The authors thank Benoit Sorre, Damien Cuvelier, Pierre Nassoy, Fran&#231;ois Quemeneur, and Gil Toombes for their essential contributions to establish the nanotube method in the group. The P.B. group belongs to the CNRS consortium CellTiss, to the Labex CelTisPhyBio (ANR-11-LABX0038), and to Paris Sciences et Lettres (ANR-10-IDEX-0001-02). F.-C. Tsai was funded by the EMBO Long-Term fellowship (ALTF 1527-2014) and Marie Curie actions (H2020-MSCA-IF-2014, project membrane-ezrin-actin). M.S. is a Junior Fellow of the Simons Society of Fellows.

The authors have nothing to disclose.

The method of pulling tubes from GUVs gives rich information on the membrane-protein system, as it is not only the means to measure the fundamental mechanical properties of the membrane, but it helps to shed light on the coupling between proteins and membrane curvature. As discussed in the Introduction, other techniques exist to measure the effects of membrane-curving proteins, either by incubating the proteins with sub-micron liposomes tethered to a passivated surface18,19 or by observing the spontaneous formation of tubules in micropipette-aspirated GUVs upon the injection of a protein20,21. The method described here provides direct geometric connection to endocytosis and is unique in providing the important physical measurements of the system at the same time, such as protein density, membrane tension, and membrane force.
An important limitation of the method is that it is not trivial to construct and calibrate all the components involved in the assay. However, while a combination of confocal microscopy, optical tweezers, and micromanipulation provides maximal information, some components can be substituted as discussed in the Introduction, depending on the information that needs to be learned about the system. The minimal components required to successfully probe the curvature coupling of a protein, or any other potential curvature-coupled molecule, are a micropipette aspirator and a confocal microscope, with the optical tweezers being dispensable. Importantly, many of the calibrations (such as the calibration for membrane tube radius and for determining protein surface density) are typically only needed to be performed once. Another limitation of the method is that it can measure only one GUV at a time, amounting to two to three GUVs per hour (with practice); however each measurement provides a wealth of data.
For higher throughput measurements or if it is essential to study the interaction of proteins with spherical and not tubular geometries, other assays will need to be explored or developed. Future efforts will be devoted to: (1) designing a higher-throughput system in which multiple tubes (from multiple GUVs) could be pulled and measured at the same time, and (2) integrating super-resolution microscopy so to study the details of the protein scaffold that forms on the nanotubes.

