The authors acknowledge the support from Charles H. Hood Foundation Child Health Research Award and generous support from the Department of Molecular Biology at Massachusetts General Hospital.

1. Preparation of lipid mixtures
<ol>
	<li>Prepare a lipid stock solution by dissolving 1,2-dioleoyl-sn-glycero-3phosphocholine (DOPC), 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine (POPE), L-&#945;-phosphatidylinositol (Liver PI), cardiolipin, and 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-[methoxy(polyethylene glycol)-2000] (18:1 PEG2000 PE) into chloroform at the concentration of 25 mg/mL. Dissolve fluorescent dye-conjugated lipid (TexasRed DHPE and Cy5 DOPE) chloroform at a concentration of 1 mg/mL. Store the lipid solution in amber vials with chloroform resistant liner, further sealed with polytetrafluoroethylene tape. The solution can be kept at -20 &#176;C for up to 6 months.</li>
	<li>Make solutions A and B.
	<ol>
		<li>Mix lipid to prepare solution A (final concentration 1 mg/mL) that contains DOPC (52.8 mol%), POPE (20 mol%), Liver PI (7 mol%) and cardiolipin (20 mol%), and 0.2 mol% fluorophore.</li>
		<li>Make solution B (final concentration 1 mg/mL) containing DOPC (47.8 mol%), POPE (20 mol%), liver PI (7 mol%), cardiolipin (20 mol%) and DOPE-PEG2000 (5 mol%), and 0.2 mol% fluorophore.</li>
		<li>Generate the lipid mixture by adding the calculated volume of storage solution into amber vials using a glass syringe. Match the final volume by adding extra chloroform into the vials. 
		NOTE: For FCS (fluorescence correlation spectroscopy), decrease the ratio of dye conjugated lipid to 0.002 mol% and replace the rest by DOPC.</li>
	</ol>
	</li>
</ol>
2. Fabrication of lipid bilayers
<ol>
	<li>Bake microscope cover glass slides at 520 &#176;C for 30 min. After baking, cool down the cover slides to room temperature.</li>
	<li>Weigh approximately 10 g of sodium hydroxide and add to 500 mL of methanol while stirring. Stir for 2 h, continue to add sodium hydroxide in the solution until precipitates start to show. Make sure to wear appropriate PPE during the whole process.</li>
	<li>Clean the glass slides in 10% sodium dodecyl sulfate solution; methanol saturated with sodium hydroxide; and 50 mM hydrochloric acid, sequentially (bath sonication under each condition for 30 min). Clean the glass slide in ultrapure water for 10 min between each condition. 
	NOTE: Though highly recommended to use fresh solutions for glass slide preparation, each solution can be reused up to 5x or within 1 month, whichever comes first. Make sure to stir-mix the solution before each use.</li>
	<li>Store the cleaned cover glass sealed in HCl solution up to 2 weeks to ensure good bilayer quality. If stored in ultrapure water, use the slides within a week.</li>
	<li>Clean the polytetrafluoroethylene trough of the Langmuir-Blodgett dipping system using chloroform and ultrapure water until no wetting is observed on the trough. Spray chloroform on the trough surface, wipe thoroughly with cellulose wipes 3x. Rinse with ultrapure water and remove the water via suctioning. Repeat 3x.</li>
	<li>Cover the surface of the trough with clean ultrapure water.</li>
	<li>Take 2 pieces of surface-treated cover glass from cleaning solution or ultrapure water, and rinse the glass slide with ultrapure water for approximated 30 s.</li>
	<li>Place the cover glass in a back-to-back manner. Use the substrate clamp to hold the glass slides. Immerse the glass slide underneath the water surface by manually clicking &#8220;dipper down&#8221; on the Langmuir control software.</li>
	<li>Zero the film balance, carefully spreading Solution B drop by drop at the air-water interface (Figure 2A). Make sure lipids are only spreading at the air-water interface, with no chloroform and lipid droplets sinking to the bottom of the polytetrafluoroethylene surface.&#160; Failure to ensure this will&#160;create a lipid &#8220;channel&#8221; and prevent monolayer formation.</li>
	<li>Stop adding lipids till film balance readout around ~15-20 mN/m, wait for ~10-15 min. Initiate the barrier controller to alter the surface area by clicking &#8220;start experiments,&#8221; till film balance readout to 37 mN/m. Keep the pressure for ~20-30 min (Figure 2B).</li>
	<li>Raise the cover glass at the speed of 22 mm/min while maintaining the surface tension at 37 mN/m. A lipid monolayer with polymer tethering will be transferred from the air-water interface to the surface of cover glass through the Blodgett dipping process (Figure 2C). This forms the bottom leaflet of the lipid bilayer.</li>
	<li>Clean the air-water interface by suction, rinse the trough with ultrapure water.</li>
	<li>Clean a one-welled glass slide (e.g., Shaefer slide) using chloroform, ethanol, and ultrapure water before use. Set the clean glass slide on the trough with ultrapure water underneath the water layer. Make sure the well is facing up toward the air-water interface and pour fresh ultrapure water until the glass slide is fully covered. Repeat step 2.8.</li>
	<li>Hold the cover glass with lipid monolayer from step 2.4 using a silicon suction cup (make sure monolayer side is away from the suction cup), gently push the lipid monolayer to the air-water interface, hold the cover glass for ~2-3 s at the interface, then push the cover glass against the slide (Figure 2D). Take the slide out with a cover slide. 
	NOTE: The lipid bilayer will be held at the surface of the cover-glass facing the sandwiched area between the two slides (Figure 2E).</li>
