The authors were supported by a Russian Science Foundation grant 20-74-00133.

1. Fluorescent protein labeling
<ol>
	<li>Material preparation
	<ol>
		<li>Prepare 1 M Sodium bicarbonate buffer, pH 9.0, store it at 4 &#176;C, and use it within one week.</li>
		<li>Prepare 1.5 M hydroxylamine hydrochloride buffer, pH 8.5, immediately before use.</li>
		<li>Prepare a 10 mg/mL solution of fluorescent dye (see the Table of Materials) in dimethylsulfoxide. 
		NOTE: This solution can be stored for a month at -20 &#176;C in the dark.</li>
		<li>Prepare solutions of purified antibodies or other proteins at 1-10 mg/mL. 
		NOTE: Avoid buffers containing ammonium ions or primary amines. Replace the buffers containing Tris or glycerol with phosphate-buffered saline (PBS) by dialysis. Neither sodium azide (&#8804;3 mM) nor thimerosal (&#8804;1 mM) will significantly affect the conjugation reaction.</li>
		<li>Incubate the gel filtration medium (see the Table of Materials) for protein purification in PBS overnight at room temperature or for 2 h at 60 &#176;C. Apply the gel filtration medium onto spin columns with 0.2 &#181;m membranes.</li>
	</ol>
	</li>
	<li>Calculate the amount of reactive dye to be used for each reaction according to the concentration of protein to be labeled by using Eq (1). 
	<img alt="Equation 1" src="/files/ftp_upload/63459/63459eq1v4.jpg" /> 
	Where Vdye is the volume of the dye stock solution; Cpr, Vpr and MWpr is the concentration, volume and molar weight of the protein; MWdye is the dye molar weight; 100 is a unit conversion factor; MR is the molar ratio of dye to protein in the reaction mixture. 
	NOTE: The following MRs are recommended for IgG labeling reactions: MR = 40 if the antibody is at 1-3 mg/mL or MR = 30 if the antibody is at 4-10 mg/mL. For coagulation factors, MR = 5 is usually used.</li>
	<li>Conjugation reaction
	<ol>
		<li>In a reaction tube, mix the protein solution with a 10x lower volume of 1 M bicarbonate solution.</li>
		<li>Add the required amount of fluorescent dye (see step 1.2) with continuous stirring.</li>
		<li>Incubate the reaction mixture at room temperature for approximately 1 h, protected from light and with continuous stirring.</li>
		<li>For each 200 &#181;L of protein solution, add 5 &#181;L of 1.5 M hydroxylamine hydrochloride.</li>
		<li>Incubate the reaction mixture at room temperature for approximately 30 min, protected from light and with continuous stirring.</li>
	</ol>
	</li>
	<li>Prepare the spin column.
	<ol>
		<li>Add 500 &#181;L of gel filtration medium for protein purification to a column. Centrifuge the column for 3 min at 1,000 &#215; g.</li>
		<li>Discard the buffer from the collection tube. If the column is not full, add more gel filtration medium, and centrifuge the column for 3 min at 1,000 &#215; g. Repeat this step until the column is full.</li>
	</ol>
	</li>
	<li>Purification
	<ol>
		<li>Centrifuge the reaction mixture (from step 1.3.5) for 5 min at 17,000 &#215; g and remove the precipitate.</li>
		<li>Transfer the supernatant to the spin column with gel filtration medium. Allow the solution to absorb into the gel bed.</li>
		<li>Use an empty collection tube for the spin column and centrifuge it for 5 min at 1,000 &#215; g. After centrifugation, collect the labeled protein from the collection tube.</li>
	</ol>
	</li>
	<li>Determination of the degree of labeling
	<ol>
		<li>Correct for the contribution of the dye to the absorbance at A280 by measuring the absorbance of free dye at 280 nm (A280) and the &#955;max for the dye (Amax) (see Eq (2)). 
		<img alt="Equation 2" src="/files/ftp_upload/63459/63459eq02.jpg" />&#160; &#160; (2)</li>
		<li>Measure the absorbance of the protein-dye conjugate at 280 nm (A280) and the &#955;max for the dye (Amax) and calculate the protein concetration using Eq (3). 
		<img alt="Equation 3" src="/files/ftp_upload/63459/63459eq3v2.jpg" />&#160; &#160; (3) 
		Where &#949;M is the molar extinction coefficient of the protein at 280 nm; d is the optical path length during the measurement of the absorbance; CF is the contribution of the dye to the absorbance at A280 (step 1.6.1).</li>
		<li>Calculate the degree of labeling (DOL) using Eq (4). 
		<img alt="Equation 4" src="/files/ftp_upload/63459/63459eq4v2.jpg" />&#160; &#160; (4) 
		Where &#949;M dye is the molar extinction coefficient of the dye at &#955;max nm; d is the optical path length during the measurement of the absorbance; Cprotein is the concentration of the protein (step 1.6.2).</li>
	</ol>
	</li>
</ol>
2. Preparation of phospholipid vesicles
<ol>
	<li>Preparation and storage of the lipid mixture
	<ol>
		<li>Combine the lipids in the appropriate ratio (phosphatidylserine/phosphatidylcholine&#160;at a ratio of 20 mol% to 80 mol%).</li>
		<li>Dry the lipid mixture after lyophilization or evaporation and store it under an inert atmosphere in glass ampules.</li>
	</ol>
	</li>
	<li>Lipid film production
	<ol>
		<li>Open the ampule, and resuspend the lipid mixture in a small amount (~100 &#181;L) of chloroform. 
		NOTE: Do not use too much chloroform as it does not evaporate completely.</li>
		<li>Add DiIC16(3) in ethanol at 0.2 mol%. Transfer the lipid mixture to a round bottom flask. Spread the mixture thinly over the sides of the flask by rotating it. Dry the lipid mix for 30 min under an argon stream.</li>
	</ol>
	</li>
	<li>Hydration of the lipid mixture
	<ol>
		<li>Add a suitably warm (~55 &#176;C) aqueous buffer (HEPES 20 mM, NaCl 140 mM, pH 7.4) in a volume corresponding to the expected lipid concentration to the flask with the lipid film. Incubate the mixture with vortexing at 55 &#176;&#1057; for 30 min for complete hydration.</li>
		<li>Placing the sample tube in a freezer or warm thermostat to make the lipid suspension undergo 3-5 freeze-thaw cycles.</li>
	</ol>
	</li>
	<li>Formation of lipid vesicles by extrusion
	<ol>
		<li>Prepare the extruder according to the manufacturer&#39;s instructions. Warm up all the extruder components to the phase transition temperature of the lipid mixture.</li>
		<li>Fill one of the extruder syringes with the hydrated lipid mixture. Wait for 5-10 min for the temperature of the lipid suspension to equilibrate with the temperature of the extruder.</li>
		<li>Extrude the lipid mixture through the membrane at least 10 times. For the final extrusion, place the lipid suspension in the alternate syringe and look for a change in appearance from a slightly nebulous to a clear solution.</li>
		<li>Store the resulting mixture of lipid vesicles at 4 &#176;C, preferably in an inert atmosphere of argon or nitrogen, for 3-4 days. Do not freeze.</li>
	</ol>
	</li>
</ol>
3. Isolation of platelets from whole blood
<ol>
	<li>Collect whole blood from healthy donors in tubes containing 3.2% sodium citrate.</li>
	<li>Add prostaglandin E1 (PGE1) (1 &#181;M) and apyrase (0.1 U/mL) to the blood, followed by centrifugation at room temperature at 100 &#215; g for 8 min.</li>
	<li>After centrifugation, take the platelet-rich plasma and add sodium citrate solution (106 mM, pH 5.5) to a plasma/citrate ratio of 3:1. Centrifuge the plasma at room temperature at 400 &#215; g for 5 min.</li>
	<li>Remove the supernatant, and resuspend the pellet in 300 &#181;L of Tyrode&#39;s buffer without BSA (20 mM HEPES, 150 mM NaCl, 2.7 mM KCl, 1 mM MgCl2, 0.4 mM NaH2PO4, 2.5 mM CaCl2, 5 mM glucose, pH 7.4). Purify the platelets from plasma proteins by gel chromatography on the gel filtration medium for platelet purification (see the Table of Materials).</li>
</ol>
4. Detection of protein - lipid interaction by flow cytometry
<ol>
	<li>Kinetic binding experiments
	<ol>
		<li>Dilute phospholipid vesicles (from step 2.4.4) in Tyrode&#39;s buffer (20 mM HEPES, 150 mM NaCl, 2.7 mM KCl, 1 mM MgCl2, 0.4 mM NaH2PO4, 2.5 mM CaCl2, 5 mM glucose, 0.5% BSA, pH 7.4) to a concentration of 1 &#181;M and total volume of 250 &#181;L.</li>