Many cellular processes, such as endocytosis, trafficking, the formation of filopodia, infection, etc., are accompanied by a dramatic change in the shape of cell membranes1,2. In the cell, a number of proteins participate in these processes by binding to the membrane and altering their shape. The most notable examples are members of the Bin/Amphiphysin/Rvs (BAR) protein family, containing a characteristic intrinsically curved BAR domain3,4,5,6,7. Typically, they interact with the membrane by adhering the BAR domain to the surface and, in many cases, also shallowly inserting amphipathic helices into the bilayer. The shape, size, and charge of the BAR domain together with the number of amphipathic helices determines: (1) the direction of membrane curvature (i.e., whether they will induce invaginations or protrusions), and (2) the magnitude of membrane curvature5,8. Of note, here positive curvature is defined as the convex side of the curved membrane, i.e., the bulge toward the interacting particle, and negative otherwise. Moreover, quantitative studies of BAR proteins revealed that their effect on the membrane depends on a set of physical parameters: surface density of proteins, membrane tension, and membrane shape (flat versus tubular versus spherical shape)7. Depending on these parameters BAR proteins can: (1) act as sensors of membrane curvature, (2) bend membranes, or (3) induce membrane scission7.
Due to the sheer number of components involved in membrane reshaping in the cell, studying the quantitative aspects of the phenomena, such as endocytosis, in vivo is extremely challenging. In vitro reconstitution of minimal components mimicking curved membranes in the cell provides means to gain a mechanistic understanding of how membrane-curving proteins operate. This article describes a protocol to reconstitute a membrane nanotube in vitro using micromanipulation, confocal microscopy, and optical tweezers. The approach can be used to study, in a quantitative way, how proteins, lipids, or small molecules interact with curved membranes. Lipid GUVs are used as models of a cell membrane, whose curvature is negligible compared to the size of interacting membrane-curving molecules. They are prepared using the electroformation method9 in which the vesicles are formed by hydrating a lipid film and swelling it into GUVs under an alternating current (AC)10. Most common substrates on which GUVs are grown are either semi-conductive plates coated with indium tin oxide (ITO) or platinum wires (Pt-wires)11. In this work, GUVs are grown on Pt-wires as this method has been shown to work much better than the alternative in making GUVs in the presence of salts in the buffer12. Although the electroformation protocol is described here in sufficient detail to reproduce it, we refer the reader to previous articles in which similar and other methods of making GUVs have been described in detail13,14. In our hands, electroformation on Pt-wires has successfully yielded GUVs from a mix of synthetic lipids or from natural lipid extracts in a buffer containing ~100 mM NaCl. Furthermore, it was also possible to encapsulate proteins inside GUVs during growth. An example electroformation chamber is shown in Figure 1A; it comprises two ~10-cm-long Pt-wires inserted into a holder made from polytetrafluoroethylene (PTFE) that can be sealed on both sides with glass coverslips ~1 - 2 cm apart (Figure 1A).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56086/56086fig1.jpg" /> 
Figure 1: Experimental setup. (A) The GUV electroformation chamber with electrical connectors attached to Pt-wires. (B) Left: the experimental system showing the microscope, the experimental chamber above the objective and two micropipettes (left and right) attached to the micromanipulators and inserted into the experimental chamber for tube pulling and protein injection. Right: a close-up view of the experimental chamber mounted above the objective showing the tips of the aspiration and the injection micropipettes inserted. (C) A syringe equipped with a thin dispenser inserted into a micropipette at its back end. The bottom is a close-up view of the dispenser inside the micropipette with the blue dotted line outlining the micropipette. This system is used to fill the micropipette with casein to passivate the glass surface and also to back fill with mineral oil when needed. (D) A system used to aspirate &#181;L quantities of the protein solution. The needle is connected to a syringe and to tubing which is connected to the injection micropipette. The micropipette tip is carefully immersed into the protein solution and aspirated so to fill the micropipette tip. The micropipette is then back filled with mineral oil using the system shown in panel C. <a href="//cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56086/56086fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
A membrane nanotube, ranging in radius from 7 nm to several hundred nm, can be pulled from a GUV by an external force. This method was initially designed to measure the elastic properties of cell membranes and vesicles, such as the bending rigidity15,16. In most recent works, the method was extended to study the interaction of proteins with curved membranes by microinjecting the proteins near the pulled nanotube7,17. Other methods have been developed for studying membrane-curving proteins. In one method, proteins are incubated with differently sized liposomes tethered to a passivated surface. Confocal microscopy is used to measure the protein binding as a function of liposome diameter, which can indicate curvature-induced sorting18,19. In another method, proteins are injected near a micro-aspirated GUV to measure their ability to spontaneously induce tubules20,21. The method described in this protocol is uniquely suited to study membrane-curving proteins involved in endocytosis, where most proteins typically encounter preformed membrane nanotubes connecting the cargo-containing membrane invagination with the underlying flat plasma membrane. Furthermore, in this method, unlike in the assay with tethered small liposomes, the membrane nanotube is continuously connected to the membrane; therefore, it is in mechanical equilibrium with the GUV, a situation expected in vivo. Hence, fundamental membrane physics applies and we can infer a plethora of mechanical properties from our measurements22,23,24.
For a full implementation of this method, the necessary equipment includes a confocal microscope, optical tweezers, and one or two micropipettes connected to a water tank (Figure 1B). By combining all three, it is possible to simultaneously measure membrane tension, membrane curvature, surface density of proteins, and tube force25. Micropipette aspiration is essential and it is easily constructed by inserting a glass micropipette into a holder connected to a water tank, which, via hydrostatic pressure, controls the aspiration pressure26. The micropipette and the holder are controlled by a micromanipulator and, ideally, in one direction by a piezo-actuator for precision movement. To pull a nanotube, the microaspirated GUV is briefly stuck to a micron-sized bead then pulled away creating a nanotube. In this implementation, the bead is held by optical tweezers, which can be constructed by following a published protocol27. It is possible to dispense of the optical tweezers and pull nanotubes in different ways, although at the cost of accurate force measurements. If it is too challenging to build an optical trap or if force measurements are not essential, such as if one simply wants to check the preference of proteins for curved membranes, a tube can be pulled using a bead aspirated at the tip of a second micropipette28. It is also possible to pull tubes using gravitational force29 or flow30,31. Furthermore, confocal microscopy is not essential either; however, it is preferred so to measure the surface density of proteins. It also allows measuring the nanotube radius from fluorescence intensity of lipids in the tube, thus independently of membrane force and tension. Inferring tube radius from fluorescence is particularly important if the relationship between these quantities deviates from well-established equations due to the presence of membrane-adhered proteins25. Importantly, one cannot dispense of both the optical trap and confocal microscopy, as it will not be possible to measure the tube curvature.
The method as described in this protocol has been used to study the curvature-induced sorting of various peripheral membrane proteins on nanotubes, mostly those from the BAR family25,32,33,34. It was also shown that the conically shaped transmembrane potassium channel KvAP is enriched on curved nanotubes in the same way as BAR proteins35. By optimizing the method to encapsulate proteins inside GUVs, the interaction of proteins with negative curvature has been recently investigated as well36. Furthermore, this method has been used to elucidate the formation of protein scaffolds25,37 and to study the mechanism of membrane scission by either line tension38, protein dynamin39, or by BAR proteins40,41. In addition to proteins, small molecules or ions can also induce curvature. Using this method, calcium ions were shown to induce positive curvature under salt-free conditions42. Interestingly, it has also been shown that lipids can undergo curvature sorting, although only for compositions that are near a demixing point43,44. In sum, the method can be used by researchers interested in investigating how either integral membrane components (e.g., lipids or transmembrane proteins) or peripherally binding molecules (either inside or outside GUVs) interact with cylindrically curved membranes, from mechanical and quantitative points of view. It is also intended for those interested in measuring the mechanical properties of the membrane itself22,23,45.

