This work was supported by the AXA research fund and the Warwick-Wellcome Quantitative Biomedicine Programme (Wellcome ISSF, RMRCB0058) for DVK, NCBS-TIFR for AB and ST, and the Wellcome-DBT Margdarshi fellowship (IA/M/15/1/502018) for SM. DVK would also like to thank the Biophysical Society for enabling the virtual networking event &#34;Challenges in understanding multi-component cytoskeletal networks from the molecular to the meso-scale&#34;, which contributed to the creation of this protocol collection.

<ol>
	<li>K&#246;ster, D. V., Mayor, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cortical+actin+and+the+plasma+membrane:+Inextricably+intertwined.">Cortical actin and the plasma membrane: Inextricably intertwined.</a> Current Opinion in Cell Biology. 38, 81-89 (2016).</li><li>Rao, M., Mayor, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Active+organization+of+membrane+constituents+in+living+cells.">Active organization of membrane constituents in living cells.</a> Current Opinion in Cell Biology. 29, 126-132 (2014).</li><li>Honigmann, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+lipid+bound+actin+meshwork+organizes+liquid+phase+separation+in+model+membranes.">A lipid bound actin meshwork organizes liquid phase separation in model membranes.</a> eLife. 3, 01671 (2014).</li><li>Das, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stratification+relieves+constraints+from+steric+hindrance+in+the+generation+of+compact+actomyosin+asters+at+the+membrane+cortex.">Stratification relieves constraints from steric hindrance in the generation of compact actomyosin asters at the membrane cortex.</a> Science Advances. 6 (11), (2020).</li><li>K&#246;ster, D. V., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Actomyosin+dynamics+drive+local+membrane+component+organization+in+an+in+vitro+active+composite+layer.">Actomyosin dynamics drive local membrane component organization in an in vitro active composite layer.</a> Proceedings of the National Academy of Sciences. 113 (12), 1645-1654 (2016).</li><li>Vogel, S. 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S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Myosin+II+filament+dynamics+in+actin+networks+revealed+with+interferometric+scattering+microscopy.">Myosin II filament dynamics in actin networks revealed with interferometric scattering microscopy.</a> Biophysical Journal. 118 (8), 1946-1957 (2020).</li><li>Seara, D. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Entropy+production+rate+is+maximized+in+non-contractile+actomyosin.">Entropy production rate is maximized in non-contractile actomyosin.</a> Nature Communications. 9 (1), 4948 (2018).</li><li>Linsmeier, I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Disordered+actomyosin+networks+are+sufficient+to+produce+cooperative+and+telescopic+contractility.">Disordered actomyosin networks are sufficient to produce cooperative and telescopic contractility.</a> Nature Communications. 7, 12615 (2016).</li><li>Vogel, S. K., Petrasek, Z., Heinemann, F., Schwille, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Myosin+motors+fragment+and+compact+membrane-bound+actin+filaments.">Myosin motors fragment and compact membrane-bound actin filaments.</a> eLife. 2, 00116 (2013).</li><li>Ditlev, J. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+composition-dependent+molecular+clutch+between+T+cell+signaling+condensates+and+actin.">A composition-dependent molecular clutch between T cell signaling condensates and actin.</a> eLife. 8, 42695 (2019).</li><li>Banjade, S., Rosen, M. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phase+transitions+of+multivalent+proteins+can+promote+clustering+of+membrane+receptors.">Phase transitions of multivalent proteins can promote clustering of membrane receptors.</a> eLife. 3, 04123 (2014).</li><li>Heinemann, F., Vogel, S. K., Schwille, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lateral+membrane+diffusion+modulated+by+a+minimal+actin+cortex.">Lateral membrane diffusion modulated by a minimal actin cortex.</a> Biophysical Journal. 104 (7), 1465-1475 (2013).</li><li>Vogel, S. K., Greiss, F., Khmelinskaia, A., Schwille, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Control+of+lipid+domain+organization+by+a+biomimetic+contractile+actomyosin+cortex.">Control of lipid domain organization by a biomimetic contractile actomyosin cortex.</a> eLife. 6, 24350 (2017).</li><li>Block, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Brownian+motion+at+lipid+membranes:+A+comparison+of+hydrodynamic+models+describing+and+experiments+quantifying+diffusion+within+lipid+bilayers.">Brownian motion at lipid membranes: A comparison of hydrodynamic models describing and experiments quantifying diffusion within lipid bilayers.</a> Biomolecules. 8 (2), 30 (2018).</li><li>JoVE, <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=JoVE+Methods+Collection.+In+vitro+reconstitution+of+cytoskeleton+networks+for+biomaterials,+biophysics+and+active+matter+research.">JoVE Methods Collection. In vitro reconstitution of cytoskeleton networks for biomaterials, biophysics and active matter research.