		<li>Mix fluorescent-labeled coagulation factor X (fX-fd) from step 1 at a concentration of 500 nM with the phospholipid vesicles from step 4.1.1 in a 1:1 ratio (final vesicle concentration 0.5 &#181;M, fX-fd concentration is 250 nM of fX) to a total volume of 500 &#181;L.</li>
		<li>Immediately inject the 500 &#181;L of the mixed suspension (~20 min for analysis with a low flowing rate) into the flow cytometer. Use a low flow rate and ensure that the threshold for channel FL2 (ex&#1089;itation 488 nm, emission filter 585/42 nm) is as value 200. Measure the mean fluorescence in channel FL4 (ex&#1089;itation 633 nm, emission filter 660/20) for the fluorescence dye from the Table of Materials. 
		NOTE: Choose a cytometer without an autosampler. This will speed up the process of injection of the sample into the measuring cell.</li>
		<li>When saturation of binding is achieved (no significant increase in fluorescence within 5 min), rapidly dilute the sample 20-fold with Tyrode&#39;s buffer, and monitor the dissociation until baseline fluorescence is reached (complete dissociation) or until a plateau is reached (no significant decrease in fluorescence within 5 min). 
		NOTE: As a control, add 10 &#181;M EDTA and monitor complete dissociation for 5 min.</li>
	</ol>
	</li>
	<li>Equilibrium binding experiments 
	NOTE: Use the kinetic curve of binding to determine the time to reach saturation; the time for saturation for fX-fd and artificial vesicles is 20 min.&#160;
	<ol>
		<li>Incubate artificial phospholipid vesicles (5 &#181;M) for the binding assay with different concentrations of fX-fd (from 0 to 1,000 nM) in Tyrode&#39;s buffer for 20 min.</li>
		<li>Dilute each sample from step 4.2.1&#160;by 20x to a final volume of 200 &#181;L with Tyrode&#39;s buffer. Immediately analyze the diluted sample by flow cytometry within 30 s. Use settings from step 4.1.3. 
		NOTE: As a control for nonspecific binding, use similar samples with EDTA (10 &#181;M) and incubate them for 5 min.</li>
	</ol>
	</li>
</ol>
5. Analysis of flow cytometry data
<ol>
	<li>Export experiments in FSC format from cytometry data acquisition software to cytometer software for data analysis (see the Table of Materials). Choose File | Export | FCS files. Open FSC files in cytometer software for data analysis by selecting the files on the computer and dragging them to the workspace of the program.</li>
	<li>For gating of the microvesicles, identify the region of microvesicles by the fluorescence of the lipophilic dye DiIC16(3). Use menu commands or plot button in the worksheet to create dot plot SSC from FL2 (dye DilC16(3)) in log coordinates. Choose the Rectangular Gate button to draw a gating region so that events from a sample without vesicles are not included in this region (Figure 1B,C).</li>
	<li>Analyze the kinetic experiments.
	<ol>
		<li>Create a dot-plot using the coordinates of fluorescence (FL4) over time for the region of the vesicles (double-click in the region of vesicles in step 5.2)</li>
		<li>Export the data on the change in fluorescence over time in csv format. Choose Sample | Right-click | Export | Choose FL4 and Time in Parameters | Select directory for saving | Select CSV format | Export.</li>
		<li>Open the CSV file in any statistical software (see the Table of Materials). Calculate a simple moving average of fluorescence and time for every 1,000 events.</li>
		<li>Approximate a graph of the dependence of the simple moving average fluorescence on time under the assumption of exponential dependence (Analysis &#62; Fitting &#62;Nonlinear Curve Fit) and use this to calculate the kinetic association constant using Eq (5). 
		<img alt="Equation 5" src="/files/ftp_upload/63459/63459eq05v2.jpg" />&#160; &#160; (5) 
		Where [XB] is the bound factor concentration at each moment of time (user-defined units) according to the simple moving average from step 5.3.3; [X] is the added factor concentration; [X]max is the maximum bound factor concentration; k is the association constant; t is the time.</li>
		<li>Repeat the same set of actions (5.3.1-5.3.4) to calculate the kinetic dissociation constant using Eq (6). 
		<img alt="Equation 6" src="/files/ftp_upload/63459/63459eq06v2.jpg" />&#160; &#160; (6) 
		Where [XB] is the bound factor concentration at each moment of time; [X]0 is the bound factor concentration at the initial moment of time; k is the dissociation constant; t is the time.</li>
	</ol>
	</li>
	<li>Equilibrium binding assay
	<ol>
		<li>Determine the average fluorescence of fX-fd in the region of the vesicles for each selected concentration of fX-fd.</li>
		<li>Approximate the dependence of the bound factor fluorescence from the concentration of the added factor in the assumption of simple single-site binding. Calculate the average binding parameters using Eq (7) from three independent repeats at a minimum. 
		<img alt="Equation 7" src="/files/ftp_upload/63459/63459eq07v2.jpg" />&#160; &#160; (7) 
		Where [XB] is the bound factor concentration; [X] is the added factor concentration; nx&#160;is the apparent number of binding sites per vesicle; Kd is the apparent dissociation constant.</li>
	</ol>
	</li>
</ol>
6. Converting fluorescence intensity to the mean number of binding sites
<ol>
	<li>Prepare calibrated beads.
	<ol>
		<li>Incubate gel-filtered platelets (see step 3.3) with A23187 (10 &#181;M) in the presence of CaCl2 (2.5 mM) for 10 min at room temperature.</li>
		<li>Add to the activated platelets the various concentrations of fX-fd (0 to 1,000 nM). Add 2% v/v formaldehyde and incubate for 1 h. Stop the reaction by incubating the platelets with 3 M glycine and 5% BSA for 30 min at room temperature.</li>
		<li>Purify the mixture from the unreacted dye. Centrifuge the platelets for 5 min at 400 &#215; g, remove the supernatant, and resuspend the pellet in Tyrode&#39;s buffer (containing 0.5% BSA). 
		NOTE: Repeat step 6.1.3 three times.</li>
	</ol>
	</li>
	<li>Measure the fluorescence level of the calibration beads in each sample first using a spectrofluorometer (for fluorescent dye from the Table of Materials, excitation 633 nm, emission 670 nm) and then using the flow cytometer (in channel FL4: excitation 633 nm, emission filter 660/20). Using a cell counter, determine the number of beads in each sample.</li>
	<li>Convert the fluorescence intensity of each respective bead sample to the concentration of soluble fluorescent dye using a spectrofluorometer. Recalculate the fluorescent dye concentration for the number of fluorophore molecules using Eq (8). 
	<img alt="Equation 8" src="/files/ftp_upload/63459/63459eq08.jpg" />&#160; &#160;(8) 
	Where Nx is the number of fluorophore molecules; C is the fluorescent dye concentration; NA&#160;is Avogadro constant; NA = 6.02214076&#215;1023 mol -1.</li>
	<li>Create a dependence graph of the average fluorescence of the beads in a flow cytometer (step 6.2) on the number of fluorophore molecules (see step 6.3) for each sample using any statistical software (see the Table of Materials). Approximate this dependence by line proportionality (Analysis | Fitting | Fit linear). From the approximation in Eq (9), calculate the conversion factor of the mean fluorescence to binding sites. 
	<img alt="Equation 9" src="/files/ftp_upload/63459/63459eq09.jpg" />&#160; &#160; (9) 
	Where MF is the mean fluorescence of beads by flow cytometry; Nx is the number of fluorophore molecules per bead; CF represents the conversion factor of the mean fluorescence to binding sites. CF and b are obtained from the results of fitting the graph by linear proportionality.</li>
	<li>Calculate the apparent number of binding sites per vesicle of interest using Eq (10). 
	<img alt="Equation 10" src="/files/ftp_upload/63459/63459eq10.jpg" />&#160; &#160; (10) 
	Where nx is the apparent number of binding sites per vesicle of interest; MF is the mean fluorescence of the vesicles of interest by flow cytometry; CF and b are conversion factors from the Eq (8).</li>
</ol>