<table>
	<tbody>
		<tr>
			<td>1,2-Dioleoyl-sn-glycero-3-phosphatidylcholine (DOPC)</td>
			<td>Avanti</td>
			<td>850375</td>
			<td>Example lipid used in data for Figure 3</td>
		</tr>
		<tr>
			<td>1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[biotinyl(polyethyleneglycol)-2000] [DSPE-PEG(2000)-biotin]</td>
			<td>Avanti</td>
			<td>880129</td>
			<td>biotinylated lipid</td>
		</tr>
		<tr>
			<td>BODIPY-TR-C5-ceramide</td>
			<td>Molecular Probes (ThermoFisher)</td>
			<td>D7540</td>
			<td>fluorescent lipid</td>
		</tr>
		<tr>
			<td>BODIPY- FLC5-hexadecanoyl phosphatidylcholine (HPC*)</td>
			<td>Molecular Probes (ThermoFisher)</td>
			<td></td>
			<td>fluorescent lipid for protein density calibration</td>
		</tr>
		<tr>
			<td>egg L-&#945;-phosphatidylcholine (eggPC)</td>
			<td>Avanti</td>
			<td>840051</td>
			<td>used for calibrating the tube radius constant</td>
		</tr>
		<tr>
			<td>&#946;-casein from bovine milk (&#62;99%)</td>
			<td>Sigma Aldrich</td>
			<td>C6905</td>
			<td>used for passivating the micropipette and the experimental chamber</td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Sigma Aldrich</td>
			<td>S7903</td>
			<td></td>
		</tr>
		<tr>
			<td>D(+) glucose</td>
			<td>Sigma Aldrich</td>
			<td>G7021</td>
			<td></td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma Aldrich</td>
			<td>S7653</td>
			<td></td>
		</tr>
		<tr>
			<td>Tris</td>
			<td>Sigma Aldrich</td>
			<td>10708976001</td>
			<td></td>
		</tr>
		<tr>
			<td>osmometer</td>
			<td>Loser</td>
			<td>n/a</td>
			<td></td>
		</tr>
		<tr>
			<td>Pt-wires, 0.5 mm diameter, 99.99% pure</td>
			<td>Goodfellow USA</td>
			<td>PT005139</td>
			<td>used for GUV electroformation</td>
		</tr>
		<tr>
			<td>function generator</td>
			<td></td>
			<td>n/a</td>
			<td>used to create current for GUV electroformation</td>
		</tr>
		<tr>
			<td>putty sealant</td>
			<td>Vitrex (from CML France)</td>
			<td>CRIT 140013</td>
			<td>used to seal the electroformation chamber</td>
		</tr>
		<tr>
			<td>bath sonicator</td>
			<td></td>
			<td>n/a</td>
			<td>useful to clean the electroformation chamber, but not crucial</td>
		</tr>
		<tr>
			<td>Nikon TE2000 inverted microscope, eC1 confocal system (Nikon), with two laser lines (&#955; = 488 nm and 543 nm); optical tweezers induced by a 5 W ytterbium fiber continuous wave laser (&#955; &#62; 1070 nm; IPG GmBH Germany)</td>
			<td></td>
			<td></td>
			<td>an example of a confocal microscopy system equipped with optical tweezers</td>
		</tr>
		<tr>
			<td>micromanipulators</td>
			<td></td>
			<td>n/a</td>
			<td></td>
		</tr>
		<tr>
			<td>borosilicate capillaries (with internal and external radii of 0.78 mm and 1 mm, respectively)</td>
			<td>Harvard apparatus</td>
			<td>30-0036</td>
			<td></td>
		</tr>
		<tr>
			<td>micropipette puller</td>
			<td>Sutter Instrument</td>
			<td>P-2100</td>
			<td></td>
		</tr>
		<tr>
			<td>microforge</td>
			<td>Narishige</td>
			<td>MF-800</td>
			<td></td>
		</tr>
		<tr>
			<td>piezoelectric actuator</td>
			<td>Physik Instrumente</td>
			<td>n/a</td>
			<td></td>
		</tr>
		<tr>
			<td>polystyrene streptavidin coated beads (diameter 3.2 &#181;m)</td>
			<td>Spherotech</td>
			<td>SVP-30-5</td>
			<td></td>
		</tr>
	</tbody>
</table>