	<li>Take the cover glass with the bilayer to an epifluorescence microscope. Image the lipid bilayer. If a homogeneous distribution of lipid dye is observed, photobleach a small area of the bilayer for 30 s, turn off the light source for ~30 s-1 min., then image again to observe recovery. Lipid bilayer will show fluorescence recovery. 
	NOTE: Membranes with defects or bad fluorescence recovery should not be used for further experiments.</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61620/61620fig2v2.jpg" /> 
Figure 2: Steps in making a polymer-tethered lipid bilayer. 
Steps of making lipid bilayers using Langmuir-Blodgett dipping (A-C) and Langmuir-Schaefer transfer (D) techniques. (E) The final &#8220;sandwich&#8221; containing the lipid bilayer. <a href="https://www.jove.com/files/ftp_upload/61620/61620fig2v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
3. Protein reconstitution into the polymer-tethered lipid bilayer
<ol>
	<li>Prepare a crystallization dish containing ultrapure water. Prepare a clean microscope image ring and place underneath the dish.</li>
	<li>Immerse the &#8220;sandwich&#8221; of the Schaefer slide and cover glass that containing lipid bilayer underneath the water, gently separate the Schaefer slide and cover glass, hold the cover glass slide from the bottom, away from the bilayer side, transfer the cover glass into the image ring, close the image ring. 
	NOTE: Make sure that the cover glass with lipid bilayer is always in water, and the ring is well sealed.</li>
	<li>Replace the ultrapure water in the image ring with Bis-Tris NaCl buffer, make sure the lipid bilayer is not exposed to any air bubbles. Add 1.1 x 10-9 M n-Octyl-&#946;-D-Glucopyranoside to the lipid bilayer. Immediately add the mixture of 1.2 x 10-9 M of DDM and 1.3 x 10-12 mol purified l-Opa136 into the image ring. Incubate sample on a benchtop shaker at low speed for 2 h (Figure 3). 
	NOTE: Detergents may vary depending on protein to be reconstituted.</li>
	<li>Distribute 30 mg SM-2 Resin beads into 3 mL of Bis-Tris buffer and shake before applying. Use a plastic pipette to add 5~10 &#181;L of SM-2 Resin beads to image ring, incubate for 10 min, remove resin beads by rinsing. The final volume of the buffer in the image ring is 1.5 mL.</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61620/61620fig3v2.jpg" /> 
Figure 3: Procedure for reconstituting l-Opa1 into a polymer-tethered lipid bilayer. <a href="https://www.jove.com/files/ftp_upload/61620/61620fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
4. Preparation of proteoliposomes
<ol>
	<li>Prepare 1 mg of lipid mixture A in chloroform solution. Evaporate chloroform under nitrogen flow for 20 min and keep under vacuum overnight and form a lipid film.</li>
	<li>Prepare 50 mM calcein containing buffer by dissolving 15.56 g of calcein to 50 mL of 1.5 mol NaOH solution, stir at room temperature until calcein is completely dissolved, added 12.5 mM Bis-Tris and ultrapure water to the final volume of 500 mL. Adjust the pH to 7.5.</li>
	<li>Suspend lipid film in calcein containing buffer, fully hydrate the lipid by heating the suspension at 65 &#730;C for 20 min. 200 nm liposomes are formed through extrusion using a polycarbonate membrane.</li>
	<li>Add 2 &#181;g of l-Opa1 in 0.5 &#181;M DDM to 0.2 mg liposome and incubate at 4 &#730;C for 1.5 h. Remove the surfactant by dialysis using a 3.5 kDa dialysis cassette against 250 ml of 25 mM Bis-Tris, 150 mM NaCl and 50 mM calcein buffer at 4 &#730;C overnight, changing the buffer twice.</li>
	<li>Remove extra calcein using a PD-10 desalting column.</li>
</ol>
5. Imaging and data analysis
<ol>
	<li>Acquire TIRF images using a 100x oil-immersion objective (N.A 1.4). Use 543 nm laser and a 488 nm laser for the analysis of TexasRed-PE labeled liposomes and proteoliposomes encapsulated with calcein. Use&#160;a 633 nm laser for the analysis of Cy5-PE embedded in the planar lipid bilayer.</li>
	<li>Align the TIRF angle using a lipid bilayer to obtain maximum emission. The quality of lipid bilayer after reconstitution is observed using a 100x oil objective at 25 &#176;C. The diffusion coefficient of phospholipid and reconstituted bilayer are determined using FCS with a protocol describe elsewhere37.</li>
	<li>Add 10 &#181;L of 2 mg/mL proteoliposomes to the image ring and set for 10 mins before the image. GTP, GMPPCP, or GDP are added into the reaction ring with 1 mM MgCl2 and 1 mM of nucleotide.</li>
	<li>To determine the influence of s-Opa1 in membrane fusion, titrate s-Opa1 into a proteoliposome/supported bilayer sample containing l-Opa1, and record fusion events.</li>
	<li>Simultaneous imaging of TexasRed-DHPE and calcein is achieved through a beam-splitting system. Both 488 nm and 543 nm lasers are simultaneously applied to the sample as flourescent excitation sources. The emission light is then divided using a 560 nm beam splitter. The split emission light then is filtered by a 510 nm filter with a bandwidth of 42 nm and a 609 nm filter with a bandwidth of 40 nm. The filtered beam is projected to two adjacent areas on camera chip.</li>
	<li>Fluorescent emission is simultaneously recorded through a 609-emission filter with a bandwidth of 40 nm, and a 698-emission filter with a bandwidth of 70 nm. The microscope system is equipped with a CMOS camera maintained at -10 &#176;C.</li>
	<li>Particle identification of the liposomes can be performed using a Gaussian-based particle recognition algorithm. The particle distribution and intensity are analyzed channel-by-channel. A lipid bilayer signal is used as a mask to isolate particles.</li>
</ol>