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The authors have no conflicts of interest to disclose.

The proposed method can be adapted for a rough characterization of the interaction of proteins with phospholipid membranes from various sources and compositions. The quantitative flow cytometry described here concedes to surface plasmon resonance (SPR) in several parameters. In particular, it has a lower sensitivity and time resolution and requires fluorescent labeling of proteins. Fluorescent labeling can lead to a change in conformation and loss of activity for many proteins and therefore requires careful control. However, this technique has significant advantages over others. This method provides an opportunity to explore the interaction of proteins with the native cell membrane, which is not readily implemented using SPR. Moreover, the approach allows the estimation of the number of protein binding sites on the membrane surface and can be efficient for some analysis tasks. 
Commercially available beads are available for some fluorescent dyes to count binding sites. However, there are no such beads for many widely used dyes. Hence, self-made beads are the best way to resolve this. The same cells were used to prepare these beads as for all other experiments. However, as the beads require high centrifugation speeds during washing, phospholipid vesicles cannot be used. However, cells or vesicles can be replaced with beads with amino-reactive groups, which can be conjugated to the chosen dye. The sequence of actions will be similar to those described in step 6.1.
The limitations of this quantitative flow cytometry method are related to the technical capabilities of the used cytometer. Three different models of flow cytometers (with the variable lasers, detectors, pumps) were applied to reproduce this technique without any complication. However, the selection of a fluorescent label suitable for the cytometer used must be carefully considered because the set of lasers, detectors, and optical filters differ from device to device, even within the same model. It is necessary to focus on the ability of the cytometer to measure microparticles of a specific diameter; not all instruments are equally capable of detecting particles at resolutions below 200 nm (to determine this, use the fixed-size calibration beads supplied by the instrument's manufacturer). Additionally, some flow cytometers, which are sampled using a syringe pump, cannot measure continuous binding kinetics in principle. In this case, the kinetics can be recorded only point by point, taking separate samples for measurement at certain points in time4,6.
Flow cytometry is used to investigate the expression of antigens on various cells-the presence/absence of an antigen and the percentage of cell populations expressing and not expressing this antigen. The ability of flow cytometry to concurrently discriminate free and bound ligands without additional separation procedures also affords the opportunity for quantitative assessment of ligand-binding dynamics. The method proposed herein describes the preparation of self-made calibration beads to quantify the binding of a fluorescent ligand to artificial phospholipid vesicles. This approach does not limit the choice of commercially available fluorophores. Beyond that, the technique described here for the acquisition and analysis of kinetic data enables improved time resolution. Thus, the method described herein can be used alone as a first step in characterizing protein-membrane interactions or in combination with other methods (for example, SPR or microscopy) to improve measurement accuracy.