pulling membrane nanotubes from giant unilamellar vesicles

The reshaping of the cell membrane is an integral part of many cellular phenomena, such as endocytosis, trafficking, the formation of filopodia, etc. Many different proteins associate with curved membranes because of their ability to sense or induce membrane curvature. Typically, these processes involve a multitude of proteins making them too complex to study quantitatively in the cell. We describe a protocol to reconstitute a curved membrane in vitro, mimicking a curved cellular structure, such as the endocytic neck. A giant unilamellar vesicle (GUV) is used as a model of a cell membrane, whose internal pressure and surface tension are controlled with micropipette aspiration. Applying a point pulling force on the GUV using optical tweezers creates a nanotube of high curvature connected to a flat membrane. This method has traditionally been used to measure the fundamental mechanical properties of lipid membranes, such as bending rigidity. In recent years, it has been expanded to study how proteins interact with membrane curvature and the way they affect the shape and the mechanics of membranes. A system combining micromanipulation, microinjection, optical tweezers, and confocal microscopy allows measurement of membrane curvature, membrane tension, and the surface density of proteins, concurrently. From these measurements, many important mechanical and morphological properties of the protein-membrane system can be inferred. In addition, we lay out a protocol of creating GUVs in the presence of physiological salt concentration, and a method of quantifying the surface density of proteins on the membrane from fluorescence intensities of labeled proteins and lipids.

The tube-pulling experiment can give vital mechanical information about the membrane. In the absence of proteins or other molecules that couple with membrane curvature, the membrane force and tube radius can be related with membrane tension by applying the Canham-Helfrich Hamiltonian equation to a tube pulled from a GUV50,51
<img alt="Equation 3" src="/files/ftp_upload/56086/56086eq3v6.jpg" /> (Equation 1)
where <img alt="Equation 4" src="/files/ftp_upload/56086/56086eq4v3.jpg" /> is the membrane free energy of the tube, K membrane bending rigidity, r and R0 are, respectively, the mean and the spontaneous radii of curvature, &#963; membrane tension, A the area of the tube, L the length of the tube, and f the point force exerted by the tube on the bead. At equilibrium, r is the tube radius, denoted as R, and f becomes the tube-retraction force, F, i.e., the equilibrium force by which the tube is pulling the bead in the optical trap. To compute F and R, we minimize <img alt="Equation 4" src="/files/ftp_upload/56086/56086eq4v3.jpg" /> with respect to L and r, yielding22,23
<img alt="Equation 5" src="/files/ftp_upload/56086/56086eq5v4.jpg" /> (Equation 2)
<img alt="Equation 6" src="/files/ftp_upload/56086/56086eq6v5.jpg" /> (Equation 3)
R0 is neglected if both leaflets of the membrane have the same composition, although it is speculated that other factors, such as charge repulsions can induce some spontaneous curvature42. Force and radius measurements on tubes pulled from 100% DOPC GUVs are in excellent agreement with theoretical predictions in Equation 2 and Equation 3 (Figure 4). The two measurements are independent as the force is measured from the displacement of the bead in the optical trap and the radius from fluorescence intensity, therefore providing two ways of calculating the membrane bending rigidity. Fitting Equation 2 and Equation 3 to the shown data yields: K = (25.1 &#177; 1.4) (mean &#177; SD) kBT and (22.1 &#177; 1.5) kBT, respectively, as previously measured42.
If experimenting with curvature-coupling proteins, it is suggested to use the more sophisticated Hamiltonian&#39;s equation than the one in Equation 1, as Equation 1 only accounts for an effective spontaneous curvature and does not account for the effect of proteins on the membrane&#39;s mechanical properties or possible protein-protein interactions. See, e.g.,25,35,36,52,53. Although advanced theoretical models are essential for an in-depth understanding of a specific protein-membrane system, without resorting to them, it is still possible to obtain very useful information from the tube-pulling method.
For example, to test if proteins sense curvature, the relative intensity of proteins on the tube is measured compared to the GUV, which is considered flat. The interaction with positive curvature is studied by injecting the proteins near the tube with the second micropipette25. The interaction with negative curvature or the behavior of transmembrane proteins is studied by encapsulating or reconstituting proteins during GUV formation35,36. The relative enrichment of BAR and KvAP proteins on membrane tubes over the underlying GUVs indicates that they prefer curved membranes (Figure 5). To be more quantitative, this enrichment may be calculated as a sorting coefficient, S, according to:
<img alt="Equation 7" src="/files/ftp_upload/56086/56086eq7v4.jpg" /> (Equation 4)
where Iprot,tub and Iprot,GUV are the fluorescence intensities of proteins on the tube and on the GUV, respectively, and Ilip,tub and Ilip,GUV are the fluorescence intensities of lipids on the tube and on the GUV, respectively.
Another example is measuring the tube force as the protein binds to the membrane. Figure 5B,C shows that an N-BAR protein endophilin A2 gradually reduces the tube force, decreasing it to zero, indicating that it has formed a three-dimensional structure that keeps the tube stable, which is termed a scaffold. Such structures play important roles in endocytosis, by fixing the tube radius but also in enabling scission7,41. Measurement of S, as well as the mechanical and morphological effects that proteins impose on membranes provides important insights into how they operate in the membrane-bending phenomena in the cell.

