This study was supported by the Velux Stiftung (A.K.M.), NIH grant EY024207 (A.K.M.) and the Austrian Science Fund (FWF) project J3686 (B.P.).

1. Preparation of Liposomes and Proteoliposomes
<ol>
	<li>Liposome Formation
	<ol>
		<li>Using a glass syringe, add 1,435 &#181;l POPC (25 mg/ml, in chloroform) and 160 &#181;l POPG (25 mg/ml, in chloroform) to a round bottom flask to obtain 52.5 &#181;mol lipids in a molar ratio of POPC:POPG = 9:1.</li>
		<li>Dry the lipids for 30 min using a rotary evaporator at a rotation speed of 145 rpm (no water bath is needed for this volume of solvent), then transfer the flask to a vacuum desiccator for at least 3 hr, or overnight, at room temperature (RT).</li>
		<li>Hydrate the dried lipid film with 10 ml of 50 mM HEPES pH 7.4, 100 mM NaCl (henceforth referred to as buffer A) by gently swirling the flask until a homogenous, turbid suspension results; 
		NOTE: The lipid concentration at this stage is expected to be 5.25 mM as no losses should have occurred so far.</li>
		<li>Sonicate the suspension in a water bath for 10 min at room temperature with a frequency of 40 kHz. The solution will look somewhat clearer.</li>
		<li>Using an extruder, pass the suspension 10 times through a membrane with a 400 nm pore size, followed by a second cycle of extrusion with 4 passes through a membrane with a 200 nm pore size. 
		NOTE: The mean diameter of the resulting LUVs is ~175 nm. If necessary, the size and homogeneity of the LUVs can be checked by Dynamic Light Scattering according to the manufacturer&#39;s instructions.</li>
		<li>Quantify the phospholipid concentration of the LUV suspension as described in section 1.2). 
		NOTE: Because of losses during extrusion, the concentration is usually around 3.6 mM.</li>
		<li>Store the LUVs at 4 &#176;C for about 2 weeks if not used immediately.</li>
	</ol>
	</li>
	<li>Phospholipid Quantification 
	NOTE: To determine the phospholipid concentration of the LUV suspension used for reconstitution, as well as that of the proteoliposomes that are eventually generated, an aliquot of the sample is subjected to oxidation by perchloric acid. This procedure breaks down phospholipids to release inorganic phosphate that is then quantified by a colorimetric assay in comparison with standards 23.
	<ol>
		<li>Prepare a 40 mM stock solution of sodium phosphate (Na2HPO4) in deionized distilled water.</li>
		<li>Dilute the stock solution with deionized distilled water to obtain 4 mM and 0.4 mM working solutions that will serve as calibration standards.</li>
		<li>Using the working solutions, prepare standards in 13 x 100 mm2 glass tubes ranging from 0 to 80 nmol sodium phosphate in a final volume of 50 &#181;l.</li>
		<li>Take 10 &#181;l each of the LUV and proteoliposome samples to be quantified and dilute with 40 &#181;l of ddH2O in 13 x 100 mm2 glass tubes. 
		NOTE: As the lipid concentration of the LUVs and proteoliposomes is in the range 2.5-5 &#181;M, the 10 &#181;l of sample should contain 25-50 nmol lipid phosphorus.</li>
		<li>Add 300 &#181;l perchloric acid to each of the standards and samples and heat for 1 hr at 145 &#176;C in a heating block. Put marbles on the tubes to prevent evaporation.</li>
		<li>Let the tubes cool to room temperature and add 1 ml of ddH2O.</li>
		<li>Add 400 &#181;l each of freshly prepared 12 g/L ammonium molybdate and 50 g/L sodium ascorbate and vortex to mix.</li>
		<li>Heat for 10 min at 100 &#176;C with marbles on top of the tubes. Remove the tubes from the heating block and let them cool to room temperature.</li>
		<li>Measure the absorbance of the samples against the blank (standard sample containing 0 nmol sodium phosphate) with a spectrometer at a wavelength of 797 nm.</li>
		<li>Determine the phosphate content of samples against the calibration standard curve.</li>
	</ol>
	</li>
	<li>Reconstitution of Opsin 
