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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<li>Ge, Y., Siegel, A. P., Jordan, R., Naumann, C. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Ligand+binding+alters+dimerization+and+sequestering+of+urokinase+receptors+in+raft-mimicking+lipid+mixtures%5BTitle%5D)+AND+%22Biophysical+Journal%22%5BJournal%5D)">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/pubmed?term=((Model+systems%2C+lipid+rafts%2C+and+cell+membranes%5BTitle%5D)+AND+%22Annual+Review+of+Biophysics+and+Biomolecular+Structure%22%5BJournal%5D)">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/pubmed?term=((Reconstituting+SNARE-mediated+membrane+fusion+at+the+single+liposome+level%5BTitle%5D)+AND+%22Methods+in+Cell+Biology%22%5BJournal%5D)">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/pubmed?term=((Native+ligands+change+integrin+sequestering+but+not+oligomerization+in+raft-mimicking+lipid+mixtures%5BTitle%5D)+AND+%22Biophysics+Journal%22%5BJournal%5D)">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&#039;,3&#039;-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 (&amp;Delta;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 &amp;amp; Langmuir-Blodgett Trough</td><td>Biolin Scientifc</td><td>KN2002</td><td></td></tr><tr><td>L-&amp;alpha;-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-&amp;beta;-D-Maltopyranoside</td><td>Anatrace</td><td>Cat #: D310 25 GM</td><td>surfactant for reconstitution</td></tr><tr><td>n-Octyl-&amp;alpha;-D-Glucopyranoside</td><td>Anatrace</td><td>Cat #: O311HA 25 GM</td><td>surfactant for reconstitution</td></tr><tr><td>PC Membranes 0.2&amp;mu;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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7.6K 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

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

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

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

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

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

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

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

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

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

In Vitro Reconstitution of Self-Organizing Protein Patterns on Supported Lipid Bilayers

Many aspects of the fundamental spatiotemporal organization of cells are governed by reaction-diffusion type systems. In vitro reconstitution of such systems allows for detailed studies of their underlying mechanisms which would not be feasible in vivo. Here, we provide a protocol for the in vitro reconstitution of the MinCDE system of Escherichia coli, which positions the cell division septum in the cell middle. The assay is designed to supply only the components necessary for self-organization, namely a membrane, the two proteins MinD and MinE and energy in the form of ATP. We therefore fabricate an open reaction chamber on a coverslip, on which a supported lipid bilayer is formed. The open design of the chamber allows for optimal preparation of the lipid bilayer and controlled manipulation of the bulk content. The two proteins, MinD and MinE, as well as ATP, are then added into the bulk volume above the membrane. Imaging is possible by many optical microscopies, as the design supports confocal, wide-field and TIRF microscopy alike. In a variation of the protocol, the lipid bilayer is formed on a patterned support, on cell-shaped PDMS microstructures, instead of glass. Lowering the bulk solution to the rim of these compartments encloses the reaction in a smaller compartment and provides boundaries that allow mimicking of in vivo oscillatory behavior. Taken together, we describe protocols to reconstitute the MinCDE system both with and without spatial confinement, allowing researchers to precisely control all aspects influencing pattern formation, such as concentration ranges and addition of other factors or proteins, and to systematically increase system complexity in a relatively simple experimental setup.

In Vitro Reconstitution of Self-Organizing Protein Patterns on Supported Lipid Bilayers

We provide a protocol for in vitro self-organization assays of MinD and MinE on a supported lipid bilayer in an open chamber. Additionally, we describe how to enclose the assay in lipid-clad PDMS microcompartments to mimic in vivo conditions by reaction confinement.

Many aspects of the fundamental spatiotemporal organization of cells are governed by reaction-diffusion type systems. In vitro reconstitution of such ...