Biomembranes separate the inner contents of animal cells and extracellular space. Note that membranes also surround microvesicles formed during the cell&#39;s life cycle and organelles. The cell membrane is predominantly composed of lipids and proteins. Membrane proteins perform signaling, structural, transport, and adhesive functions. However, the lipid bilayer is also essential for the interrelation of the animal cell with the extracellular space. This paper proposes a method for studying the peripheral interaction of external proteins with the lipid membrane.
The most striking example of reactions occurring on the outer membrane layer of an animal cell is the blood coagulation reaction. An important feature of blood coagulation is that all the main reactions proceed on the phospholipid membranes of cells and microvesicles arising from these cells and not in the plasma1,2,3. Membrane-dependent reactions include the process of starting coagulation (on the cell membranes of the subendothelium, inflamed endothelium, or activated immune cells, with the participation of a tissue factor), all reactions of the main cascade-activation of factors IX, X, prothrombin; activation of factor XI by thrombin (on the membranes of activated platelets, erythrocytes, lipoproteins, and microvesicles); reactions of the protein C pathway; inactivation of coagulation enzymes (on the membranes of endothelial cells with the participation of thrombomodulin cofactors, endothelial protein C receptor, heparan sulfate); and contact pathway reactions (on membranes of platelets and some microvesicles with the participation of unknown cofactors). Thus, it is impossible to investigate blood coagulation without studying the interaction of various plasma proteins with the membrane of blood cells.
This paper describes a flow-cytometry-based method for characterizing the interaction of proteins with lipid membranes of cells or microvesicles. This approach was initially proposed to study the interaction of blood plasma with platelets and artificial phospholipid vesicles. Moreover, most of the studied proteins interact directly with negatively charged membrane phospholipids, particularly with phosphatidylserine4,5. Additionally, there are proteins whose interaction with the membrane is mediated by special receptors6.
An important ability of flow cytometry is discriminating between free and bound ligands without additional separation. This feature of cytometry allows the study of ligand equilibrium binding at the endpoint and helps perform continuous kinetic measurements. The technique is unsophisticated and does not require complex sample preparation. Flow cytometry is actively used to quantitatively study the dynamics of interaction between fluorescent peptides, receptors, and G-proteins in intact and detergent-permeable neutrophils7. This approach is also applicable for exploring protein-DNA interactions and the kinetics of endonuclease activity in real time8. Over time, this method was used to quantitatively study high-affinity protein-protein interactions with purified lipid vesicles9, or, more generally, with membrane proteins expressed in a highly efficient Sf9 cell expression system10. Quantitative methods have also been described for characterizing protein-liposome interactions using flow cytometry for transmembrane proteins11.
This technique uses self-made calibration beads to avoid using commercially available beads7. The calibration beads used previously7 were intended to work with fluorescein, which substantively restricted the assortment of accessible fluorescent ligands on the proteins. In addition, this paper offers a new way to acquire and analyze kinetic data for reasonable time resolution. Although this method is described for artificial phospholipid vesicles, there are no obvious limitations for its adaptability to cells, natural vesicles, or artificial phospholipid vesicles with a different lipid composition. The method described herein allows the estimation of the parameters of interaction (kon, koff) and equilibrium (Kd) and facilitates quantitative characterization of the number of protein binding sites on the membrane. Note that this technique provides an approximate estimate of the number of binding sites. The advantages of the method are its relative simplicity, accessibility, and adaptability to native cells and natural and artificial microvesicles.