	NOTE: It is necessary to determine optimal swelling conditions for reconstitution as these depend on the nature of the detergent, as well as the lipid composition and concentration of the LUVs. As LUVs change their light scattering properties on swelling the process can be monitored by measuring absorbance (Figure 2) as reviewed by Rigaud and Levy 24 and Geertsma et al. 25.
	<ol>
		<li>Pipette 800 &#181;l of LUVs (from section 1.1; as lipid recovery after extrusion is 70%, the expected concentration of LUVs is 3.6 mM phospholipid) into a 2 ml microfuge tube.</li>
		<li>Add 5.3 &#181;l of buffer A and 34.7 &#181;l of 10% (w/v) DDM dissolved in buffer A.</li>
		<li>Incubate for 3 hr at room temperature with end-over-end mixing.</li>
		<li>Meanwhile prepare the polystyrene beads:
		<ol>
			<li>Use 400 mg of beads per sample and weigh them out in a glass beaker.</li>
			<li>Wash them twice with methanol, three times with water and once with buffer A. For each washing step use 5 ml of liquid and stir slowly for 10 min. 
			NOTE: It is recommended to prepare the polystyrene beads for several samples at once, e.g., weigh in 6 g of beads and wash with 75 ml of liquid. Excess beads can be stored in a refrigerator for several days.</li>
		</ol>
		</li>
		<li>During the last 30 min of vesicle destabilization dry the NBD-labeled phospholipid in a screw-cap glass tube (&#39;reconstitution glass tube&#39;): Per sample, 9.5 &#181;l of NBD-PC (1 mg/ml in chloroform, yielding 0.4 mol% of total phospholipids) are dried under a stream of nitrogen in a glass tube and subsequently dissolved in 45 &#181;l of 0.1% (w/v) DDM in buffer A.</li>
		<li>After 3 hr of vesicle destabilization add the dissolved-NBD-labeled phospholipid, the DDM-solubilized protein and buffer A such that the final volume of 1 ml contains 0.36% (w/v), i.e., 7 mM, DDM.
		<ol>
			<li>Thus, to generate protein-free liposomes (used as a control sample) add 45 &#181;l of NBD-PC (dissolved in 0.1% DDM), 60 &#181;l of 0.1% DDM and 55 &#181;l of buffer A; for proteoliposomes add, for example, 40 &#181;l of protein (from a typical stock solution of ~110 ng/&#181;l) in 0.1% DDM, 45 &#181;l of NBD-PC (dissolved in 0.1% DDM), 20 &#181;l of 0.1% DDM and 55 &#181;l of buffer A. 
			NOTE: The order of addition should be as listed.</li>
		</ol>
		</li>
		<li>Mix the sample for an additional hr end over end at room temperature.</li>
		<li>Add 80 mg of the prepared polystyrene beads and incubate the sample with end-over-end mixing for 1 hr at RT.</li>
		<li>Next add an additional 160 mg of polystyrene beads and incubate with end-over-end mixing for a further 2 hr at RT.</li>
		<li>Transfer the sample (leaving the spent polystyrene beads behind) to a glass screw-cap tube containing 160 mg of fresh polystyrene beads and mix overnight at 4 &#176;C. 
		NOTE: The easiest way to transfer the sample and avoid sucking up beads is to use a Pasteur pipette that is pushed to the bottom of the glass tube with a little positive pressure. Once the tip of the pipette is firmly at the bottom of the tube, then the sample can be withdrawn easily without interference from the beads.</li>
		<li>The next morning, transfer the sample to a microfuge tube without carrying over beads and place on ice in preparation for the scramblase activity assay.</li>
	</ol>
	</li>
</ol>
2. Scramblase Activity Assay
NOTE: The fluorescence intensity of liposomes or proteoliposomes diluted with buffer A is monitored over time upon the addition of dithionite in a fluorescence spectrometer. To obtain a stable starting intensity, the fluorescence is recorded for at least 50 sec (or until a stable signal is achieved) before adding dithionite to a constantly stirred sample and is then followed for at least 500 sec after adding dithionite.
<ol>
	<li>Add 1,950 &#181;l of buffer A to a plastic cuvette containing a mini stir-bar.</li>
	<li>Add 50 &#181;l of the prepared proteo(liposomes) and let the sample equilibrate in the fluorescence spectrometer with constant stirring for several sec.</li>