Reconstitution of Septin Assembly at Membranes to Study Biophysical Properties and Functions

Most cells can sense and change their shape to carry out fundamental cell processes. In many eukaryotes, the septin cytoskeleton is an integral component in coordinating shape changes like cytokinesis, polarized growth, and migration. Septins are filament-forming proteins that assemble to form diverse higher-order structures and, in many cases, are found in different areas of the plasma membrane, most notably in regions of micron-scale positive curvature. Monitoring the process of septin assembly in vivo is hindered by the limitations of light microscopy in cells, as well as the complexity of interactions with both membranes and cytoskeletal elements, making it difficult to quantify septin dynamics in living systems. Fortunately, there has been substantial progress in the past decade in reconstituting the septin cytoskeleton in a cell-free system to dissect the mechanisms controlling septin assembly at high spatial and temporal resolutions. The core steps of septin assembly include septin heterooligomer association and dissociation with the membrane, polymerization into filaments, and the formation of higher-order structures through interactions between filaments. Here, we present three methods to observe septin assembly in different contexts: planar bilayers, spherical supports, and rod supports. These methods can be used to determine the biophysical parameters of septins at different stages of assembly: as single octamers binding the membrane, as filaments, and as assemblies of filaments. We use these parameters paired with measurements of curvature sampling and preferential adsorption to understand how curvature sensing operates at a variety of length and time scales.

Cell-free reconstitution has been a key tool to understand the cytoskeleton assembly, and work in the last decade has established approaches to study septin dynamics in minimal systems. Presented here are three complementary methods to observe septin assembly in different membrane contexts: planar bilayers, spherical supports, and rod supports.

Most cells can sense and change their shape to carry out fundamental cell processes. In many eukaryotes, the septin cytoskeleton is an integral component ...

Single-Molecule Imaging of Lateral Mobility and Ion Channel Activity in Lipid Bilayers using Total Internal Reflection Fluorescence (TIRF) Microscopy

High-resolution imaging techniques have shown that many ion channels are not static, but subject to highly dynamic processes, including the transient association of pore-forming and auxiliary subunits, lateral diffusion, and clustering with other proteins. However, the relationship between lateral diffusion and function is poorly understood. To approach this problem, we describe how lateral mobility and activity of individual channels in supported lipid membranes can be monitored and correlated using total internal reflection fluorescence (TIRF) microscopy. Membranes are fabricated on an ultrathin hydrogel substrate using the droplet interface bilayer (DIB) technique. Compared to other types of model membranes, these membranes have the advantage of being mechanically robust and suitable for highly sensitive analytical techniques. This protocol measures Ca2+ ion flux through single channels by observing the fluorescence emission of a Ca2+-sensitive dye in close proximity to the membrane. In contrast to classical single-molecule tracking approaches, no fluorescent fusion proteins or labels, which can interfere with lateral movement and function in the membrane, are required. Possible changes in ion flux associated with conformational changes of the protein are only due to protein lateral motion in the membrane. Representative results are shown using the mitochondrial protein translocation channel TOM-CC and the bacterial channel OmpF. In contrast to OmpF, the gating of TOM-CC is very sensitive to molecular confinement and the nature of lateral diffusion. Hence, supported droplet-interface bilayers are a powerful tool to characterize the link between lateral diffusion and the function of ion channels.

Single-Molecule Imaging of Lateral Mobility and Ion Channel Activity in Lipid Bilayers using Total Internal Reflection Fluorescence TIRF Microscopy

This protocol describes how to use TIRF microscopy to track individual ion channels and determine their activity in supported lipid membranes, thereby defining the interplay between lateral membrane movement and channel function. It describes how to prepare the membranes, record the data, and analyze the results.

High-resolution imaging techniques have shown that many ion channels are not static, but subject to highly dynamic processes, including the transient ...

Unveiling Mitochondrial Ultrastructure in Neurons Using STED Microscopy

Using Live Cell STED Imaging to Visualize Mitochondrial Inner Membrane Ultrastructure in Neuronal Cell Models

Mitochondria play many essential roles in the cell, including energy production, regulation of Ca2+ homeostasis, lipid biosynthesis, and production of reactive oxygen species (ROS). These mitochondria-mediated processes take on specialized roles in neurons, coordinating aerobic metabolism to meet the high energy demands of these cells, modulating Ca2+ signaling, providing lipids for axon growth and regeneration, and tuning ROS production for neuronal development and function. Mitochondrial dysfunction is therefore a central driver in neurodegenerative diseases. Mitochondrial structure and function are inextricably linked. The morphologically complex inner membrane with structural infolds called cristae harbors many molecular systems that perform the signature processes of the mitochondrion. The architectural features of the inner membrane are ultrastructural and therefore, too small to be visualized by traditional diffraction-limited resolved microscopy. Thus, most insights on mitochondrial ultrastructure have come from electron microscopy on fixed samples. However, emerging technologies in super-resolution fluorescence microscopy now provide resolution down to tens of nanometers, allowing visualization of ultrastructural features in live cells. Super-resolution imaging therefore offers an unprecedented ability to directly image fine details of mitochondrial structure, nanoscale protein distributions, and cristae dynamics, providing fundamental new insights that link mitochondria to human health and disease. This protocol presents the use of stimulated emission depletion (STED) super-resolution microscopy to visualize the mitochondrial ultrastructure of live human neuroblastoma cells and primary rat neurons. This procedure is organized into five sections: (1) growth and differentiation of the SH-SY5Y cell line, (2) isolation, plating, and growth of primary rat hippocampal neurons, (3) procedures for staining cells for live STED imaging, (4) procedures for live cell STED experiments using a STED microscope&#160;for reference, and (5) guidance for segmentation and image processing using examples to measure and quantify morphological features of the inner membrane.