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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ligand+binding+alters+dimerization+and+sequestering+of+urokinase+receptors+in+raft-mimicking+lipid+mixtures.">Ligand binding alters dimerization and sequestering of urokinase receptors in raft-mimicking lipid mixtures.</a> Biophysical Journal. 107 (9), 2101-2111 (2014).</li><li>Simons, K., Vaz, W. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Model+systems,+lipid+rafts,+and+cell+membranes.">Model systems, lipid rafts, and cell membranes.</a> Annual Review of Biophysics and Biomolecular Structure. 33, 269-295 (2004).</li><li>Kiessling, V., Liang, B., Tamm, L. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reconstituting+SNARE-mediated+membrane+fusion+at+the+single+liposome+level.">Reconstituting SNARE-mediated membrane fusion at the single liposome level.</a> Methods in Cell Biology. 128, 339-363 (2015).</li><li>Siegel, A. P., Kimble-Hill, A., Garg, S., Jordan, R., Naumann, C. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Native+ligands+change+integrin+sequestering+but+not+oligomerization+in+raft-mimicking+lipid+mixtures.">Native ligands change integrin sequestering but not oligomerization in raft-mimicking lipid mixtures.</a> Biophysics Journal. 101 (7), 1642-1650 (2011).</li></ol>

In vitro model-membrane systems can describe complex membrane processes under well-defined conditions. These systems can distinguish minimal components necessary for complex molecular processes to reveal molecular mechanisms6,15,20,38. For membrane proteins, liposomes and planar supported bilayers are common reconstitution systems. In contrast to solid-supported lipid bilayers, the polymer cushion between the substrate and supported membrane in polymer-tethered bilayers allows free mobility of large membrane proteins, and transmembrane-proteins to diffuse and assemble freely34. These features helped us investigate the kinetics of mitochondria inner-membrane fusion36.
We prepared a polymer-tethered lipid bilayer using Langmuir-Blodgett/Langmuir-Schaefer (LB/LS) techniques. This allows us to prepare a bilayer with asymmetric lipid components. Cellular membranes have asymmetric leaflet composition, and the LB/LS approach allows the study of such bilayers. With Schaefer transfer, an entire glass substrate can be covered by a lipid bilayer. It is critical to prepare a clean surface for bilayer preparation. Additionally, it takes practice to perform a Schaefer transfer correctly. Unsuccessful Schaefer transfer can create unwanted defects in a lipid bilayer. In this protocol, the pressure added to the film balance is applicable for a bilayer containing 20% cardiolipin. For bilayers with other components, refer to the surface pressure-area isotherm of the key components. An alternative method is the Langmuir-Blodgett/vesicle fusion (LB/VF) method, where the bottom lipid monolayer is transferred from the air-water interface of a Langmuir trough onto a clean substrate, then liposomes fuse to the top of the supported lipid monolayer and form the final bilayer39. Reconstitution of membrane proteins using the LB/VF method is more straightforward than LB/LS, as reconstitution can be performed through the fusion of proteoliposomes. However, vesicle fusion requires the addition of excess liposomes, which may complicate the study of membrane events dependent on concentration-dependent protein-protein interactions.
The successful reconstitution of transmembrane proteins into both polymer-tethered lipid bilayers and liposomes in a preferred functional orientation is important, yet difficult to enforce. Experimental controls are needed to account for this. For polymer-tethered lipid bilayers, it is also important to maintain the integrity of lipid bilayer during reconstitution. Surfactant concentrations must be kept relatively low to prevent dissolving the lipid bilayer, but high enough to prevent denaturation of the protein of interest37,40. The method described here is ideal for reconstituting membrane proteins for single-molecule studies but is not necessarily scalable for larger-scale studies. Surfactant choice is another important consideration. Frequently, the surfactant used for purification and storage is a good starting point. The maximum concentration of surfactant is usually ~200 times less of the CMC36, in a range where the surfactant maintains protein stability and prevents protein aggregation, while maintaining the integrity of membrane36. Cocktails containing 2 or 3 surfactants may be considered. For reconstitution into liposomes, a low concentration of surfactant is not necessary. However, surfactant concentrations below CMC are preferable to maintain uniform size and morphology distribution for the liposomes. To prevent leakage of content dye, it is necessary to dialyze against a dye-containing buffer.
In contrast to liposome-based fusion assays, the platform we established provides an approach to investigate the kinetics of each step of membrane fusion. This method provides the ability to study transmembrane fusion proteins under near-native conditions. Model membrane platforms can be applied to study membrane protein assembly and oligomerization, membrane &#8220;sculpting&#8221;, and protein-lipid interactions of proteins in subcellular environments, like the mitochondrial inner-membrane. This method also allows exploration of important physiological conditions in the membrane-protein interplay, such as bilayer composition asymmetry. The role of a key mitochondrial lipid, cardiolipin, in the bilayer properties of both liposomes and polymer-supported bilayers remains to be defined. Properties such as the ionic strength, membrane thickness, membrane stiffness, membrane curvature, and membrane elastic-viscosity properties all may influence the ability of proteins to assembly into specific functional states. Future studies creatively applying model membrane systems have potential to uncover new aspects of membrane protein organization and function.

