptide substrates. In this study, we used peptides mimicking the cleavage site motifs of the coronavirus spike protein. We tested human and camel derived Middle East Respiratory Syndrome coronaviruses (MERS-CoV) to demonstrate that single and double substitutions in the cleavage site can alter the activity of furin and dramatically change cleavage efficiency. We also used this method in combination with bioinformatics to test furin cleavage activity of feline coronavirus spike proteins from different serotypes and strains. This peptide-based method is less labor- and time intensive than conventional methods used for the analysis of proteolytic activity for viruses, and results can be obtained within a single day.

We present a fluorogenic peptide cleavage assay that allows a rapid screening of the proteolytic activity of proteases on peptides representing the cleavage site of viral fusion peptides. This method can also be used on any other amino acid motif within a protein sequence to test for the protease activity.

Enveloped viruses such as coronaviruses or influenza virus require proteolytic cleavage of their fusion protein to be able to infect the host cell. Often ...

Single Liposome Measurements for the Study of Proton-Pumping Membrane Enzymes Using Electrochemistry and Fluorescent Microscopy

Proton-pumping enzymes of electron transfer chains couple redox reactions to proton translocation across the membrane, creating a proton-motive force used for ATP production. The amphiphilic nature of membrane proteins requires particular attention to their handling, and reconstitution into the natural lipid environment is indispensable when studying membrane transport processes like proton translocation. Here, we detail a method that has been used for the investigation of the proton-pumping mechanism of membrane redox enzymes, taking cytochrome bo3 from Escherichia coli as an example. A combination of electrochemistry and fluorescence microscopy is used to control the redox state of the quinone pool and monitor pH changes in the lumen. Due to the spatial resolution of fluorescent microscopy, hundreds of liposomes can be measured simultaneously while the enzyme content can be scaled down to a single enzyme or transporter per liposome. The respective single enzyme analysis can reveal patterns in the enzyme functional dynamics that might be otherwise hidden by the behavior of the whole population. We include a description of a script for automated image analysis.

Here, we present a protocol to study the molecular mechanism of proton translocation across lipid membranes of single liposomes, using cytochrome bo3 as an example. Combining electrochemistry and fluorescence microscopy, pH changes in the lumen of single vesicles, containing single or multiple enzyme, can be detected and analyzed individually.

Proton-pumping enzymes of electron transfer chains couple redox reactions to proton translocation across the membrane, creating a proton-motive force used ...