	<li>Meanwhile prepare a solution of 1 M dithionite in 0.5 M unbuffered Tris (e.g., for two samples weigh out 20 mg of dithionite in a microfuge tube and dissolve in 114 &#181;l ice-cold 0.5 M Tris directly before use and keep on ice for the next sample). 
	NOTE: The dithionite solution must be prepared freshly and should not be used more than 20 min after preparation; if many measurements are to be made, aliquots of dithionite can be weighed out in advance and dissolved right before use.</li>
	<li>Start the fluorescence monitoring (excitation 470 nm, emission 530 nm, slit width 0.5 nm).</li>
	<li>Add 40 &#181;l of the 1 M dithionite solution to the cuvette 50 sec after starting the fluorescence recording (use the septum in the lid of the cuvette chamber if possible) and continue to record the fluorescence for a further 400-600 sec.</li>
	<li>Analyze the data as described in section 3.</li>
</ol>
3. Data Analysis
<ol>
	<li>Kinetics of Scrambling
	<ol>
		<li>Characterize the fluorescence trace of each sample obtained by the scramblase activity assay by defining the initial fluorescence, Fi, prior to adding dithionite, and the end-point fluorescence, F, reached after &#62;400 sec. Fi is determined for each sample as the mean value of fluorescence for the 30 sec period prior to addition of dithionite.</li>
		<li>Determine the end-point data corresponding to the extent of fluorescence reduction, R = 100&#8226;F/Fi. We use the terms RL for protein-free liposomes and RP for opsin-containing proteoliposomes.</li>
	</ol>
	</li>
	<li>Determining the Molecular Weight of the Functionally Reconstituted Scramblase
	<ol>
		<li>Convert the fluorescence reduction data according to the following equation: 
		p(&#8805;1 scramblase) = (RP &#8722; RL)/(Rmax &#8722; RL) (Equation 1) 
		NOTE: Where Rmax is the maximum reduction that is obtained when sufficient protein is reconstituted such that all vesicles in the sample possess at least one functional scramblase, and p is the probability that a particular vesicle in a reconstituted sample is &#39;scramblase-active&#39;, i.e., it possesses at least one functional scramblase. The value for RL is typically 45% 3 whereas Rmax is typically 82.5% 3 , instead of the expected 100% (Rmax can be experimentally determined for opsin proteoliposomes with a PPR of &#62;1 mg/mmol). As Rmax &#60;100% it is assumed that a sub-population of vesicles is refractory to reconstitution. For Rmax = 82.5%, the fraction of vesicles that is unable to accept protein is 0.35.</li>
		<li>Describe the relationship between p(&#8805;1 scramblase) and PPR (mg protein/mmol phospholipid) by Poisson statistics as follows: 
		p(&#8805;1 scramblase) = 1 - e-m = 1 - exp(-PPR/&#945;) (Equation 2) 
		NOTE: Where m = number of scramblases per vesicle and &#945; = mono-exponential fit constant in units of mg protein/mmol phospholipid.</li>
		<li>As a fraction of the vesicles does not contribute to scrambling even at high PPR (see discussion), modify the equation to: 
		p(&#8805;1 scramblase) = 1 - exp(-PPR*/&#945;) (Equation 3) 
		NOTE: Where PPR* = PPR/(1-f), where f is the refractory population of vesicles or, in this case, PPR* = PPR/0.65 (Equation 4). 
		NOTE: The fit constant &#945; is determined by fitting a graph of p(&#8805;1 scramblase) versus PPR* with a mono-exponential function. If opsin (molecular weight 41.7 kDa) functionally reconstitutes into 175-nm-diameter vesicles (each vesicle has 280,000 phospholipids 26) as a monomer, &#945; = 0.187 mg mmol&#8722;1. If opsin dimerizes prior to reconstitution 3 to yield scramblase-active vesicles, then &#945; = 0.37 mg mmol&#8722;1. If PPR rather than PPR* were to be used for the analysis, then the corresponding &#945; values would be 0.122 and 0.244 mg mmol&#8722;1. These predicted values for &#945; assume that all opsin molecules are functionally competent. If only a fraction of the molecules is competent to scramble lipids, then the corresponding values of &#945; will be larger.</li>
	</ol>
	</li>
</ol>