Membrane compartmentalization is a hallmark of eukaryotic cells1 (Figure 1A). Biological membranes are increasingly recognized as more than a two-dimensional solvent, and are considered as an environment playing critical roles in regulating protein function and macromolecular complex assembly2,3. Native lipids are ligands that regulate membrane protein activity3,4. Membrane spatial organization and the ability of membranes to be sculpted into diverse shapes are important physical properties for selecting new functions3,5.
Model membrane platforms are biomimetic systems that can help us understand cellular membrane structure, dynamics, and function6,7,8. Model membranes typically comprise a lipid mixture of well-defined composition, with defined biophysical properties (stiffness, thickness, and elasticity). Coupled to fluorescence imaging, model membrane platforms allow quantitative analysis of membrane structure and function9,10,11. Lipid bilayer reconstitution strategies have been used to study SNARE-mediated membrane fusion9,10, DNA-mediated membrane fusion12, and viral fusion11,13. An advantage of such methods is the potential to obtain kinetic information for intermediate steps preceding an observable reaction event14.
The plasma membrane has been extensively studied using model membranes. Bilayers with lipid phase separation have been developed to study lipid raft structures important in cellular signaling11,15,16. Micropatterned lipid planar bilayers17,18 have been used to investigate the organization of cell receptors. Polymer or gel-supported membranes have been used as biomimetic systems to study the membrane-cytoskeleton organization, membrane protein partitioning during cell signaling, and migration at cell-cell contacts19.
Artificial membrane systems are also being applied to study subcellular organelles20. Organelles feature characteristic morphologies that create distinct sub-environments. The endoplasmic reticulum (ER) network is one example. Upon reconstitution of reticulons into liposomes, tubular membrane structures with properties similar to the cellular ER are formed21. The addition of atlastin, an ER fusion protein, can induce lipid tubules from liposomes to form a network20. This is one example for how proteoliposomes can provide functional insight into organelle morphology and dynamics.
Mitochondrial membrane fusion and fission are essential for the health of the mitochondrial population22,23,24,25. A set of dynamin family GTPases catalyzes mitochondria membrane fusion. Mfn 1/2 catalyzes outer-membrane fusion. Opa1 mediates inner-membrane fusion26 (Figure 1B). Opa1 has two forms: a long form (l-Opa1), transmembrane-anchored to the mitochondrial inner-membrane, and a &#8216;soluble&#8217; short form (s-Opa1), present in the intermembrane space. The ratio of the two&#160;Opa1 forms is regulated by the activity of two proteases, Oma1 and Yme1L27,28,29,30. Important questions in Opa1 regulation include: how the two forms of Opa1, (short and long) mediate membrane fusion and their regulatory interplay28,29,31,32,33.
Here we describe a reconstitution strategy successfully applied to investigate mitochondrial inner-membrane fusion that clarified the roles of l- and s-Opa1 in inner-membrane fusion. We developed a platform mimicking the mitochondrial inner-membrane using a polymer-tethered lipid bilayer and 200 nm unilamellar vesicles. The benefits of a polymer tether beneath the lipid bilayer include the following. First, it preserves the reconstituted transmembrane protein, which would otherwise may be disrupted by the proximity to the glass slide34. Secondly, it serves a thick water layer between the lipid bilayer and glass substrate, which facilitates studies of pore opening9, and thirdly the viscoelastic nature of the PEG polymer allows membrane curvature changes35. We used three-color fluorescence imaging to characterize steps in membrane fusion (Figure 1C-F).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61620/61620fig1v2.jpg" /> 
Figure 1: Monitoring mitochondrial membrane fusion. 