<table><tbody><tr><td>A23187</td><td>Sigma Aldrich</td><td>C7522-10MG</td><td></td></tr><tr><td>Alexa Fluor 647 NHS Ester (Succinimidyl Ester)</td><td>Thermo Fisher Scientific</td><td>A37573</td><td>fluorescent dye</td></tr><tr><td>Apyrase from potatoes</td><td>Sigma Aldrich</td><td>A2230</td><td></td></tr><tr><td>BD FACSCantoII</td><td>BD Bioscience</td><td></td><td></td></tr><tr><td>bovine serum albumin</td><td>VWR Life Science AMRESCO</td><td>Am-O332-0.1</td><td></td></tr><tr><td>Calcium chloride, anhydrous, powder, &amp;ge;97%</td><td>Sigma Aldrich</td><td>C4901-100G</td><td></td></tr><tr><td>Cary Eclipse Fluorescence Spectrometer</td><td>Agilent</td><td></td><td></td></tr><tr><td>D-(+)-Glucose</td><td>Sigma Aldrich</td><td>G7528-1KG</td><td></td></tr><tr><td>DiIC16(3) (1,1&#039;-Dihexadecyl-3,3,3&#039;,3&#039;-Tetramethylindocarbocyanine Perchlorate)</td><td>Thermo Fisher Scientific</td><td>D384</td><td></td></tr><tr><td>DMSO</td><td>Sigma Aldrich</td><td>D8418</td><td></td></tr><tr><td>EDTA disodium salt</td><td>VWR Life Science AMRESCO</td><td>Am-O105B-0.1</td><td></td></tr><tr><td>FACSDiva</td><td>BD Bioscience</td><td></td><td>cytometry data acquisition software</td></tr><tr><td>FlowJo</td><td>Tree Star</td><td></td><td>cytometer software for data analysis</td></tr><tr><td>HEPES</td><td>Sigma Aldrich</td><td>H4034-500G</td><td></td></tr><tr><td>Human Factor X</td><td>Enzyme research</td><td>HFX 1010</td><td></td></tr><tr><td>Hydroxylamine hydrochloride</td><td>Panreac</td><td>141914.1209</td><td></td></tr><tr><td>L-&amp;alpha;-phosphatidylcholine (Brain, Porcine)</td><td>Avanti Polar Lipids</td><td>840053P</td><td></td></tr><tr><td>L-&amp;alpha;-phosphatidylserine (Brain, Porcine) (sodium salt)</td><td>Avanti Polar Lipids</td><td>840032P</td><td></td></tr><tr><td>Magnesium chloride</td><td>Sigma Aldrich</td><td>M8266-100G</td><td></td></tr><tr><td>Mini-Extruder</td><td>Avanti Polar Lipids</td><td>610020-1EA</td><td></td></tr><tr><td>OriginPro 8 SR4 v8.0951</td><td>OriginLab Corporation</td><td></td><td>Statistical software</td></tr><tr><td>Phosphate Buffered Saline (PBS) Tablets, Biotechnology Grade</td><td>VWR Life Science AMRESCO</td><td>97062-732</td><td></td></tr><tr><td>Potassium chloride</td><td>Sigma Aldrich</td><td>P9541-500G</td><td></td></tr><tr><td>Prostaglandin E1</td><td>Cayman Chemical</td><td>13010</td><td></td></tr><tr><td>Sephadex G25</td><td>GE Healthcare</td><td>GE17-0033-01</td><td>gel filtration medium for protein purification</td></tr><tr><td>Sepharose CL-2B</td><td>Sigma Aldrich</td><td>CL2B300-500ML</td><td>gel filtration medium for platelet purification</td></tr><tr><td>Sodium bicarbonate</td><td>Corning</td><td>61-065-RO</td><td></td></tr><tr><td>Sodium chloride</td><td>Sigma Aldrich</td><td>S3014-500G</td><td></td></tr><tr><td>Sodium phosphate monobasic</td><td>Sigma Aldrich</td><td>S3139-250G</td><td></td></tr><tr><td>Spin collumns with membrane 0.2 &amp;micro;m</td><td>Sartorius</td><td>VS0171</td><td></td></tr><tr><td>Trisodium citrate dihydrate</td><td>Sigma Aldrich</td><td>S1804-1KG</td><td></td></tr></tbody></table>

analyzing the interaction of fluorescent-labeled proteins with artificial phospholipid microvesicles using quantitative flow cytometry

In the human body, most of the major physiologic reactions involved in the immune response and blood coagulation proceed on the membranes of cells. An important first step in any membrane-dependent reaction is binding of protein on the phospholipid membrane. An approach to studying protein interaction with lipid membranes has been developed using fluorescently labeled proteins and flow cytometry. This method allows the study of protein-membrane interactions using live cells and natural or artificial phospholipid vesicles. The advantage of this method is the simplicity and availability of reagents and equipment. In this method, proteins are labeled using fluorescent dyes. However, both self-made and commercially available, fluorescently labeled proteins can be used. After conjugation with a fluorescent dye, the proteins are incubated with a source of the phospholipid membrane (microvesicles or cells), and the samples are analyzed by flow cytometry. The obtained data can be used to calculate the kinetic constants and equilibrium Kd. In addition, it is possible to estimate the approximate number of protein binding sites on the phospholipid membrane using special calibration beads.

The flow cytometry method described herein is used to characterize the binding of plasma coagulation proteins to activated platelets. In addition, phospholipid vesicles PS:PC 20:80 were applied as a model system. This paper mainly focuses on artificial phospholipid vesicles as an example. The parameters of the cytometer, in particular, the photomultiplier tube (PMT) voltage and the compensation must be selected for each specific device, the object of study (cells, artificial or natural microvesicles), and the dyes used. Figure 1B,C show an example of gating artificial phospholipid vesicles that are ~1 &#181;m in size with the incorporated lipophilic fluorescent dye DiIC16 (3). Large vesicle size and lipophilic fluorescent dye helped detect vesicles using a cytometer. The gate was set based on a sample containing the same-size artificial lipid vesicles but without the fluorescent dye (Figure 1B). Only events inside this gate were used in the analysis.
The kinetics of protein binding to vesicles was analyzed at the first stage. The sample for this was collected continuously as described in step 4.1. A typical dot-plot is shown in Figure 1D-F. The data obtained were analyzed using the flow cytometry software. The resulting curve is shown in Figure 1G. Solid lines show the curves of approximation, from which the kinetic constants of association (kon) and dissociation (koff) were obtained. As factor X binds to phospholipid vesicles reversibly and Ca2+-dependently, samples with EDTA controlled the specificity and reversibility of this binding. The resulting constants are shown in Table 1.
Based on the kinetics of binding, a time of 20 min was chosen for further equilibrium experiments to describe saturation in binding accurately. The mean fluorescence intensity of the factor was subsequently determined in the region of the vesicles. Each sample was analyzed in the presence and absence of EDTA. The fluorescence intensity in the presence of EDTA was taken as background and subtracted from the total signal as the binding of fX to the membrane in the absence of Ca2+ ions is considered nonspecific. The resulting fluorescence was converted to the number of binding sites per vesicle using the calibration beads.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63459/63459fig1v2.jpg" /> 
Figure 1: Specific binding of fX to artificial phospholipid vesicles. (A) Scheme of experiment. (B, C) Typical dot plots of phospholipid vesicles without (B) or with (C) lipophilic fluorescent dye DiIC16 (3). (D-F) Typical dot plots of factor X interaction with phospholipid vesicles before (D) or after (E) 20-fold dilution and in presence EDTA (F). (G) Kinetics of FX (250 nM) binding and dissociation to phospholipid vesicles. (H) Equilibrium interaction of factor X to phospholipid vesicles. Results are the means &#177;SD for n=3 different samples. Abbreviations: FX = Factor X; Ph vesicles = phospholipid vesicles; SSC = side scatter; a.u. = arbitrary unit). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63459/63459fig1v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Apparent Kd &#177; SEM (nM)</td>
			<td>kon &#177; SEM (&#956;M-1s-1)</td>
			<td>Koff &#177; SEM (s-1)</td>
			<td>Apparent number of binding sites per vesicle &#177; SEM</td>
		</tr>
		<tr>
			<td>400 &#177; 80</td>
			<td>0.371 &#177; 0.012</td>
			<td>0.019 &#177; 0.004</td>
			<td>8,000 &#177; 800</td>
		</tr>
	</tbody>
</table>
Table 1: Parameters of fX interaction with artificial phospholipid vesicles. Parameters were determined from the curves (see Figure 1F,G). Results are the mean &#177; SEM for n = 3.