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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phospholipid+flippase+activity+of+the+reconstituted+P-glycoprotein+multidrug+transporter.">Phospholipid flippase activity of the reconstituted P-glycoprotein multidrug transporter.</a> Biochemistry. 40, 6937-6947 (2001).</li><li>Zhou, X., Graham, T. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reconstitution+of+phospholipid+translocase+activity+with+purified+Drs2p,+a+type-IV+P-type+ATPase+from+budding+yeast.">Reconstitution of phospholipid translocase activity with purified Drs2p, a type-IV P-type ATPase from budding yeast.</a> Proc. Natl. Acad. Sci. U. S. A. 106, 16586-16591 (2009).</li></ol>

The authors have nothing to disclose.

The scramblase activity assay enabled us originally to determine that opsin has phospholipid scramblase activity 2. The assay also allowed us to characterize opsin&#39;s scramblase activity by testing specificity (we used a variety of NBD-labeled reporter lipids such as NBD-phosphatidylethanolamine, labeled with NBD on an acyl chain as shown for NBD-PC in Figure 1A, or on the headgroup, NBD-sphingomyelin or NBD- phosphatidylserine 2), the effect of vesicle lipid composition (e.g...

The photoreceptor rhodopsin, a prototypical G protein-coupled receptor (reviewed for example in reference 1), is the first phospholipid scramblase to be identified and biochemically verified 2,3. Scramblases are phospholipid transporters that increase the intrinsically slow rate of transbilayer phospholipid movement to physiologically appropriate levels in a bidirectional, ATP-independent manner 4-6. Examples of their actions can be found in the endoplasmic reticulum and bacterial cytoplasmic membrane where constitutive scrambling is needed for membrane homeostasis and growth, as well as for a variety of glycosylation pathways 5. Regulated phospholipid scrambling is needed to expose phosphatidylserine (PS) on the surface of apoptotic cells where it acts as an &#34;eat-me&#34;-signal for macrophages 7 and provides a procoagulant surface on activated blood platelets to catalyze the production of protein factors needed for blood clotting. In photoreceptor disc membranes, rhodopsin&#39;s scrambling activity has been suggested to counteract the phospholipid imbalance between the two membrane leaflets of the bilayer that is generated by the ATP-dependent, unidirectional lipid flippase ABCA4 4,8,910-12.
Despite the physiological importance of scramblases, their identity remained elusive until rhodopsin was reported as a scramblase in photoreceptor discs 2, members of the TMEM16 protein family were identified as Ca2+-dependent scramblases needed for PS exposure at the plasma membrane (reviewed in reference 13), and the bacterial protein FtsW was proposed as a Lipid II scramblase required for peptidoglycan synthesis 14. These discoveries were based on the reconstitution of purified proteins in liposomes and demonstration of scramblase activity in the resulting proteoliposomes using the methodology described here. Other potential scramblases 15-21&#160;&#8212; the MurJ and AmJ proteins implicated in peptidoglycan biosynthesis, WzxE and related proteins implicated in scrambling O-antigen precursors, MprF protein needed to translocate aminoacylated phosphatidylglycerol across the bacterial cytoplasmic membrane, and Xkr8 family members that have been proposed to expose PS on the surface of apoptotic cells&#160;&#8212; remain to be tested biochemically. This highlights the importance of a robust assay to identify and characterize scramblase activity.
Here, we describe the reconstitution of purified opsin, the apoprotein of the photoreceptor rhodopsin, into large unilamellar vesicles (LUVs), and subsequent analysis of scramblase activity in the resulting proteoliposomes using a fluorescence-based assay. There are several well-described protocols available in the literature for the heterologous expression and purification of opsin, therefore we will not describe it in this protocol; we use the protocols described in Goren et al. 3 which yields FLAG-tagged, thermostable opsin at about 100 ng/&#181;l in 0.1% (w/v) dodecylmaltoside (DDM).
Reconstitution is achieved by treating LUVs with sufficient detergent so that they swell but do not dissolve. Under these conditions, a membrane protein&#160;&#8212; supplied in the form of protein-detergent micelles&#160;&#8212; will integrate into the liposomes and become reconstituted into the liposome membrane upon detergent removal, resulting in proteoliposomes. To reconstitute opsin (obtained as a purified protein in 0.1% (w/v) DDM), LUVs are prepared from a mixture of POPC (1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine) and POPG (1-palmitoyl-2-oleoyl-sn-glycero-3-[phospho-rac-(1-glycerol)]) and saturated with DDM before adding opsin and NBD-PC. The detergent is then removed by treating the sample with polystyrene beads.
The principle underlying the fluorescence-based assay is shown in Figure 1B. LUVs are symmetrically reconstituted with a trace amount of NBD-PC or other NBD-labeled fluorescent phospholipid reporter (Figure 1A). On adding dithionite, a membrane-impermeant dianion, NBD-PC molecules in the outer leaflet of the LUVs are rendered non-fluorescent as the nitro-group of NBD is reduced to a non-fluorescent amino-group. As neither NBD-PC molecules nor dithionite are able to traverse the membrane on the time-scale of the experiment (&#60;10 min), this results in 50% reduction of the fluorescent signal. However, if the liposomes are reconstituted with a scramblase, NBD-PC molecules in the inner leaflet can scramble rapidly to the outside where they are reduced. This results in the total loss of fluorescence in the ideal case (Figure 1C).
<img alt="Figure 1" src="/files/ftp_upload/54635/54635fig1.jpg" /> 
Figure 1: Schematic representation of the scramblase activity assay. The assay uses a fluorescent NBD-labeled reporter lipid; NBD-PC is shown (A). Large unilamellar vesicles are reconstituted with a trace amount of NBD-PC. Reconstitution produces symmetric vesicles, with NBD-PC distributed equally in the outer and inner leaflets. Dithionite (S2O42-) chemically reduces the nitro-group of NBD to a non-fluorescent amino-group. Treatment of protein-free liposomes with dithionite (B, top) causes a 50% reduction of fluorescence since only the NBD-PC molecules in the outer leaflet are reduced: dithionite is negatively charged and cannot cross the membrane to react with NBD-PC molecules in the inner leaflet. Dithionite treatment of opsin-containing proteoliposomes (B, bottom), i.e., scramblase-active proteoliposomes, results in 100% loss of fluorescence as opsin facilitates movement of NBD-PC between the inner and the outer leaflet. (C) shows idealized fluorescence traces obtained on treating protein-free liposomes and opsin-containing proteoliposomes with dithionite. The rate of fluorescence loss is the same in both cases indicating that the chemical reduction of NBD by dithionite is rate-limiting, and that scrambling occurs at a rate equal to or greater than the rate of the chemical reaction. Traces obtained from an actual experiment are shown in Figure 3. <a href="https://www.jove.com/files/ftp_upload/54635/54635fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The methods we describe can be used to reconstitute and assay other purified proteins, as well as mixtures of membrane proteins obtained, for example, by extracting microsomes with detergent 22.