Author Spotlight: Decoding Mitochondrial Aging 

This protocol presents a workflow for the propagation, differentiation, and staining of cultured SH-SY5Y cells and primary rat hippocampal neurons for mitochondrial ultrastructure visualization and analysis using stimulated emission depletion (STED) microscopy.

Mitochondria play many essential roles in the cell, including energy production, regulation of Ca2+ homeostasis, lipid biosynthesis, and production of ...

Neuroscience

MATLAB Optimization for Analyzing Mitochondrial Networks in Neuroscience

Optimized Automated Analysis of Live Neuronal Mitochondria Homeostasis Modulation by Isoform-Specific Retinoic Acid Receptors

The complex mitochondrial network makes it very challenging to segment, follow, and analyze live cells. MATLAB tools allow the analysis of mitochondria in timelapse files, considerably simplifying and speeding up the process of image processing. Nonetheless, existing tools produce a large output volume, requiring individual manual attention, and basic experimental setups have an output of thousands of files, each requiring extensive and time-consuming handling. 
To address these issues, a routine optimization was developed, in both MATLAB code and live-script forms, allowing for swift file analysis and significantly reducing document reading and data processing. With a speed of 100 files/min, the optimization allows an overall rapid analysis. The optimization achieves the results output by averaging frame-specific data for individual mitochondria throughout time frames, analyzing data in a defined manner, consistent with those output from existing tools. Live confocal imaging was performed using the dye tetramethylrhodamine methyl ester, and the routine optimization was validated by treating neuronal cells with retinoic acid receptor (RAR) agonists, whose effects on neuronal mitochondria are established in the literature. The results were consistent with the literature and allowed further characterization of mitochondrial network behavior in response to isoform-specific RAR modulation. 
This new methodology allowed rapid and validated characterization of whole-neuron mitochondria network, but it also allows for differentiation between axon and cell body mitochondria, an essential feature to apply in the neuroscience field. Moreover, this protocol can be applied to experiments using fast-acting treatments, allowing the imaging of the same cells before and after treatments, transcending the field of neuroscience.

Author Spotlight: An Optimized Automated Method for Investigating Retinoic Acid Receptors in Neuronal Mitochondria

The mitochondrial network is extremely complex, making it very challenging to analyze. A novel MATLAB tool analyzes live confocal imaged mitochondria in timelapse images but results in a large output volume requiring individual manual attention. To address this issue, a routine optimization was developed, allowing for speedy file analysis.

The complex mitochondrial network makes it very challenging to segment, follow, and analyze live cells. MATLAB tools allow the analysis of mitochondria in ...

Measurement of Mitochondrial Membrane Potential In Vivo using a Genetically Encoded Voltage Indicator

Mitochondrial membrane potential (MMP, &#916;&#936;m) is critical for mitochondrial functions, including ATP synthesis, ion transport, reactive oxygen species (ROS) generation, and the import of proteins encoded by the nucleus. Existing methods for measuring &#916;&#936;m typically use lipophilic cation dyes, such as Rhodamine 800 and tetramethylrhodamine methyl ester (TMRM), but these are limited by low specificity and are not well-suited for in vivo applications. To address these limitations, we have developed a novel protocol utilizing genetically encoded voltage indicators (GEVIs). Genetically encoded voltage indicators (GEVIs), which generate fluorescent signals in response to membrane potential changes, have demonstrated significant potential for monitoring plasma membrane and neuronal potentials. However, their application to mitochondrial membranes remains unexplored. Here, we developed protein-based mitochondrial-targeted GEVIs capable of detecting &#916;&#936;m fluctuations in cells and the motor cortex of living animals. The mitochondrial potential indicator (MPI)offers a non-invasive approach to study &#916;&#936;m dynamics in real-time, providing a method to investigate mitochondrial function under both normal and pathological conditions.