Watch this Scientific Journal Video about Analyzing the Interaction of Fluorescent-Labeled Proteins with Artificial Phospholipid Microvesicles using Quantitative Flow Cytometry at JoVE.com

Analyzing the Interaction of Fluorescent-Labeled Proteins with Artificial Phospholipid Microvesicles using Quantitative Flow Cytometry

Here, we describe a set of methods for characterizing the interaction of proteins with membranes of cells or microvesicles.

In the human body, most of the major physiologic reactions involved in the immune response and blood coagulation proceed on the membranes of cells. An ...

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Nanyang Technological University

Research

JoVE Journal

Biochemistry

2.3K Views.  Russian Academy of Sciences. Met behulp van deze hoeveelheidsstroomcytometriemethode kunnen onderzoekers de kinetische en evenwichtsgebruiken van ingebrachte membraaninteractie berekenen en het aantal eiwitten schatten dat wordt binnengebracht door locaties op het PPH-membraan. De voordelen van deze methode zijn de eenvoud en beschikbaarheid van reagentia en apparatuur. Deze methode is uitsluitend bedoeld voor fundamenteel onderzoek.Het protocol is ontwikkeld om eiwitlipide-interacties in bloedberekening te bestud...

Video: Analyzing the Interaction of Fluorescent-Labeled Proteins with Artificial Phospholipid Microvesicles using Quantitative Flow Cytometry

Fluorescence-Based Measurements of Phosphatidylserine/Phosphatidylinositol 4-Phosphate Exchange Between Membranes

Several members of the evolutionarily conserved oxysterol-binding protein (OSBP)-related proteins(ORP)/OSBP homologs (Osh) family have recently been found to represent a novel lipid transfer protein (LTP) group in yeast and human cells. They transfer phosphatidylserine (PS) from the endoplasmic reticulum (ER) to the plasma membrane (PM) via PS/phosphatidylinositol 4-phosphate (PI(4)P) exchange cycles. This finding allows a better understanding of how PS, which is critical for signaling processes, is distributed throughout the cell and the investigation of the link between this process and phosphoinositide (PIP) metabolism. The development of new fluorescence-based protocols has been instrumental in the discovery and characterization of this new cellular mechanism in vitro at the molecular level. This paper describes the production and the use of two fluorescently labelled lipid sensors, NBD-C2Lact and NBD-PHFAPP, to measure the ability of a protein to extract PS or PI(4)P and to transfer these lipids between artificial membranes. First, the protocol describes how to produce, label, and obtain high-purity samples of these two constructs. Secondly, this paper explains how to use these sensors with a fluorescence microplate reader to determine whether a protein can extract PS or PI(4)P from liposomes, using Osh6p as a case study. Finally, this protocol shows how to accurately measure the kinetics of PS/PI(4)P exchange between liposomes of defined lipid composition and to determine lipid transfer rates by fluorescence resonance energy transfer (FRET) using a standard fluorometer.

Here, we describe protocols using fluorescent lipid sensors and liposomes to determine whether a protein extracts and transports phosphatidylserine or phosphatidylinositol 4-phosphate in vitro.

Several members of the evolutionarily conserved oxysterol-binding protein (OSBP)-related proteins(ORP)/OSBP homologs (Osh) family have recently been found ...

SNARE-mediated Fusion of Single Proteoliposomes with Tethered Supported Bilayers in a Microfluidic Flow Cell Monitored by Polarized TIRF Microscopy

In the ubiquitous process of membrane fusion the opening of a fusion pore establishes the first connection between two formerly separate compartments. During neurotransmitter or hormone release via exocytosis, the fusion pore can transiently open and close repeatedly, regulating cargo release kinetics. Pore dynamics also determine the mode of vesicle recycling; irreversible resealing results in transient, &#34;kiss-and-run&#34; fusion, whereas dilation leads to full fusion. To better understand what factors govern pore dynamics, we developed an assay to monitor membrane fusion using polarized total internal reflection fluorescence (TIRF) microscopy with single molecule sensitivity and ~15 msec time resolution in a biochemically well-defined in vitro system. Fusion of fluorescently labeled small unilamellar vesicles containing v-SNARE proteins (v-SUVs) with a planar bilayer bearing t-SNAREs, supported on a soft polymer cushion (t-SBL, t-supported bilayer), is monitored. The assay uses microfluidic flow channels that ensure minimal sample consumption while supplying a constant density of SUVs. Exploiting the rapid signal enhancement upon transfer of lipid labels from the SUV to the SBL during fusion, kinetics of lipid dye transfer is monitored. The sensitivity of TIRF microscopy allows tracking single fluorescent lipid labels, from which lipid diffusivity and SUV size can be deduced for every fusion event. Lipid dye release times can be much longer than expected for unimpeded passage through permanently open pores. Using a model that assumes retardation of lipid release is due to pore flickering, a pore &#34;openness&#34;, the fraction of time the pore remains open during fusion, can be estimated. A soluble marker can be encapsulated in the SUVs for simultaneous monitoring of lipid and soluble cargo release. Such measurements indicate some pores may reseal after losing a fraction of the soluble cargo.

Here, we present a protocol to detect single, SNARE-mediated fusion events between liposomes and supported bilayers in microfluidic channels using polarized TIRFM, with single molecule sensitivity and ~15 msec time resolution. Lipid and soluble cargo release can be detected simultaneously. Liposome size, lipid diffusivity, and fusion pore properties are measured.

In the ubiquitous process of membrane fusion the opening of a fusion pore establishes the first connection between two formerly separate compartments. ...