<table><tbody><tr><td>1-palmitoyl-2-oleoyl-&lt;em&gt;sn&lt;/em&gt;-glycero-3-phosphocholine</td><td>Avanti Polar Lipids</td><td>850457C</td><td>POPC</td></tr><tr><td>1-Palmitoyl-2-oleoyl-&lt;em&gt;sn&lt;/em&gt;-glycero-3-phospho-rac-(1-glycerol) (sodium salt)</td><td>Avanti Polar Lipids</td><td>840457C</td><td>POPG</td></tr><tr><td>1-palmitoyl-2-{6-[7-nitro-2-1,3-benzoxadiazol-4-yl)amino]hexanoyl}-&lt;em&gt;sn&lt;/em&gt;-glycero-3-phosphocholine</td><td>Avanti Polar Lipids</td><td>810130C</td><td>NBD-PC</td></tr><tr><td>4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid</td><td>VWR Scientific</td><td>EM-5330</td><td>HEPES</td></tr><tr><td>NaCl</td><td>Sigma</td><td>S7653-1KG</td><td>NaCl</td></tr><tr><td>Dodecyl-&amp;beta;-D-maltoside</td><td>Anatrace</td><td>D310 5 GM</td><td>DDM</td></tr><tr><td>Fluorimeter cuvettes</td><td>sigma</td><td>C0918-100EA</td><td>cuvettes</td></tr><tr><td>Spectrofluorometer</td><td>Photon Technology International, Inc.</td><td>fluorimeter</td><td></td></tr><tr><td>Sodium hydrosulfite technical grade, 85%</td><td>Sigma</td><td>157953-5G</td><td>dithionite</td></tr><tr><td>GraphPad Prism 5 software</td><td></td><td></td><td>Prism</td></tr><tr><td>Tris Base</td><td>VWR</td><td>JTX171-3</td><td>Tris</td></tr><tr><td>LIPEX 10 ml extruder&amp;nbsp;</td><td>Northern Lipids, Inc.</td><td></td><td>Extruder</td></tr><tr><td>Whatman, Drain disc, PE, 25 mm</td><td>Sigma</td><td>28156-243</td><td>Disc support</td></tr><tr><td>Whatman Nuclepore Track-Etched Membranes, 0.4 &amp;micro;m, 25 mm diameter</td><td>Sigma</td><td>WHA110607</td><td>400 nm membrane</td></tr><tr><td>Whatman Nucleopore Track-Etched Membranes, 0.2 &amp;micro;m, 25 mm diameter</td><td>Sigma</td><td>WHA110606</td><td>200 nm membrane</td></tr><tr><td>sodium phosphate</td><td>Sigma</td><td>S3264-500G</td><td></td></tr><tr><td>VWR Culture Tubes, Disposable, Borosilicate Glass, 13 x 100 mm</td><td>VWR Scientific</td><td>47729-572</td><td>glass tubes</td></tr><tr><td>Perchloric acid</td><td>Sigma</td><td>30755-500ML</td><td></td></tr><tr><td>Ammonium Molybdate Tetrahydrate</td><td>Sigma</td><td>A-7302</td><td>ammonium molybdate</td></tr><tr><td>(+)-Sodium L-ascorbate</td><td>Sigma</td><td>A7631-25G</td><td>sodium ascorbate</td></tr><tr><td>Bio-Beads SM2 adsorbents</td><td>Bio Rad</td><td>1523920</td><td>polystyrene beads</td></tr><tr><td>&amp;nbsp;2.0 ml Microtubes clear</td><td>VWR Scientific</td><td>10011-742</td><td>Reconstitution tubes</td></tr><tr><td>Reconstitution glass tube</td><td>VWR Scientific</td><td>53283-800</td><td>Reconstitution glass tubes</td></tr><tr><td>Zetasizer&amp;nbsp;</td><td>Malvern&amp;nbsp;</td><td></td><td>DLS</td></tr></tbody></table>