<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56086/56086fig4.jpg" /> 
Figure 4: Representative results of mechanical measurements in the absence of proteins. Shown are linearized force and tube radius dependence on membrane tension for a 100% DOPC membrane, from three independent measurements, published previously42. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56086/56086fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/56086/56086fig5.jpg" /> 
Figure 5: Representative results of curvature sensing and scaffold formation by BAR proteins. (A) Curvature sensing by the I-BAR protein IRSp53. The protein senses negative membrane curvature and is bound on the inside of the GUV, with negligible amount of protein present on outer membrane leaflet. Scale bar = 5 &#181;m. Reprinted from36, under the Creative Commons CC-BY license, Copyright 2015, Nature Publishing Group. (B) Forming a protein scaffold by the N-BAR protein endophilin. Proteins are injected near the tube and are therefore bound on the outer membrane leaflet. As described in37, endophilin binds to the tube&#39;s base and forms a scaffold that continuously grows along the tube from the GUV toward the optical trap (OT, white circle). (C) A kymogram of endophilin scaffold growth from the GUV to the bead, with the lipid and protein channels overlaid. The plot shows tube force, F, as a function of time, t. The timescales in the kymogram and the plot are the same (x-axis is shared). In B and C, Scale bar = 2 &#181;m. Reprinted with permission from reference37, Copyright (2016) National Academy of Sciences. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56086/56086fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Pulling Membrane Nanotubes from Giant Unilamellar Vesicles at JoVE.com

Pulling Membrane Nanotubes from Giant Unilamellar Vesicles

Many proteins in the cell sense and induce membrane curvature. We describe a method to pull membrane nanotubes from lipid vesicles to study the interaction of proteins or any curvature-active molecule with curved membranes in vitro.

The reshaping of the cell membrane is an integral part of many cellular phenomena, such as endocytosis, trafficking, the formation of filopodia, etc. Many ...

pulling-membrane-nanotubes-from-giant-unilamellar-vesicles

Universidad San Sebastian

Research

JoVE Journal

Biology

10.9K Views.  PSL Research University, CNRS UMR168. Many proteins in the cell sense and induce membrane curvature. We describe a method to pull membrane nanotubes from lipid vesicles to study the interaction of proteins or any curvature-active molecule with curved membranes in vitro.

Video: Pulling Membrane Nanotubes from Giant Unilamellar Vesicles

Crystallizing Membrane Proteins for Structure Determination using Lipidic Mesophases

A detailed protocol for crystallizing membrane proteins by using lipidic mesophases is described. This method has variously been referred to as the lipidic cubic phase or in meso method. The method has been shown to be quite versatile in that it has been used to solve X-ray crystallographic structures of prokaryotic and eukaryotic proteins, proteins that are monomeric, homo- and hetero-multimeric, chromophore-containing and chromophore-free, and alpha-helical and beta-barrel proteins. Recent successes using in meso crystallization are the human engineered beta2-adrenergic and adenosine A2a G protein-coupled receptors. Protocols are presented for reconstituting the membrane protein into the monoolein-based mesophase, and for setting up crystallizations in the manual mode. Additional steps in the overall process, such as crystal harvesting, are to be addressed in future video articles. The time required to prepare the protein-loaded mesophase and to set up a crystallization plate manually is about one hour.

Herein is described the procedure implemented in the Caffrey Membrane Structural and Functional Biology Group to set up manually crystallization trials of membrane proteins in lipidic mesophases.

A detailed protocol for crystallizing membrane proteins by using lipidic mesophases is described. This method has variously been referred to as the ...

Measuring Peptide Translocation into Large Unilamellar Vesicles

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

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

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

Use of a Robot for High-throughput Crystallization of Membrane Proteins in Lipidic Mesophases

Structure-function studies of membrane proteins greatly benefit from having available high-resolution 3-D structures of the type provided through macromolecular X-ray crystallography (MX). An essential ingredient of MX is a steady supply of ideally diffraction-quality crystals. The in meso or lipidic cubic phase (LCP) method for crystallizing membrane proteins is one of several methods available for crystallizing membrane proteins. It makes use of a bicontinuous mesophase in which to grow crystals. As a method, it has had some spectacular successes of late and has attracted much attention with many research groups now interested in using it. One of the challenges associated with the method is that the hosting mesophase is extremely viscous and sticky, reminiscent of a thick toothpaste. Thus, dispensing it manually in a reproducible manner in small volumes into crystallization wells requires skill, patience and a steady hand. A protocol for doing just that was developed in the Membrane Structural &amp; Functional Biology (MS&amp;FB) Group1-3. JoVE video articles describing the method are available1,4. 
The manual approach for setting up in meso trials has distinct advantages with specialty applications, such as crystal optimization and derivatization. It does however suffer from being a low throughput method. Here, we demonstrate a protocol for performing in meso crystallization trials robotically. A robot offers the advantages of speed, accuracy, precision, miniaturization and being able to work continuously for extended periods under what could be regarded as hostile conditions such as in the dark, in a reducing atmosphere or at low or high temperatures. An in meso robot, when used properly, can greatly improve the productivity of membrane protein structure and function research by facilitating crystallization which is one of the slow steps in the overall structure determination pipeline. 
In this video article, we demonstrate the use of three commercially available robots that can dispense the viscous and sticky mesophase integral to in meso crystallogenesis. The first robot was developed in the MS&amp;FB Group5,6. The other two have recently become available and are included here for completeness.	
An overview of the protocol covered in this article is presented in Figure 1. All manipulations were performed at room temperature (~20 &deg;C) under ambient conditions.