Neuroscience

A Quantitative Fluorescence Microscopy-based Single Liposome Assay for Detecting the Compositional Inhomogeneity Between Individual Liposomes

Most research employing liposomes as membrane model systems or drug delivery carriers relies on bulk read-out techniques and thus intrinsically assumes all liposomes of the ensemble to be identical. However, new experimental platforms able to observe liposomes at the single-particle level have made it possible to perform highly sophisticated and quantitative studies on protein-membrane interactions or drug carrier properties on individual liposomes, thus avoiding errors from ensemble averaging. Here we present a protocol for preparing, detecting, and analyzing single liposomes using a fluorescence-based microscopy assay, facilitating such single-particle measurements. The setup allows for imaging individual liposomes in a massive parallel manner and is employed to reveal intra-sample size and compositional inhomogeneities. Additionally, the protocol describes the advantages of studying liposomes at the single liposome level, the limitations of the assay, and the important features to be considered when modifying it to study other research questions.

This protocol describes the fabrication of liposomes and how these can be immobilized on a surface and imaged individually in a massive parallel manner using fluorescence microscopy. This allows for the quantification of the size and compositional inhomogeneity between single liposomes of the population.

Most research employing liposomes as membrane model systems or drug delivery carriers relies on bulk read-out techniques and thus intrinsically assumes ...

Bioengineering

Quantification of Efferocytosis by Single-cell Fluorescence Microscopy

Studying the regulation of efferocytosis requires methods that are able to accurately quantify the uptake of apoptotic cells and to probe the signaling and cellular processes that control efferocytosis. This quantification can be difficult to perform as apoptotic cells are often efferocytosed piecemeal, thus necessitating methods which can accurately delineate between the efferocytosed portion of an apoptotic target versus residual unengulfed cellular fragments. The approach outlined herein utilizes dual-labeling approaches to accurately quantify the dynamics of efferocytosis and efferocytic capacity of efferocytes such as macrophages. The cytosol of the apoptotic cell is labeled with a cell-tracking dye to enable monitoring of all apoptotic cell-derived materials, while surface biotinylation of the apoptotic cell allows for differentiation between internalized and non-internalized apoptotic cell fractions. The efferocytic capacity of efferocytes is determined by taking fluorescent images of live or fixed cells and quantifying the amount of bound versus internalized targets, as differentiated by streptavidin staining. This approach offers several advantages over methods such as flow cytometry, namely the accurate delineation of non-efferocytosed versus efferocytosed apoptotic cell fractions, the ability to measure efferocytic dynamics by live-cell microscopy, and the capacity to perform studies of cellular signaling in cells expressing fluorescently-labeled transgenes. Combined, the methods outlined in this protocol serve as the basis for a flexible experimental approach that can be used to accurately quantify efferocytic activity and interrogate cellular signaling pathways active during efferocytosis.

Efferocytosis, the phagocytic removal of apoptotic cells, is required to maintain homeostasis and is facilitated by receptors and signaling pathways that allow for the recognition, engulfment, and internalization of apoptotic cells. Herein, we present a fluorescence microscopy protocol for the quantification of efferocytosis and the activity of efferocytic signaling pathways.

Studying the regulation of efferocytosis requires methods that are able to accurately quantify the uptake of apoptotic cells and to probe the signaling ...

Immunology and Infection

Flow Cytometric Analysis of Bimolecular Fluorescence Complementation: A High Throughput Quantitative Method to Study Protein-protein Interaction

Among methods to study protein-protein interaction inside cells, Bimolecular Fluorescence Complementation (BiFC) is relatively simple and sensitive. BiFC is based on the production of fluorescence using two non-fluorescent fragments of a fluorescent protein (Venus, a Yellow Fluorescent Protein variant, is used here). Non-fluorescent Venus fragments (VN and VC) are fused to two interacting proteins (in this case, AKAP-Lbc and PDE4D3), yielding fluorescence due to VN-AKAP-Lbc-VC-PDE4D3 interaction and the formation of a functional fluorescent protein inside cells.
BiFC provides information on the subcellular localization of protein complexes and the strength of protein interactions based on fluorescence intensity. However, BiFC analysis using microscopy to quantify the strength of protein-protein interaction is time-consuming and somewhat subjective due to heterogeneity in protein expression and interaction. By coupling flow cytometric analysis with BiFC methodology, the fluorescent BiFC protein-protein interaction signal can be accurately measured for a large quantity of cells in a short time. Here, we demonstrate an application of this methodology to map regions in PDE4D3 that are required for the interaction with AKAP-Lbc. This high throughput methodology can be applied to screening factors that regulate protein-protein interaction.

Flow cytometric analysis of Bimolecular Fluorescence Complementation provides a high throughput quantitative method to study protein-protein interaction. This methodology can be applied to mapping protein binding sites and for screening factors that regulate protein-protein interaction.

Among methods to study protein-protein interaction inside cells, Bimolecular Fluorescence Complementation (BiFC) is relatively simple and sensitive. BiFC ...

Biology

Membrane Transport Processes Analyzed by a Highly Parallel Nanopore Chip System at Single Protein Resolution

Membrane protein transport on the single protein level still evades detailed analysis, if the substrate translocated is non-electrogenic. Considerable efforts have been made in this field, but techniques enabling automated high-throughput transport analysis in combination with solvent-free lipid bilayer techniques required for the analysis of membrane transporters are rare. This class of transporters however is crucial in cell homeostasis and therefore a key target in drug development and methodologies to gain new insights desperately needed.
The here presented manuscript describes the establishment and handling of a novel biochip for the analysis of membrane protein mediated transport processes at single transporter resolution. The biochip is composed of microcavities enclosed by nanopores that is highly parallel in its design and can be produced in industrial grade and quantity. Protein-harboring liposomes can directly be applied to the chip surface forming self-assembled pore-spanning lipid bilayers using SSM-techniques (solid supported lipid membranes). Pore-spanning parts of the membrane are freestanding, providing the interface for substrate translocation into or out of the cavity space, which can be followed by multi-spectral fluorescent readout in real-time. The establishment of standard operating procedures (SOPs) allows the straightforward establishment of protein-harboring lipid bilayers on the chip surface of virtually every membrane protein that can be reconstituted functionally. The sole prerequisite is the establishment of a fluorescent read-out system for non-electrogenic transport substrates.
High-content screening applications are accomplishable by the use of automated inverted fluorescent microscopes recording multiple chips in parallel. Large data sets can be analyzed using the freely available custom-designed analysis software. Three-color multi spectral fluorescent read-out furthermore allows for unbiased data discrimination into different event classes, eliminating false positive results.
The chip technology is currently based on SiO2 surfaces, but further functionalization using gold-coated chip surfaces is also possible.