(A) Organelles are cellular membrane compartments. (B) Sequential steps of mitochondrial membrane fusion. Fusion of the outer membrane of mitochondria is catalyzed by Mfn1 and/or Mfn2, while inner-membrane fusion is mediated by Opa1. (C-F) Schematic of the in vitro reconstitution platform to study mitochondrial membrane fusion. The platform includes two parts: a proteoliposome and a polymer-tethered lipid bilayer, both with reconstituted l-Opa1. Fluorescent labels, including two different fluorescent membrane dyes and a content marker, help distinguish steps during membrane fusion. The two membrane markers (Cy5-PE (red) and TexasRed PE (orange) make a FRET pair, which can report on close membrane docking. Diffusion of TexasRed-PE that labels proteoliposome is an indicator of lipid demixing (hemifusion). Content release is monitored through the dequenching of the calcein signal (shown in green). Panels A and B created using Biorender. <a href="https://www.jove.com/files/ftp_upload/61620/61620fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table>
	<tbody>
		<tr>
			<td>1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-(Cyanine 5)</td>
			<td>Avanti polar lipid</td>
			<td>Cat #: 810335C1mg</td>
			<td>membrane fluorescent markers</td>
		</tr>
		<tr>
			<td>1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-[methoxy(polyethylene glycol)-2000] (ammonium salt)</td>
			<td>Avanti Polar lipids</td>
			<td>Cat #: 880130P</td>
			<td>lipid molecules</td>
		</tr>
		<tr>
			<td>1&#39;,3&#39;-bis[1,2-dioleoyl-sn-glycero-3-phospho]-glycerol (sodium salt)</td>
			<td>Avanti Polar lipids</td>
			<td>Cat #: 710335P</td>
			<td>lipid molecules</td>
		</tr>
		<tr>
			<td>18:1 (&#916;9-Cis) PC (DOPC)</td>
			<td>Avanti Polar lipids</td>
			<td>Cat #: 850375P</td>
			<td>lipid molecules</td>
		</tr>
		<tr>
			<td>1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine</td>
			<td>Avanti Polar lipids</td>
			<td>Cat #: 850757P</td>
			<td>lipid molecules</td>
		</tr>
		<tr>
			<td>Alexa Fluor 488 Antibody Labeling Kit</td>
			<td>ThermoFisher Scientific</td>
			<td>A20181</td>
			<td></td>
		</tr>
		<tr>
			<td>Amber vial with PTFE liner</td>
			<td>Fisher scientific</td>
			<td>14-955-332</td>
			<td>sample vials to keep lipid solutions</td>
		</tr>
		<tr>
			<td>Calcein</td>
			<td>Sigma-Aldrich</td>
			<td>Cat #: C0875; PubChem Substance ID: 24892279</td>
			<td>fluorescent dye</td>
		</tr>
		<tr>
			<td>Chloroform</td>
			<td>Fisher scientific</td>
			<td>298-500/ C295-4</td>
			<td>Fisher brand Chloroform is usually quite reliable for lipid works.</td>
		</tr>
		<tr>
			<td>Concavity slide (1 well)</td>
			<td>Electron Microscopy Science</td>
			<td>71878-05</td>
			<td>applied as Schaefer Slide</td>
		</tr>
		<tr>
			<td>FCS analysis tool</td>
			<td>Smith Lab, University of Akron</td>
			<td></td>
			<td>software tool</td>
		</tr>
		<tr>
			<td>Fiji /ImageJ</td>
			<td>Fiji</td>
			<td>SCR_002285</td>
			<td>software tool</td>
		</tr>
		<tr>
			<td>Fisherbrand Cover Glasses: Circles</td>
			<td>Fisher scientific</td>
			<td>12-545-102</td>
			<td>Cover glass for solid supported lipid bilayers, the item is now discontinued as authors prepared the manuscript. An alternative is Fisher brand premium cover glass with catalog number: 12-548-5M</td>
		</tr>
		<tr>
			<td>GTP Disodium salt</td>
			<td>SIGMA-ALDRICH INC</td>
			<td>Cat #: 10106399001</td>
			<td></td>
		</tr>
		<tr>
			<td>Langmuir &#38; Langmuir-Blodgett Trough</td>
			<td>Biolin Scientifc</td>
			<td>KN2002</td>
			<td></td>
		</tr>
		<tr>
			<td>L-&#945;-lysophosphatidylinositol (Liver, Bovine) (sodium salt)</td>
			<td>Avanti Polar lipids</td>
			<td>Cat #: 850091P</td>
			<td>lipid molecules</td>
		</tr>
		<tr>
			<td>Mini Extruder</td>
			<td>Avanti Polar lipids</td>
			<td>610020</td>
			<td></td>
		</tr>
		<tr>
			<td>n-Dodecyl-&#946;-D-Maltopyranoside</td>
			<td>Anatrace</td>
			<td>Cat #: D310 25 GM</td>
			<td>surfactant for reconstitution</td>
		</tr>
		<tr>
			<td>n-Octyl-&#945;-D-Glucopyranoside</td>
			<td>Anatrace</td>
			<td>Cat #: O311HA 25 GM</td>
			<td>surfactant for reconstitution</td>
		</tr>
		<tr>
			<td>PC Membranes 0.2&#956;m</td>
			<td>Avanti Polar Lipids</td>
			<td>610006</td>
			<td></td>
		</tr>
		<tr>
			<td>Rabbit Anti-Opa1 antibody</td>
			<td>NOVUS BIOLOGICALS</td>
			<td>Cat #: NBP2-59770</td>
			<td>antibody for Opa1 C-terminal detection</td>
		</tr>
		<tr>
			<td>Slidebook</td>
			<td>Intelligent imaging</td>
			<td>RRID: SCR_014300</td>
			<td>software tool</td>
		</tr>
		<tr>
			<td>Teflon threaded seal tape</td>
			<td>Fisher Scientific</td>
			<td>NC0636085</td>
			<td>taflon tape for sample storage</td>
		</tr>
		<tr>
			<td>Texas Red 1,2-Dihexadecanoyl-sn-Glycero-3-Phosphoethanolamine, Triethylammonium Salt (Texas Red DHPE)</td>
			<td>ThermoFisher Scientific</td>
			<td>Cat #: T1395MP</td>
			<td>membrane fluorescent markers</td>
		</tr>
	</tbody>
</table>