Herein is described a robotic approach to high-throughput crystallization of membrane proteins in lipidic mesophases for use in structure determination using macromolecular X-ray crystallography. Three robots capable of handling the viscous and sticky protein-laden mesophase integral to the method are introduced.

Structure-function studies of membrane proteins greatly benefit from having available high-resolution 3-D structures of the type provided through ...

Examining BCL-2 Family Function with Large Unilamellar Vesicles

The BCL-2 (B cell CLL/Lymphoma) family is comprised of approximately twenty proteins that collaborate to either maintain cell survival or initiate apoptosis1. Following cellular stress (e.g., DNA damage), the pro-apoptotic BCL-2 family effectors BAK (BCL-2 antagonistic killer 1) and/or BAX (BCL-2 associated X protein) become activated and compromise the integrity of the outer mitochondrial membrane (OMM), though the process referred to as mitochondrial outer membrane permeabilization (MOMP)1. After MOMP occurs, pro-apoptotic proteins (e.g., cytochrome c) gain access to the cytoplasm, promote caspase activation, and apoptosis rapidly ensues2.
In order for BAK/BAX to induce MOMP, they require transient interactions with members of another pro-apoptotic subset of the BCL-2 family, the BCL-2 homology domain 3 (BH3)-only proteins, such as BID (BH3-interacting domain agonist)3-6. Anti-apoptotic BCL-2 family proteins (e.g., BCL-2 related gene, long isoform, BCL-xL; myeloid cell leukemia 1, MCL-1) regulate cellular survival by tightly controlling the interactions between BAK/BAX and the BH3-only proteins capable of directly inducing BAK/BAX activation7,8. In addition, anti-apoptotic BCL-2 protein availability is also dictated by sensitizer/de-repressor BH3-only proteins, such as BAD (BCL-2 antagonist of cell death) or PUMA (p53 upregulated modulator of apoptosis), which bind and inhibit anti-apoptotic members7,9. As most of the anti-apoptotic BCL-2 repertoire is localized to the OMM, the cellular decision to maintain survival or induce MOMP is dictated by multiple BCL-2 family interactions at this membrane.
 Large unilamellar vesicles (LUVs) are a biochemical model to explore relationships between BCL-2 family interactions and membrane permeabilization10. LUVs are comprised of defined lipids that are assembled in ratios identified in lipid composition studies from solvent extracted Xenopus mitochondria (46.5% phosphatidylcholine, 28.5% phosphatidylethanoloamine, 9% phosphatidylinositol, 9% phosphatidylserine, and 7% cardiolipin)10. This is a convenient model system to directly explore BCL-2 family function because the protein and lipid components are completely defined and tractable, which is not always the case with primary mitochondria. While cardiolipin is not usually this high throughout the OMM, this model does faithfully mimic the OMM to promote BCL-2 family function. Furthermore, a more recent modification of the above protocol allows for kinetic analyses of protein interactions and real-time measurements of membrane permeabilization, which is based on LUVs containing a polyanionic dye (ANTS: 8-aminonaphthalene-1,3,6-trisulfonic acid) and cationic quencher (DPX: p-xylene-bis-pyridinium bromide)11. As the LUVs permeabilize, ANTS and DPX diffuse apart, and a gain in fluorescence is detected. Here, commonly used recombinant BCL-2 family protein combinations and controls using the LUVs containing ANTS/DPX are described.

Biochemically-defined large unilamellar vesicles (LUVs) are a convenient model system to analyze BCL-2 family interactions with immediate implications in better understanding the mitochondrial pathway of apoptosis. A method to produce LUVs, along with standard BCL-2 family protein combinations and controls to examine LUV permeabilization, are presented.

The BCL-2 (B cell CLL/Lymphoma) family is comprised of approximately twenty proteins that collaborate to either maintain cell survival or initiate ...

Giant Liposome Preparation for Imaging and Patch-Clamp Electrophysiology

The reconstitution of ion channels into chemically defined lipid membranes for electrophysiological recording has been a powerful technique to identify and explore the function of these important proteins. However, classical preparations, such as planar bilayers, limit the manipulations and experiments that can be performed on the reconstituted channel and its membrane environment. The more cell-like structure of giant liposomes permits traditional patch-clamp experiments without sacrificing control of the lipid environment.
Electroformation is an efficient mean to produce giant liposomes &gt;10 &mu;m in diameter which relies on the application of alternating voltage to a thin, ordered lipid film deposited on an electrode surface. However, since the classical protocol calls for the lipids to be deposited from organic solvents, it is not compatible with less robust membrane proteins like ion channels and must be modified. Recently, protocols have been developed to electroform giant liposomes from partially dehydrated small liposomes, which we have adapted to protein-containing liposomes in our laboratory.
We present here the background, equipment, techniques, and pitfalls of electroformation of giant liposomes from small liposome dispersions. We begin with the classic protocol, which should be mastered first before attempting the more challenging protocols that follow. We demonstrate the process of controlled partial dehydration of small liposomes using vapor equilibrium with saturated salt solutions. Finally, we demonstrate the process of electroformation itself. We will describe simple, inexpensive equipment that can be made in-house to produce high-quality liposomes, and describe visual inspection of the preparation at each stage to ensure the best results.