a fluorescence-based assay of phospholipid scramblase activity

Scramblases translocate phospholipids across the membrane bilayer bidirectionally in an ATP-independent manner. The first scramblase to be identified and biochemically verified was opsin, the apoprotein of the photoreceptor rhodopsin. Rhodopsin is a G protein-coupled receptor localized in rod photoreceptor disc membranes of the retina where it is responsible for the perception of light. Rhodopsin&#39;s scramblase activity does not depend on its ligand 11-cis-retinal, i.e., the apoprotein opsin is also active as a scramblase. Although constitutive and regulated phospholipid scrambling play an important role in cell physiology, only a few phospholipid scramblases have been identified so far besides opsin. Here we describe a fluorescence-based assay of opsin&#39;s scramblase activity. Opsin is reconstituted into large unilamellar liposomes composed of phosphatidylcholine, phosphatidylglycerol and a trace quantity of fluorescent NBD-labeled PC (1-palmitoyl-2-{6-[7-nitro-2-1,3-benzoxadiazole-4-yl)amino]hexanoyl}-sn-glycero-3-phosphocholine). Scramblase activity is determined by measuring the extent to which NBD-PC molecules located in the inner leaflet of the vesicle are able to access the outer leaflet where their fluorescence is chemically eliminated by a reducing agent that cannot cross the membrane. The methods we describe have general applicability and can be used to identify and characterize scramblase activities of other membrane proteins.

We describe the reconstitution of opsin into LUVs to characterize its scramblase activity using a fluorescence-based assay. We analyze the results to place a lower limit on the rate of opsin-mediated phospholipid scrambling and to determine the oligomeric state in which opsin functionally reconstitutes into the vesicles.
To identify optimal reconstitution conditions, it is necessary to determine empirically the amount of deterge...

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A Fluorescence-based Assay of Phospholipid Scramblase Activity

We describe a fluorescence-based assay to measure phospholipid scrambling in large unilamellar liposomes reconstituted with opsin.

Scramblases translocate phospholipids across the membrane bilayer bidirectionally in an ATP-independent manner. The first scramblase to be identified and ...

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Biochemistry

13.8K Views.  Weill Cornell Medical College. We describe a fluorescence-based assay to measure phospholipid scrambling in large unilamellar liposomes reconstituted with opsin. 

Video: A Fluorescence-based Assay of Phospholipid Scramblase Activity

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 ...

Defining Substrate Specificities for Lipase and Phospholipase Candidates

Microorganisms produce a wide spectrum of (phospho)lipases that are secreted in order to make external substrates available for the organism. Alternatively, other (phospho)lipases may be physically associated with the producing organism causing a turnover of intrinsic lipids and frequently giving rise to a remodeling of the cellular membranes. Although potential (phospho)lipases can be predicted with a number of algorithms when the gene/protein sequence is available, experimental proof of the enzyme activities, substrate specificities, and potential physiological functions has frequently not been obtained. This manuscript describes the optimization of assay conditions for prospective (phospho)lipases with unknown substrate specificities and how to employ these optimized conditions in the search for the natural substrate of a respective (phospho)lipase. Using artificial chromogenic substrates, such as p-nitrophenyl derivatives, may help to detect a minor enzymatic activity for a predicted (phospho)lipase under standard conditions. Having encountered such a minor enzymatic activity, the distinct parameters of an enzyme assay can be varied in order to obtain a more efficient hydrolysis of the artificial substrate. After having determined the conditions under which an enzyme works well, a variety of potential natural substrates should be assayed for their degradation, a process that can be followed employing distinct chromatographic methods. The definition of substrate specificities for new enzymes, often provides hypotheses for a potential physiological role of these enzymes, which then can be tested experimentally. Following these guidelines, we were able to identify a phospholipase C (SMc00171) that degrades phosphatidylcholine to phosphocholine and diacylglycerol, in a crucial step for the remodeling of membranes in the bacterium Sinorhizobium meliloti upon phosphorus-limiting conditions of growth. For two predicted patatin-like phospholipases (SMc00930 and SMc01003) of the same organism, we could redefine their substrate specificities and clarify that SMc01003 is a diacylglycerol lipase.