</a> Journal of Visualized Experiments. , (2022).</li><li>Pardee, J. D., Spudich, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Purification+of+muscle+actin.">Purification of muscle actin.</a> Methods in Enzymology. 85, 164-181 (1982).</li><li>Spudich, J. A., Watt, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+regulation+of+rabbit+skeletal+muscle+contraction.+I.+Biochemical+studies+of+the+interaction+of+the+tropomyosin-troponin+complex+with+actin+and+the+proteolytic+fragments+of+myosin.">The regulation of rabbit skeletal muscle contraction. I. Biochemical studies of the interaction of the tropomyosin-troponin complex with actin and the proteolytic fragments of myosin.</a> Journal of Biological Chemistry. 246 (15), 4866-4871 (1971).</li><li>Pollard, T. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Myosin+purification+and+characterization.">Myosin purification and characterization.</a> Methods in Cell Biology. 24, 331-371 (1982).</li><li>JoVE Science Education Database. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Separating+protein+with+SDS-PAGE.">Separating protein with SDS-PAGE.</a> Journal of Visualized Experiments. , (2022).</li><li>Bieling, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=WH2+and+proline-rich+domains+of+WASP-family+proteins+collaborate+to+accelerate+actin+filament+elongation.">WH2 and proline-rich domains of WASP-family proteins collaborate to accelerate actin filament elongation.</a> The EMBO Journal. 37 (1), 102-121 (2018).</li><li>Shrivastava, R., K&#246;ster, D., Kalme, S., Mayor, S., Neerathilingam, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tailor-made+ezrin+actin+binding+domain+to+probe+its+interaction+with+actin+in-vitro.">Tailor-made ezrin actin binding domain to probe its interaction with actin in-vitro.</a> PLoS One. 10 (4), 0123428 (2015).</li><li>Nye, J. 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The authors have no conflicts of interest to declare.

This protocol presents a versatile platform and a starting point to design experiments to study the membrane-cortex interface of cells. Critical steps are the preparation of clean glass slides, using fresh lipids for efficient SUV formation (both affecting the quality of SLBs), and the use of freshly recycled myosin II proteins for dynamic actin filament reorganization. When imaging dynamics over a long time, it is very important to incorporate an oxygen scavenger system (e.g., protocatechuic acid and protocatechuate 3 4-dioxygenase5,31).
The open-chamber design allows the sequential addition of components to an existing system without inducing lipid flows. This can be an important advantage over commonly used, closed-chamber approaches or work using encapsulated proteins within liposomes36. Contrary effects such as protein-induced membrane deformation cannot be studied with glass-adsorbed lipid bilayers.
The lipid bilayers can be formed with a wide range of lipid compositions. It begins with adsorption of the lipid vesicles to the hydrophilic glass surface, followed by either spontaneous vesicle rupture due to surface-vesicle and direct vesicle-vesicle interactions or the adsorbed vesicles reaching a critical coverage after which a small fraction of vesicles rupture, forming active edges, which eventually leads to the bilayer formation32. Besides glass, various substrates can be used to form supported lipid bilayers, such as Mica (e.g., for atomic force microscopy), soft substrates (e.g., poly-di-methyl-siloxane), polymer cushions33,34,35, spanning between holes of electron microscopy grids14. Droplet interface bilayers are another interesting method to create stable, free-standing lipid bilayers36. The inclusion of acto-myosin networks into vesicles or emulsions is a very powerful method to study this minimal system in a cell-like geometry37,38, and which is described in detail elsewhere39.

The plasma membrane of a living animal cell constantly interacts with the adjacent actin cytoskeleton, and together they form an active composite material that fulfills a multitude of cellular functions1,2. To study processes at this lipid membrane-actin interface, the reconstitution of cytoskeletal networks on top of supported lipid bilayers (SLBs) has proven to be very helpful. This minimal system approach allows the precise control of cytoskeleton network components and lipid composition. Compared to the free-standing lipid membranes of giant unilamellar vesicles, the planar geometry of SLBs allows efficient use of state-of-the-art microscopy techniques such as super-resolution3,4, total internal reflection fluorescence (TIRF)5,6,7, or interferometric scattering8 to study the spatial organization and dynamics of cytoskeletal networks. TIRF provides the highest contrast for fluorescently labeled components, since the signal of unbound labeled molecules in the solution contributing to the background signal is minimal.