Reconstituting functional membrane proteins into giant liposomes of defined composition is a powerful approach when combined with patch-clamp electrophysiology. However, conventional giant liposome production may be incompatible with protein stability. We describe protocols for producing giant liposomes from pure lipids or small liposomes containing ion channels.

The reconstitution of ion channels into chemically defined lipid membranes for electrophysiological recording has been a powerful technique to identify ...

Reconstitution of a Transmembrane Protein, the Voltage-gated Ion Channel, KvAP, into Giant Unilamellar Vesicles for Microscopy and Patch Clamp Studies

Giant Unilamellar Vesicles (GUVs) are a popular biomimetic system for studying membrane associated phenomena. However, commonly used protocols to grow GUVs must be modified in order to form GUVs containing functional transmembrane proteins. This article describes two dehydration-rehydration methods &#8212; electroformation and gel-assisted swelling &#8212; to form GUVs containing the voltage-gated potassium channel, KvAP. In both methods, a solution of protein-containing small unilamellar vesicles is partially dehydrated to form a stack of membranes, which is then allowed to swell in a rehydration buffer. For the electroformation method, the film is deposited on platinum electrodes so that an AC field can be applied during film rehydration. In contrast, the gel-assisted swelling method uses an agarose gel substrate to enhance film rehydration. Both methods can produce GUVs in low (e.g., 5 mM) and physiological (e.g., 100 mM) salt concentrations. The resulting GUVs are characterized via fluorescence microscopy, and the function of reconstituted channels measured using the inside-out patch-clamp configuration. While swelling in the presence of an alternating electric field (electroformation) gives a high yield of defect-free GUVs, the gel-assisted swelling method produces a more homogeneous protein distribution and requires no special equipment.

The reconstitution of the transmembrane protein, KvAP, into giant unilamellar vesicles (GUVs) is demonstrated for two dehydration-rehydration methods &#8212; electroformation, and gel-assisted swelling. In both methods, small unilamellar vesicles containing the protein are fused together to form GUVs that can then be studied by fluorescence microscopy and patch-clamp electrophysiology.

Giant Unilamellar Vesicles (GUVs) are a popular biomimetic system for studying membrane associated phenomena. However, commonly used protocols to grow ...

Microperfusion Technique to Investigate Regulation of Microvessel Permeability in Rat Mesentery

Experiments to measure the permeability properties of individually perfused microvessels provide a bridge between investigation of molecular and cellular mechanisms regulating vascular permeability in cultured endothelial cell monolayers and the functional exchange properties of whole microvascular beds. A method to cannulate and perfuse venular microvessels of rat mesentery and measure the hydraulic conductivity of the microvessel wall is described. The main equipment needed includes an intravital microscope with a large modified stage that supports micromanipulators to position three different microtools: (1) a beveled glass micropipette to cannulate and perfuse the microvessel; (2) a glass micro-occluder to transiently block perfusion and enable measurement of transvascular water flow movement at a measured hydrostatic pressure, and (3) a blunt glass rod to stabilize the mesenteric tissue at the site of cannulation. The modified Landis micro-occlusion technique uses red cells suspended in the artificial perfusate as markers of transvascular fluid movement, and also enables repeated measurements of these flows as experimental conditions are changed and hydrostatic and colloid osmotic pressure difference across the microvessels are carefully controlled. Measurements of hydraulic conductivity first using a control perfusate, then after re-cannulation of the same microvessel with the test perfusates enable paired comparisons of the microvessel response under these well-controlled conditions. Attempts to extend the method to microvessels in the mesentery of mice with genetic modifications expected to modify vascular permeability were severely limited because of the absence of long straight and unbranched microvessels in the mouse mesentery, but the recent availability of the rats with similar genetic modifications using the CRISPR/Cas9 technology is expected to open new areas of investigation where the methods described herein can be applied.

The modified Landis technique enables paired measurement of the hydraulic conductivity of individual microvessels in the mesentery of normal and genetically modified rats under control and test conditions using microperfusion techniques. It provides a convenient method to evaluate mechanisms that regulate microvessel permeability and transvascular exchange under physiological conditions.

Experiments to measure the permeability properties of individually perfused microvessels provide a bridge between investigation of molecular and cellular ...