Many predicted (phospho)lipases are poorly characterized with regard to their substrate specificities and physiological functions. Here we provide a protocol to optimize enzyme activities, search for natural substrates, and propose physiological functions for these enzymes.

Microorganisms produce a wide spectrum of (phospho)lipases that are secreted in order to make external substrates available for the organism. ...

Click-Chemistry Based Fluorometric Assay for Apolipoprotein N-acyltransferase from Enzyme Characterization to High-Throughput Screening

Lipoproteins from proteobacteria are posttranslationally modified by fatty acids derived from membrane phospholipids by the action of three integral membrane enzymes, resulting in triacylated proteins. The first step in the lipoprotein modification pathway involves the transfer of a diacylglyceryl group from phosphatidylglycerol onto the prolipoprotein, resulting in diacylglyceryl prolipoprotein. In the second step, the signal peptide of prolipoprotein is cleaved, forming an apolipoprotein, which in turn is modified by a third fatty acid derived from a phospholipid. This last step is catalyzed by apolipoprotein N-acyltransferase (Lnt). The lipoprotein modification pathway is essential in most &#947;-proteobacteria, making it a potential target for the development of novel antibacterial agents. Described here is a sensitive assay for Lnt that is compatible with high-throughput screening of small inhibitory molecules. The enzyme and substrates are membrane-embedded molecules; therefore, the development of an in vitro test is not straightforward. This includes the purification of the active enzyme in the presence of detergent, the availability of alkyne-phospholipids and diacylglyceryl peptide substrates, and the reaction conditions in mixed micelles. Furthermore, in order to use the activity test in a high-throughput screening (HTS) setup, direct readout of the reaction product is preferred over coupled enzymatic reactions. In this fluorometric enzyme assay, the alkyne-triacylated peptide product is rendered fluorescent through a click-chemistry reaction and detected in a multiwell plate format. This method is applicable to other acyltransferases that use fatty acid-containing substrates, including phospholipids and acyl-CoA.

Presented here is a sensitive fluorescence assay to monitor apolipoprotein N-acyltransferase activity using diacylglyceryl peptide and alkyne-phospholipids as substrates with click-chemistry.

Lipoproteins from proteobacteria are posttranslationally modified by fatty acids derived from membrane phospholipids by the action of three integral ...

Fluorescence Recovery after Merging a Droplet to Measure the Two-dimensional Diffusion of a Phospholipid Monolayer

We introduce a new method to measure the lateral diffusivity of a surfactant monolayer at the fluid-fluid interface, called fluorescence recovery after merging (FRAM). FRAM adopts the same principles as the fluorescence recovery after photobleaching (FRAP) technique, especially for measuring fluorescence recovery after bleaching a specific area, but FRAM uses a drop coalescence instead of photobleaching dye molecules to induce a chemical potential gradient of dye molecules. Our technique has several advantages over FRAP: it only requires a fluorescence microscope rather than a confocal microscope equipped with high power lasers; it is essentially free from the selection of fluorescence dyes; and it has far more freedom to define the measured diffusion area. Furthermore, FRAM potentially provides a route for studying the mixing or inter-diffusion of two different surfactants, when the monolayers at a surface of droplet and at a flat air/water interface are prepared with different species, independently.

We present a new technique to measure the lateral diffusion of a surface active species at the fluid-fluid interface by merging a droplet monolayer onto a flat monolayer.

We introduce a new method to measure the lateral diffusivity of a surfactant monolayer at the fluid-fluid interface, called fluorescence recovery after ...

Bioengineering

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

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

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

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

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 ...

Gramicidin-based Fluorescence Assay; for Determining Small Molecules Potential for Modifying Lipid Bilayer Properties