Here, we describe a basic protocol for the formation of actomyosin networks tethered to supported lipid bilayers, which are widely used in the field to study the physics of active, quasi-2D networks9,10,11 and their effect on membrane organisation3,5,12,13,14,15,16 (Figure 1). This approach is not limited to actin-based networks but can also be adapted easily to explore microtubules, intermediate filaments, or composite networks of mixed nature and to study a variety of interactions between lipid membrane proteins and cytoskeletal components using surface-sensitive microscopy methods.
To keep this protocol focused, we have excluded a detailed description of the purification and labeling of actin and myosin proteins or details about how to tune and control the contractility and organization of actomyosin networks. One should refer to other protocols that are published alongside this one in the JoVE Methods Collection, In Vitro Reconstitution of Cytoskeleton Networks for Biomaterials, Biophysics and Active Matter Research17.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63968/63968fig01.jpg" /> 
Figure 1: Schematic of the in vitro actin-membrane active composite system. Created with Biorender. <a href="https://www.jove.com/files/ftp_upload/63968/63968fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1,2 dipalmitoyl-sn-glycero-3-phosphoethanolamine-N- (lissamine rhodamine B sulfonyl)</td>
			<td>Avanti Polar Lipids</td>
			<td>810158</td>
			<td>16:0 RhoPE</td>
		</tr>
		<tr>
			<td>1,2-dioleoyl-sn-glycero-3- [(N-(5-amino-1 carboxypentyl) iminodiacetic acid) succinyl] (nickel salt)</td>
			<td>Avanti Polar Lipids</td>
			<td>790404</td>
			<td>DGS-NTA-Ni2+</td>
		</tr>
		<tr>
			<td>1,2-dioleoyl-sn-glycero-3-phosphocholine</td>
			<td>Avanti Polar Lipids</td>
			<td>850375</td>
			<td>DOPC</td>
		</tr>
		<tr>
			<td>1,2-dipalmitoyl-sn-glycero-3-phosphocholine</td>
			<td>Avanti Polar Lipids</td>
			<td>850355</td>
			<td>DPPC</td>
		</tr>
		<tr>
			<td>Amber glass vials</td>
			<td>ThermoFisher</td>
			<td>B7990-2A</td>
			<td></td>
		</tr>
		<tr>
			<td>ATP disodium salt</td>
			<td>Sigma Aldrich</td>
			<td>A26209</td>
			<td></td>
		</tr>
		<tr>
			<td>Attofluor cell chamber</td>
			<td>ThermoFisher</td>
			<td>A7816</td>
			<td></td>
		</tr>
		<tr>
			<td>Bath sonicator</td>
			<td>GT Sonic</td>
			<td>1860QTS</td>
			<td></td>
		</tr>
		<tr>
			<td>beta-casein</td>
			<td>Sigma Aldrich</td>
			<td>C6905</td>
			<td></td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>ThermoFisher</td>
			<td>12135</td>
			<td></td>
		</tr>
		<tr>
			<td>chloroform</td>
			<td>Sigma Aldrich</td>
			<td>650471</td>
			<td>alternatively from Electron Microscopy Sciences, 50980296</td>
		</tr>
		<tr>
			<td>Cover slips, #1, 25 mm diameter, Gold Seal</td>
			<td>Harvard Apparatus</td>
			<td>64-0705B</td>
			<td></td>
		</tr>
		<tr>
			<td>Cover slips, #1, 40x22 mm, Gold Seal</td>
			<td>ThermoFisher</td>
			<td>48404-031</td>
			<td></td>
		</tr>
		<tr>
			<td>EDTA</td>
			<td>ThermoFisher</td>
			<td>G12635</td>
			<td></td>
		</tr>
		<tr>
			<td>EGTA</td>
			<td>Himedia</td>
			<td>MB130</td>
			<td></td>
		</tr>
		<tr>
			<td>Gas tight glass syringes, with removebla needle, blunt, volumes 10 &#181;L, 100 &#181;L, 500 &#181;L</td>
			<td>Hammilton</td>
			<td>1700 series</td>
			<td></td>
		</tr>
		<tr>
			<td>Hellmanex III</td>
			<td>Hellma Analytics</td>
			<td>Z805939</td>
			<td>cleaning solution</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Himedia</td>
			<td>RM380</td>
			<td></td>
		</tr>
		<tr>
			<td>KaH2PO4</td>
			<td>ThermoFisher</td>
			<td>G13405</td>
			<td></td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>ThermoFisher</td>
			<td>G13305</td>
			<td></td>
		</tr>
		<tr>
			<td>KOH</td>
			<td>ThermoFisher</td>
			<td>G26708</td>
			<td></td>
		</tr>
		<tr>
			<td>Lipid extruder</td>
			<td>Avanti Polar Lipids</td>
			<td>61000-1EA</td>
			<td></td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>ThermoFisher</td>
			<td>G15535</td>
			<td></td>
		</tr>
		<tr>
			<td>Microtip sonicator</td>
			<td>Sonics</td>
			<td>VC750</td>
			<td>3 mm Tip diameter</td>
		</tr>
		<tr>
			<td>Na2CO3</td>
			<td>ThermoFisher</td>
			<td>G15955</td>