Ground State Depletion Super-resolution Imaging in Mammalian Cells

Advances in fluorescent microscopy and cell biology are intimately correlated, with the enhanced ability to visualize cellular events often leading to dramatic leaps in our understanding of how cells function. The development and availability of super-resolution microscopy has considerably extended the limits of optical resolution from ~250-20 nm. Biologists are no longer limited to describing molecular interactions in terms of colocalization within a diffraction limited area, rather it is now possible to visualize the dynamic interactions of individual molecules. Here, we outline a protocol for the visualization and quantification of cellular proteins by ground-state depletion microscopy&#160;for fixed cell imaging. We provide examples from two different membrane proteins, an element of the endoplasmic reticulum translocon, sec61&#946;, and a plasma membrane-localized voltage-gated L-type Ca2+ channel (CaV1.2). Discussed are the specific microscope parameters, fixation methods, photo-switching buffer formulation, and pitfalls and challenges of image processing.

This article outlines a protocol for the detection of one or more plasma and/or intracellular membrane proteins using Ground State Depletion (GSD) super-resolution microscopy in mammalian cells. Here, we discuss the benefits and considerations of using such approaches for the visualization and quantification of cellular proteins.

Advances in fluorescent microscopy and cell biology are intimately correlated, with the enhanced ability to visualize cellular events often leading to ...

Flow Cytometric Analysis of Extracellular Vesicles from Cell-conditioned Media

Flow cytometry (FC) is the method of choice for semi-quantitative measurement of cell-surface antigen markers. Recently, this technique has been used for phenotypic analyses of extracellular vesicles (EV) including exosomes (Exo) in the peripheral blood and other body fluids. The small size of EV mandates the use of dedicated instruments having a detection threshold around 50-100 nm. Alternatively, EV can be bound to latex microbeads that can be detected by FC. Microbeads, conjugated with antibodies that recognize EV-associated markers/Cluster of Differentiation CD63, CD9, and CD81 can be used for EV capture. Exo isolated from CM can be analyzed with or without pre-enrichment by ultracentrifugation. This approach is suitable for EV analyses using conventional FC instruments. Our results demonstrate a linear correlation between Mean Fluorescence Intensity (MFI) values and EV concentration. Disrupting EV through sonication dramatically decreased MFI, indicating that the method does not detect membrane debris. We report an accurate and reliable method for the analysis of EV surface antigens, which can be easily implemented in any laboratory.

The protocol describes a reproducible method designed for use with cell culture supernatants to detect surface epitopes on small extracellular vesicles (EV). It utilizes specific EV immunoprecipitation using beads coupled with antibodies that recognize surface antigen CD9, CD63, and CD81. The method is optimized for downstream flow cytometry analysis.

Flow cytometry (FC) is the method of choice for semi-quantitative measurement of cell-surface antigen markers. Recently, this technique has been used for ...

Scalable Isolation and Purification of Extracellular Vesicles from Escherichia coli and Other Bacteria

Diverse bacterial species secrete ~20-300 nm extracellular vesicles (EVs), comprised of lipids, proteins, nucleic acids, glycans, and other molecules derived from the parental cells. EVs function as intra- and inter-species communication vectors while also contributing to the interaction between bacteria and host organisms in the context of infection and colonization. Given the multitude of functions attributed to EVs in health and disease, there is a growing interest in isolating EVs for in vitro and in vivo studies. It was hypothesized that the separation of EVs based on physical properties, namely size, would facilitate the isolation of vesicles from diverse bacterial cultures. 
The isolation workflow consists of centrifugation, filtration, ultrafiltration, and size-exclusion chromatography (SEC) for the isolation of EVs from bacterial cultures. A pump-driven tangential flow filtration (TFF) step was incorporated to enhance scalability, enabling the isolation of material from liters of starting cell culture. Escherichia coli was used as a model system expressing EV-associated nanoluciferase and non-EV-associated mCherry as reporter proteins. The nanoluciferase was targeted to the EVs by fusing its N-terminus with cytolysin A. Early chromatography fractions containing 20-100 nm EVs with associated cytolysin A - nanoLuc were distinct from the later fractions containing the free proteins. The presence of EV-associated nanoluciferase was confirmed by immunogold labeling and transmission electron microscopy. This EV isolation workflow is applicable to other human gut-associated gram-negative and gram-positive bacterial species. In conclusion, combining centrifugation, filtration, ultrafiltration/TFF, and SEC enables scalable isolation of EVs from diverse bacterial species. Employing a standardized isolation workflow will facilitate comparative studies of microbial EVs across species.

Scalable Isolation and Purification of Extracellular Vesicles from Escherichia coli and Other Bacteria

Bacteria secrete nanometer-sized extracellular vesicles (EVs) carrying bioactive biological molecules. EV research focuses on understanding their biogenesis, role in microbe-microbe and host-microbe interactions and disease, as well as their potential therapeutic applications. A workflow for scalable isolation of EVs from various bacteria is presented to facilitate standardization of EV research.

Diverse bacterial species secrete ~20-300 nm extracellular vesicles (EVs), comprised of lipids, proteins, nucleic acids, glycans, and other molecules ...