a model membrane platform for reconstituting mitochondrial membrane dynamics

Mitochondrial dynamics is essential for the organelle&#8217;s diverse functions and cellular responses. The crowded, spatially complex, mitochondrial membrane is a challenging environment to distinguish regulatory factors. Experimental control of protein and lipid components can help answer specific questions of regulation. Yet, quantitative manipulation of these factors is challenging in cellular assays. To investigate the molecular mechanism of mitochondria inner-membrane fusion, we introduced an in vitro reconstitution platform that mimics the lipid environment of the mitochondrial inner-membrane. Here we describe detailed steps for preparing lipid bilayers and reconstituting mitochondrial membrane proteins. The platform allowed analysis of intermediates in mitochondrial inner-membrane fusion, and the kinetics for individual transitions, in a quantitative manner. This protocol describes the fabrication of bilayers with asymmetric lipid composition and describes general considerations for reconstituting transmembrane proteins into a cushioned bilayer. The method may be applied to study other membrane systems.

The reconstituted transmembrane protein freely diffuses and is homogeneously distributed in the membrane.
Example images of a lipid bilayer and its lipid fluidity validated by epifluorescence microscopy is shown in Figure 4. Lipid distribution in bilayer before and after photobleaching is shown in Figure 4A,B. Homogeneity of the lipid bilayer was visualized using an epifluorescence microscope before and after reconstitution (Figure 4D,E). l-Opa1 reconstituted in lipid bilayer was validated by fluorescence correlation spectroscopy (FCS). We use dye conjugated lipids to evaluate the lipid diffusivity of the bilayer. Reconstituted Opa1 was labeled using a fluorescent-tagged anti-Opa1 C-terminal antibody. Bilayer lipid diffusion was measured as 1.46 &#177; 0.12 &#956;m2/s, while the diffusion coefficient of bilayer-reconstituted l-Opa1 was 0.88 &#177; 0.10 &#956;m2/s. Intensity readout from the FCS curves indicated 75% of l-Opa1 is reconstituted into the lipid bilayer (Figure 4G,H). These results suggest that l-Opa1 freely diffuses in the polymer-tethered lipid bilayer with the potential to self-assemble&#160;into functional complexes.
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61620/61620fig04.jpg" /> 
Figure 4: Distribution of lipid and reconstituted protein in the model membrane. 
(A-C) Example images of a lipid bilayer and its lipid fluidity validated by epifluorescence microscopy. (A) Homogeneous lipid distribution in bilayer prior to photobleaching. (B) Snapshot immediately after photobleaching. (C) Bilayer imaged after fluorescence recovery indicates good lipid fluidity of the membrane following reconstitution. (D,E) Representative images of lipid distribution before (D), and after (E) l-Opa1 reconstitution indicate the reconstitution process did not create defects in the bilayer. Representative TIRF image of l-Opa1 labeled with Alexa 488 conjugated antibody (F) showing a homogeneous distribution of Opa1 upon reconstitution. G. Representative raw photon counts of l-Opa1 signal by fluorescent correlation spectroscopy. In the control, no l-Opa1 was reconstituted in the bilayer, while antibody was added and rinsed. The diffusion of l-Opa1 is significantly slower than lipids in the membrane, consistent with successful reconstitution of transmembrane l-Opa1 (H). Scale bar: 10 &#181;m. <a href="https://www.jove.com/files/ftp_upload/61620/61620fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Fluorescence step bleaching indicated an average of 2-3 copies of l-Opa1 were reconstituted in a given liposome (Figure 5A,B). The size distribution of Opa1 reconstituted proteoliposomes was tested after reconstitution using DLS (Figure 5C). The reconstitution of Opa1 in proteoliposomes was also verified using FCS. The diffusion coefficient of free antibody was 164 &#177; 22 &#956;m2/s; diffusion coefficient for liposomes labeled with a lipid dye was 2.22 &#177; 0.33 &#956;m2/s, and the diffusion coefficient for l-Opa1 proteoliposomes bound to a TexasRed labeled anti-His antibody was 2.12 &#177; 0.36 &#956;m2/s.
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/61620/61620fig05.jpg" /> 
Figure 5: Fabrication and characterization of proteoliposomes. 
(A) Steps in fabricating proteoliposomes with encapsulated, quenched calcein. (B) Representative data of fluorescent step-bleaching show an average of 2-3 copies of l-Opa1 embedded in the liposome. (C) Representative size distributions of proteoliposomes (red) without any nucleotide 1 h after GTP incubation (green). <a href="https://www.jove.com/files/ftp_upload/61620/61620fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Detection of membrane tethering, lipid demixing/hemifusion, and pore opening by fluorescent microscopy.
Membrane tethering is monitored by observing the signal of TexasRed on the surface of lipid bilayer using TIRF microscopy (Figure 6A). Membrane lipid demixing (hemifusion) behavior was monitored through TexasRed as the dye diffuses from the liposomes into the lipid bilayer. Calcein dequenching helps distinguish full fusion pore formation from only lipid demixing. This allows comparison between conditions where particles stall at hemifusion (Fig 6B), and particles that proceed to full fusion (Figure 6C).
Membrane tethering is indicated by a stable lipid signal from liposomes. The distance could be evaluated based on the FRET signal between the labels of the two membranes36. Hemifusion signal features no dequenching in the calcein signal (Figure 6B, lower row), but a rapid decay of the TexasRed signal indicates diffusion of the dye into the lipid bilayer (Figure 6B upper row). Full fusion (with pore opening) features both lipid decay and content release (Figure 6C). TexasRed intensity and calcein intensity can be tracked in a time-dependent manner to provide quantitative detail for the kinetics of membrane fusion36.
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/61620/61620fig06.jpg" /> 
Figure 6: Representative results showing particle tethering (A, scale bar 10 &#181;m), hemifusion (B, scale bar 0.5 &#181;m), and fusion (C, scale bar 0.5 &#181;m). 
(A) Proteoliposomes tethered to Opa1-reconstituted lipid bilayer before GTP addition. (B) An example of hemifusion. The upper row in B shows lipid demixing (TexasRed signal, red), lower row in B shows no content release (calcein signal, green) under these conditions. (C) A representative trace of proteoliposome fusing with the lipid bilayer. Content release can be observed from images in the lower row of showing dequenching of calcein (lower row, green). <a href="https://www.jove.com/files/ftp_upload/61620/61620fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about A Model Membrane Platform for Reconstituting Mitochondrial Membrane Dynamics at JoVE.com

A Model Membrane Platform for Reconstituting Mitochondrial Membrane Dynamics

Mitochondrial fusion is an important homeostatic reaction underlying mitochondrial dynamics. Described here is an in vitro reconstitution system to study mitochondrial inner-membrane fusion that can resolve membrane tethering, docking, hemifusion, and pore opening. The versatility of this approach in exploring cell membrane systems is discussed.

Mitochondrial dynamics is essential for the organelle&#8217;s diverse functions and cellular responses. The crowded, spatially complex, mitochondrial ...

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7.4K Views.  Massachusetts General Hospital. Mitochondrial fusion is an important homeostatic reaction underlying mitochondrial dynamics. Described here is an in vitro reconstitution system to study mitochondrial inner-membrane fusion that can resolve membrane tethering, docking, hemifusion, and pore opening. The versatility of this approach in exploring cell membrane systems is discussed.

Video: A Model Membrane Platform for Reconstituting Mitochondrial Membrane Dynamics

Directed Protein Packaging within Outer Membrane Vesicles from Escherichia coli: Design, Production and Purification

An increasing interest in applying synthetic biology techniques to program outer membrane vesicles (OMV) are leading to some very interesting and unique applications for OMV where traditional nanoparticles are proving too difficult to synthesize. To date, all Gram-negative bacteria have been shown to produce OMV demonstrating packaging of a variety of cargo that includes small molecules, peptides, proteins and genetic material. Based on their diverse cargo, OMV are implicated in many biological processes ranging from cell-cell communication to gene transfer and delivery of virulence factors depending upon which bacteria are producing the OMV. Only recently have bacterial OMV become accessible for use across a wide range of applications through the development of techniques to control and direct packaging of recombinant proteins into OMV. This protocol describes a method for the production, purification, and use of enzyme packaged OMV providing for improved overall production of recombinant enzyme, increased vesiculation, and enhanced enzyme stability. Successful utilization of this protocol will result in the creation of a bacterial strain that simultaneously produces a recombinant protein and directs it for OMV encapsulation through creating a synthetic linkage between the recombinant protein and an outer membrane anchor protein. This protocol also details methods for isolating OMV from bacterial cultures as well as proper handling techniques and things to consider when adapting this protocol for use for other unique applications such as: pharmaceutical drug delivery, medical diagnostics, and environmental remediation.