This protocol describes the application of mitochondria-targeted genetically encoded voltage indicators (GEVIs). These GEVIs offer a significant advantage over traditional mitochondrial membrane potential dyes by enabling specific, in vivo, and real-time monitoring of mitochondrial membrane potential.

Mitochondrial membrane potential (MMP, &#916;&#936;m) is critical for mitochondrial functions, including ATP synthesis, ion transport, reactive oxygen ...

Reconstitution of Membrane-Tethered Minimal Actin Cortices on Supported Lipid Bilayers

The surface of a living cell provides a versatile active platform for numerous cellular processes, which arise from the interplay of the plasma membrane with the underlying actin cortex. In the past decades, reconstituted, minimal systems based on supported lipid bilayers in combination with actin filament networks have proven to be very instrumental in unraveling basic mechanisms and consequences of membrane-tethered actin networks, as well as in studying the functions of individual membrane-associated proteins. Here, we describe how to reconstitute such active composite systems in vitro that consist of fluid supported lipid bilayers coupled via membrane-associated actin-binding proteins to dynamic actin filaments and myosin motors that can be readily observed via total internal reflection fluorescence microscopy. An open-chamber design allows one to assemble the system in a step-by-step manner and to systematically control many parameters such as linker protein concentration, actin concentration, actin filament length, actin/myosin ratio, as well as ATP levels. Finally, we discuss how to control the quality of the system, how to detect and troubleshoot commonly occurring problems, and some limitations of this system in comparison with the living cell surface.

This protocol describes the formation of supported lipid bilayers and the addition of cytoskeletal filaments and motor proteins to study the dynamics of reconstituted, membrane-tethered cytoskeletal networks using fluorescence microscopy.

The surface of a living cell provides a versatile active platform for numerous cellular processes, which arise from the interplay of the plasma membrane ...

Biology

Simulating and Analyzing Lipid Bilayers Using Molecular Dynamics

Realistic Membrane Modeling Using Complex Lipid Mixtures in Simulation Studies

Lipids are structural building blocks of cell membranes; lipid species vary across cell organelles and across organisms. This variety results in different mechanical and structural properties in the membrane that directly impact the molecules and processes that occur at this interface. Lipid composition is dynamic and can serve to modulate cell signaling processes. Computational approaches are increasingly used to predict interactions between biomolecules and provide molecular insights to experimental observables. Molecular dynamics (MD) is a technique based on statistical mechanics that predicts the movement of atoms based on the forces that act on them. MD simulations can be used to characterize the interaction of biomolecules. Here, we briefly introduce the technique, outline practical steps for beginners who are interested in simulating lipid bilayers, demonstrate the protocol with beginner-friendly software, and discuss alternatives, challenges, and important considerations of the process. Particularly, we emphasize the relevance of using complex lipid mixtures to model a cell membrane of interest to capture the appropriate hydrophobic and mechanical environments in simulation. We also discuss some examples where membrane composition and properties modulate the interactions of bilayers with other biomolecules.

Author Spotlight: Advancing Cell Membrane Biophysics - Exploring Interactions and Challenges Through Experimental and Computational Approaches 

Membrane lipid diversity in structure and composition is an important contributor to cellular processes and can be a marker of disease. Molecular dynamics simulations allow us to study membranes and their interactions with biomolecules at atomistic resolution. Here, we provide a protocol to build, run, and analyze complex membrane systems.

Lipids are structural building blocks of cell membranes; lipid species vary across cell organelles and across organisms. This variety results in different ...

A Model Membrane Platform for Reconstituting Mitochondrial Membrane Dynamics

Summary

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Using Live Cell STED Imaging to Visualize Mitochondrial Inner Membrane Ultrastructure in Neuronal Cell Models

Optimized Automated Analysis of Live Neuronal Mitochondria Homeostasis Modulation by Isoform-Specific Retinoic Acid Receptors

Measurement of Mitochondrial Membrane Potential In Vivo using a Genetically Encoded Voltage Indicator

Reconstitution of Membrane-Tethered Minimal Actin Cortices on Supported Lipid Bilayers

Realistic Membrane Modeling Using Complex Lipid Mixtures in Simulation Studies