Many drugs and other small molecules used to modulate biological function are amphiphiles that adsorb at the bilayer/solution interface and thereby alter lipid bilayer properties. This is important because membrane proteins are energetically coupled to their host bilayer by hydrophobic interactions. Changes in bilayer properties thus alter membrane protein function, which provides an indirect way for amphiphiles to modulate protein function and a possible mechanism for "off-target" drug effects. We have previously developed an electrophysiological assay for detecting changes in lipid bilayer properties using linear gramicidin channels as probes 3,12. Gramicidin channels are mini-proteins formed by the transbilayer dimerization of two non-conducting subunits. They are sensitive to changes in their membrane environment, which makes them powerful probes for monitoring changes in lipid bilayer properties as sensed by bilayer spanning proteins. We now demonstrate a fluorescence assay for detecting changes in bilayer properties using the same channels as probes. The assay is based on measuring the time-course of fluorescence quenching from fluorophore-loaded large unilamellar vesicles due to the entry of a quencher through the gramicidin channels. We use the fluorescence indicator/quencher pair 8-aminonaphthalene-1,3,6-trisulfonate (ANTS)/Tl+ that has been successfully used in other fluorescence quenching assays 5,13. Tl+ permeates the lipid bilayer slowly 8 but passes readily through conducting gramicidin channels 1,14. The method is scalable and suitable for both mechanistic studies and high-throughput screening of small molecules for bilayer-perturbing, and potential "off-target", effects. We find that results using this method are in good agreement with previous electrophysiological results 12.

We introduce a fast fluorescence-based assay that monitors the rate of fluorescence quenching as a measure of gramicidin channel activity. The gramicidin channels are used as molecular force transducers to monitor changes in lipid bilayer properties as sensed by bilayer spanning proteins.

Many drugs and other small molecules used to modulate biological function are amphiphiles that adsorb at the bilayer/solution interface and thereby alter ...

Biology

Lipid Vesicle-mediated Affinity Chromatography using Magnetic Activated Cell Sorting (LIMACS): a Novel Method to Analyze Protein-lipid Interaction

The analysis of lipid protein interaction is difficult because lipids are embedded in cell membranes and therefore, inaccessible to most purification procedures. As an alternative, lipids can be coated on flat surfaces as used for lipid ELISA and Plasmon resonance spectroscopy. However, surface coating lipids do not form microdomain structures, which may be important for the lipid binding properties. Further, these methods do not allow for the purification of larger amounts of proteins binding to their target lipids.
 To overcome these limitations of testing lipid protein interaction and to purify lipid binding proteins we developed a novel method termed lipid vesicle-mediated affinity chromatography using magnetic-activated cell sorting (LIMACS). In this method, lipid vesicles are prepared with the target lipid and phosphatidylserine as the anchor lipid for Annexin V MACS. Phosphatidylserine is a ubiquitous cell membrane phospholipid that shows high affinity to the protein Annexin V. Using magnetic beads conjugated to Annexin V the phosphatidylserine-containing lipid vesicles will bind to the magnetic beads. When the lipid vesicles are incubated with a cell lysate the protein binding to the target lipid will also be bound to the beads and can be co-purified using MACS. This method can also be used to test if recombinant proteins reconstitute a protein complex binding to the target lipid. 
 We have used this method to show the interaction of atypical PKC (aPKC) with the sphingolipid ceramide and to co-purify prostate apoptosis response 4 (PAR-4), a protein binding to ceramide-associated aPKC. We have also used this method for the reconstitution of a ceramide-associated complex of recombinant aPKC with the cell polarity-related proteins Par6 and Cdc42. Since lipid vesicles can be prepared with a variety of sphingo- or phospholipids, LIMACS offers a versatile test for lipid-protein interaction in a lipid environment that resembles closely that of the cell membrane. Additional lipid protein complexes can be identified using proteomics analysis of lipid binding protein co-purified with the lipid vesicles.

Lipid Vesicle-mediated Affinity Chromatography using Magnetic Activated Cell Sorting LIMACS: a Novel Method to Analyze Protein-lipid Interaction

To test the interaction of a protein with its target lipid we used MACS and Annexin V-conjugated magnetic beads and lipid vesicles synthesized from the target lipid and Annexin V-binding phosphatidylserine. Proteins bound to the target lipid are co-purified and analyzed after elution from the beads.

The analysis of lipid protein interaction is difficult because lipids are embedded in cell membranes and therefore, inaccessible to most purification ...

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

pH Modulation Assay on Supported Lipid Bilayers to Detect Protein-Phosphoinositide Interactions

Source: Shengjuler, D., et al., <a href="https://app.jove.com/v/55869">PIP-on-a-chip: A Label-free Study of Protein-phosphoinositide Interactions.</a> J. Vis. Exp. (2017)
In this video, we demonstrate protein-phosphoinositide interactions in a microfluidic pattern using the pH modulation assay, using pH-sensitive fluorescent dye tagged to phosphatidylethanolamine lipid of a supported lipid bilayer, called a microfluidic platform. The modulations in pH after protein binding to phospholipid cause fluorescence quenching.

A Fluorescence-based Assay of Phospholipid Scramblase Activity

Samenvatting

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