Directed Protein Packaging within Outer Membrane Vesicles from Escherichia coli: Design, Production and Purification

A protocol for the production, purification, and use of enzyme packaged outer membrane vesicles (OMV) providing for enhanced enzyme stability for implementation across diverse applications is presented.

An increasing interest in applying synthetic biology techniques to program outer membrane vesicles (OMV) are leading to some very interesting and unique ...

Bacterial Inner-membrane Display for Screening a Library of Antibody Fragments

Antibodies engineered for intracellular function must not only have affinity for their target antigen, but must also be soluble and correctly folded in the cytoplasm. Commonly used methods for the display and screening of recombinant antibody libraries do not incorporate intracellular protein folding quality control, and, thus, the antigen-binding capability and cytoplasmic folding and solubility of antibodies engineered using these methods often must be engineered separately. Here, we describe a protocol to screen a recombinant library of single-chain variable fragment (scFv) antibodies for antigen-binding and proper cytoplasmic folding simultaneously. The method harnesses the intrinsic intracellular folding quality control mechanism of the Escherichia coli twin-arginine translocation (Tat) pathway to display an scFv library on the E. coli inner membrane. The Tat pathway ensures that only soluble, well-folded proteins are transported out of the cytoplasm and displayed on the inner membrane, thereby eliminating poorly folded scFvs prior to interrogation for antigen-binding. Following removal of the outer membrane, the scFvs displayed on the inner membrane are panned against a target antigen immobilized on magnetic beads to isolate scFvs that bind to the target antigen. An enzyme-linked immunosorbent assay (ELISA)-based secondary screen is used to identify the most promising scFvs for additional characterization. Antigen-binding and cytoplasmic solubility can be improved with subsequent rounds of mutagenesis and screening to engineer antibodies with high affinity and high cytoplasmic solubility for intracellular applications.

We provide a method to simultaneously screen a library of antibody fragments for binding affinity and cytoplasmic solubility by using the Escherichia coli twin-arginine translocation pathway, which has an inherent quality control mechanism for intracellular protein folding, to display the antibody fragments on the inner membrane.

Antibodies engineered for intracellular function must not only have affinity for their target antigen, but must also be soluble and correctly folded in ...

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

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

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

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

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

High Precision FRET at Single-molecule Level for Biomolecule Structure Determination

A protocol on how to perform high-precision interdye distance measurements using F&#246;rster resonance energy transfer (FRET) at the single-molecule level in multiparameter fluorescence detection (MFD) mode is presented here. MFD maximizes the usage of all &#34;dimensions&#34; of fluorescence to reduce photophysical and experimental artifacts and allows for the measurement of interdye distance with an accuracy up to ~1 &#197; in rigid biomolecules. This method was used to identify three conformational states of the ligand-binding domain of the N-methyl-D-aspartate (NMDA) receptor to explain the activation of the receptor upon ligand binding. When comparing the known crystallographic structures with experimental measurements, they agreed within less than 3 &#197; for more dynamic biomolecules. Gathering a set of distance restraints that covers the entire dimensionality of the biomolecules would make it possible to provide a structural model of dynamic biomolecules.

A protocol for high-precision FRET experiments at the single molecule level is presented here. Additionally, this methodology can be used to identify three conformational states in the ligand-binding domain of the N-methyl-D-aspartate (NMDA) receptor. Determining precise distances is the first step towards building structural models based on FRET experiments.

A protocol on how to perform high-precision interdye distance measurements using F&#246;rster resonance energy transfer (FRET) at the single-molecule ...

Brain Membrane Fractionation: An Ex Vivo Approach to Assess Subsynaptic Protein Localization

Assessing the synaptic protein composition and function constitutes an important challenge in neuroscience. However, it is not easy to evaluate neurotransmission that occurs within synapses because it is highly regulated by dynamic protein-protein interactions and phosphorylation events. Accordingly, when any method is used to study synaptic transmission, a major goal is to preserve these transient physiological modifications. Here, we present a brain membrane fractionation protocol that represents a robust procedure to isolate proteins belonging to different synaptic compartments. In other words, the protocol describes a biochemical methodology to carry out protein enrichment from presynaptic, postsynaptic, and extrasynaptic compartments. First, synaptosomes, or synaptic terminals, are obtained from neurons that contain all synaptic compartments by means of a discontinuous sucrose gradient. Of note, the quality of this initial synaptic membrane preparation is critical. Subsequently, the isolation of the different subsynaptic compartments is achieved with light solubilization using mild detergents at differential pH conditions. This allows for separation by gradient and isopycnic centrifugations. Finally, protein enrichment at the different subsynaptic compartments (i.e., pre-, post- and extrasynaptic membrane fractions) is validated by means of immunoblot analysis using well-characterized synaptic protein markers (i.e., SNAP-25, PSD-95, and synaptophysin, respectively), thus enabling a direct assessment of the synaptic distribution of any particular neuronal protein.

Brain Membrane Fractionation: An Ex Vivo Approach to Assess Subsynaptic Protein Localization

Here, we present a brain membrane fractionation protocol that represents a robust procedure to isolate proteins belonging to different synaptic compartments.

Assessing the synaptic protein composition and function constitutes an important challenge in neuroscience. However, it is not easy to evaluate ...