The presented protocol describes the analysis of membrane protein mediated transport on the single transporter level using pore-spanning solvent-free lipid bilayers. This is achieved by the creation of bulk produced nanopore array chips, combined with highly parallel data acquisition and analysis, enabling the future establishment of membrane protein effector screenings.

Membrane protein transport on the single protein level still evades detailed analysis, if the substrate translocated is non-electrogenic. Considerable ...

Flow Cytometry-Based Assays for Microglial Engulfment of Synaptic Materials

Quantification of Microglial Engulfment of Synaptic Material Using Flow Cytometry

Microglia play a pivotal role in synaptic refinement in the brain. Analysis of microglial engulfment of synapses is essential for comprehending this process; however, currently available methods for identifying microglial engulfment of synapses, such as immunohistochemistry (IHC) and imaging, are laborious and time-intensive. To address this challenge, herein we present in vitro and in vivo* assays that allow fast and high-throughput quantification of microglial engulfment of synapses using flow cytometry.
In the in vivo* approach, we performed intracellular vGLUT1 staining following fresh cell isolation from adult mouse brains to quantify engulfment of vGLUT1+ synapses by microglia. In the in vitro synaptosome engulfment assay, we used freshly isolated cells from the adult mouse brain to quantify the engulfment of pHrodo&#160;Red-labeled synaptosomes by microglia. These protocols together provide a time-efficient approach to quantifying microglial engulfment of synapses and represent promising alternatives to labor-intensive image analysis-based methods. By streamlining the analysis, these assays can contribute to a better understanding of the role of microglia in synaptic refinement in different disease models.

Author Spotlight: Advancing Understanding Through Technological Innovations in Psychoneuroimmunology

Here we present two protocols to quantify microglial engulfment of vGLUT1-positive synapses and pHRodo&#160;Red-labeled crude synaptosomes using flow cytometry.

Microglia play a pivotal role in synaptic refinement in the brain. Analysis of microglial engulfment of synapses is essential for comprehending this ...

Live Fluorescence, Inverse Imaging of Cell Ruffling, and Macropinocytosis

Macropinocytosis is a highly conserved but still incompletely understood process that is essential for the uptake and ingestion of fluid, fluid-phase nutrients and other material in cells. The dramatic extension of cell surface ruffles, their closure to form macropinosomes, and the maturation of internalized macropinosomes are key events in this pathway that can be difficult to capture using conventional confocal imaging based on tracking a bolus of fluorescent cargo. Fluorescent dextrans are commonly used experimentally as fluid phase markers for macropinosomes and for other endocytic pathways. A method the lab has adopted to optimize the imaging of dextran uptake involves using live imaging of cells bathed in high concentrations of fluorescent dextran in the medium, with the unlabeled cells appearing in relief (as black). The cell ruffles are highlighted to visualize ruffle closure, and internalized macropinosomes appear as fluorescent vacuoles in the cell interior. This method is optimal for visualizing macropinosome features and allows for easy segmentation and quantification. This paper describes dual-labeling of pathways with different sized dextrans and the co-expression of lipid probes and fluorescent membrane proteins to demark macropinosomes and other endosomes. The detection of internalized dextran at an ultrastructural level using correlative light and electron microscopy (CLEM) is also demonstrated. These cell processes can be imaged using multiple live imaging modalities, including in 3D. Taken together, these approaches optimize macropinosome imaging for many different settings and experimental systems.

This protocol demonstrates dextran imaging in live cells using continuous uptake and inverse images to optimize visualization of ruffling, macropinosome maturation, and analysis of dextran and other cell labelings.

Macropinocytosis is a highly conserved but still incompletely understood process that is essential for the uptake and ingestion of fluid, fluid-phase ...

Production and Imaging of Recombinant Protein-Filled Vesicles from E. coli

Optimized Production and Analysis of Recombinant Protein-Filled Vesicles from E. coli

This innovative system, using a short peptide tag, that exports multiple recombinant proteins in membrane bound vesicles from E. coli, provides an effective solution to a range of problems associated with bacterial recombinant protein expression. These recombinant vesicles compartmentalise proteins within a micro-environment that facilitates the production of otherwise challenging, toxic, insoluble, or disulfide-bond containing proteins from bacteria. Protein yield is increased considerably when compared to typical bacterial expression in the absence of the vesicle-nucleating peptide tag. The release of vesicle-packaged proteins supports isolation from the culture medium&#160;and permits long-term active protein storage. This technology gives rise to increased yields of vesicle-packaged, functional proteins for simplified downstream processing for a diverse range of applications from applied biotechnology to discovery science and medicine. In the present article and the associated video, a detailed protocol of the method is provided, which highlights key steps in the methodology to maximize recombinant protein-filled vesicle production.

Author Spotlight: The Production of Recombinant Proteins 

The present protocol describes a detailed method for the bacterial production of recombinant proteins, including typically insoluble or disulfide-bond containing proteins, packaged inside extracellular membrane-bound vesicles. This has the potential to be applied to versatile areas of scientific research, including applied biotechnology and medicine.

This innovative system, using a short peptide tag, that exports multiple recombinant proteins in membrane bound vesicles from E. coli, provides an ...

Quantitative Flow Cytometry to Study Labeled Protein-Phospholipid Vesicle Interactions

Source: Podoplelova, N. et al., <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/63459">Analyzing the Interaction of Fluorescent-Labeled Proteins with Artificial Phospholipid Microvesicles using Quantitative Flow Cytometry.</a> J. Vis. Exp. (2022)
In this video, we demonstrate the interaction between proteins and artificial phospholipid vesicles using quantitative flow cytometry. The binding of fluorescently-labeled proteins to fluorescently-labeled phospholipid vesicles increases the mean fluorescence intensity, and this increase is detected by a flow cytometer.

Source: Podoplelova, N. et al., Analyzing the Interaction of Fluorescent-Labeled Proteins with Artificial Phospholipid Microvesicles using Quantitative ...

Analyzing the Interaction of Fluorescent-Labeled Proteins with Artificial Phospholipid Microvesicles using Quantitative Flow Cytometry

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