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

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

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

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

Characterization of Membrane Transporters by Heterologous Expression in E. coli and Production of Membrane Vesicles

Several methods have been developed to functionally characterize novel membrane transporters. Polyamines are ubiquitous in all organisms, but polyamine exchangers in plants have not been identified. Here, we outline a method to characterize polyamine antiporters using membrane vesicles generated from the lysis of Escherichia coli cells heterologously expressing a plant antiporter. First, we heterologously expressed AtBAT1 in an E. coli strain deficient in polyamine and arginine exchange transporters. Vesicles were produced using a French press, purified by ultracentrifugation and utilized in a membrane filtration assay of labeled substrates to demonstrate the substrate specificity of the transporter. These assays demonstrated that AtBAT1 is a proton-mediated transporter of arginine, &#947;-aminobutyric acid (GABA), putrescine and spermidine. The mutant strain that was developed for the assay of AtBAT1 may be useful for the functional analysis of other families of plant and animal polyamine exchangers. We also hypothesize that this approach can be used to characterize many other types of antiporters, as long as these proteins can be expressed in the bacterial cell membrane. E. coli is a good system for the characterization of novel transporters, since there are multiple methods that can be employed to mutagenize native transporters.

Characterization of Membrane Transporters by Heterologous Expression in E. coli and Production of Membrane Vesicles

We describe a method for the characterization of proton-driven membrane transporters in membrane vesicle preparations produced by heterologous expression in E. coli and lysis of cells using a French press.

Several methods have been developed to functionally characterize novel membrane transporters. Polyamines are ubiquitous in all organisms, but polyamine ...

Screening for Thermotoga maritima Membrane-Bound Pyrophosphatase Inhibitors

Membrane-bound pyrophosphatases (mPPases) are dimeric enzymes that occur in bacteria, archaea, plants, and protist parasites. These proteins cleave pyrophosphate into two orthophosphate molecules, which is coupled with proton and/or sodium ion pumping across the membrane. Since no homologous proteins occur in animals and humans, mPPases are good candidates in the design of potential drug targets. Here we present a detailed protocol to screen for mPPase inhibitors utilizing the molybdenum blue reaction in a 96 well plate system. We use mPPase from the thermophilic bacterium Thermotoga maritima (TmPPase) as a model enzyme. This protocol is simple and inexpensive, producing a consistent and robust result. It takes only about one hour to complete the activity assay protocol from the start of the assay until the absorbance measurement. Since the blue color produced in this assay is stable for a long period of time, subsequent assay(s) can be performed immediately after the previous batch, and the absorbance can be measured later for all batches at once. The drawback of this protocol is that it is done manually and thus can be exhausting as well as require good skills of pipetting and time keeping. Furthermore, the arsenite-citrate solution used in this assay contains sodium arsenite, which is toxic and should be handled with necessary precautions.

Screening for Thermotoga maritima Membrane-Bound Pyrophosphatase Inhibitors

Here we present a screening method for membrane-bound pyrophosphatase (from Thermotoga maritima) inhibitors based on the molybdenum blue reaction in a 96 well plate format.

Membrane-bound pyrophosphatases (mPPases) are dimeric enzymes that occur in bacteria, archaea, plants, and protist parasites. These proteins cleave ...

PeptiQuick, a One-Step Incorporation of Membrane Proteins into Biotinylated Peptidiscs for Streamlined Protein Binding Assays

Membrane proteins, including transporters, channels, and receptors, constitute nearly one-fourth of the cellular proteome and over half of current drug targets. Yet, a major barrier to their characterization and exploitation in academic or industrial settings is that most biochemical, biophysical, and drug screening strategies require these proteins to be in a water-soluble state. Our laboratory recently developed the peptidisc, a membrane mimetic offering a &#8220;one-size-fits-all&#8221; approach to the problem of membrane protein solubility. We present here a streamlined protocol that combines protein purification and peptidisc reconstitution in a single chromatographic step. This workflow, termed PeptiQuick, allows for bypassing dialysis and incubation with polystyrene beads, thereby greatly reducing exposure to detergent, protein denaturation, and sample loss. When PeptiQuick is performed with biotinylated scaffolds, the preparation can be directly attached to streptavidin-coated surfaces. There is no need to biotinylate or modify the membrane protein target. PeptiQuick is showcased here with the membrane receptor FhuA and antimicrobial ligand colicin M, using biolayer interferometry to determine the precise kinetics of their interaction. It is concluded that PeptiQuick is a convenient way to prepare and analyze membrane protein-ligand interactions within one&#160;day in a detergent-free environment.

We present a method that combines membrane protein purification and reconstitution into peptidiscs in a single chromatographic step. Biotinylated scaffolds are used for direct surface attachment and measurement of protein-ligand interactions via biolayer interferometry.

Membrane proteins, including transporters, channels, and receptors, constitute nearly one-fourth of the cellular proteome and over half of current drug ...

Native Cell Membrane Nanoparticles System for Membrane Protein-Protein Interaction Analysis

Protein-protein interactions in cell membrane systems play crucial roles in a wide range of biological processes- from cell-to-cell interactions to signal transduction; from sensing environmental signals to biological response; from metabolic regulation to developmental control. Accurate structural information of protein-protein interactions is crucial for understanding the molecular mechanisms of membrane protein complexes and for the design of highly specific molecules to modulate these proteins. Many in vivo and in vitro approaches have been developed for the detection and analysis of protein-protein interactions. Among them the structural biology approach is unique in that it can provide direct structural information of protein-protein interactions at the atomic level. However, current membrane protein structural biology is still largely limited to detergent-based methods. The major drawback of detergent-based methods is that they often dissociate or denature membrane protein complexes once their native lipid bilayer environment is removed by detergent molecules. We have been developing a&#160;native cell membrane nanoparticle system for membrane protein structural biology. Here, we demonstrate the use of this system in the analysis of protein-protein interactions on the cell membrane with a case study of the oligomeric state of AcrB.

Presented here is a protocol for the determination of oligomeric state of membrane proteins that utilizes a native cell membrane nanoparticle system in conjunction with electron microscopy.

Protein-protein interactions in cell membrane systems play crucial roles in a wide range of biological processes- from cell-to-cell interactions to signal ...