			<td></td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Himedia</td>
			<td>GRM853</td>
			<td></td>
		</tr>
		<tr>
			<td>NaH2PO4</td>
			<td>ThermoFisher</td>
			<td>G15825</td>
			<td></td>
		</tr>
		<tr>
			<td>NaOH</td>
			<td>ThermoFisher</td>
			<td>G27815</td>
			<td></td>
		</tr>
		<tr>
			<td>Nikon Ti Eclipse TIRF microscope</td>
			<td>Nikon</td>
			<td></td>
			<td>With a TIRF unit connected through a polarization-conserving optical fibre to an Agilent monolithic laser combiner MLC400 with multiple laser lines with a 100X, 1.45 NA Nikon Oil Objective with two 512 x 512-pixel EMCCD cameras (Photometrics Evolve 512) with a 100X, 1.45 NA Nikon Oil Objective with two 512 x 512-pixel EMCCD cameras (Photometrics Evolve 512)</td>
		</tr>
		<tr>
			<td>NOA88</td>
			<td>Norland Products</td>
			<td>8801</td>
			<td></td>
		</tr>
		<tr>
			<td>PTFE Coated Tweezer Style #2A</td>
			<td>Structure Probe</td>
			<td>0S2AT-XD</td>
			<td></td>
		</tr>
		<tr>
			<td>Refrigerated microcentrifuge</td>
			<td>Eppendorf</td>
			<td>5424R</td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>ThermoFisher</td>
			<td>G15925</td>
			<td></td>
		</tr>
		<tr>
			<td>UV-illuminator</td>
			<td>Novascan</td>
			<td>PSD PRO-UV</td>
			<td>needs vacuum and O2 supply</td>
		</tr>
	</tbody>
</table>

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.

For representation, here a typical postbleach profile from the 1st image after photobleaching (image at t = 0 s in Figure 3A) and its fit to the following function28 (see Figure 6A) is shown:
<img alt="Equation 4" src="/files/ftp_upload/63968/63968eq04.jpg" style="margin: auto;" />
The value of re (23.94 &#181;m) calculated by the fit to this curve is very similar to the re calculated in step 2.8.4. (23.24 &#181;m). Here, K is a bleach depth parameter that can be directly estimated from F0 (described in step 2.8.4.). Similarly, Figure 6B shows the recovery profile and its fit to the following function28:
<img alt="Equation 5" src="/files/ftp_upload/63968/63968eq05.jpg" style="margin: auto;" />
We find the fitted value of the diffusion coefficient to be 1.34 &#181;m2/s, a value that closely agrees with the value of 1.39 &#181;m2/s that is calculated by the formula in step 2.8.4. Here, MF stands for the mobile fraction of the lipid bilayer that represents the fraction of the bleached population that recovers back. The mobility of lipid-anchored molecules depends, of course, on the lipid composition and its physical state (liquid or gel phase). For our experiments using DOPC-based lipid membranes, the mobility should be &#62;1 &#181;m2/s, and the mobile fraction should not be less than 0.9 to indicate a good lipid bilayer. We recommend the use of the manual fitting-free method for a quick test of the quality and mobility of the bilayer. The fitting method can be useful while automating the analysis for many FRAP curves. Further, if one wants to perform a more sophisticated FRAP experiment to systematically characterize diffusion in the system, we recommend the reader to this review from Lor&#233;n et al.30 for more detail on fitting models and potential pitfalls in experimental design.
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/63968/63968fig06.jpg" /> 
Figure 6: Quantifying the diffusion coefficient of lipid bilayers. (A)&#160;Line profile of the first image after photobleaching (t = 0 s in Figure 3A) and its fit to equation 4 to calculate the effective bleach radius. (B) The recovery profile of the bleached region and its fit to equation 5 to calculate the diffusion coefficient and mobile fraction. <a href="https://www.jove.com/files/ftp_upload/63968/63968fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
A typical result of the experiments described above showing the dynamic assembly and organization of an acto-myosin network linked on a supported lipid bilayer imaged by TIRF microscopy is depicted in Figure 7 and Supplementary Video S1.
Figure 7 shows an image montage of the linker protein, F-actin, and myosin-II.
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/63968/63968fig07.jpg" /> 
Figure 7: Contractile actomyosin flows drive local clustering of the membrane-actin linker protein HYE. TIRF snapshots of HYE (YFP-tagged), actin filaments (labeled with Atto-635 maleimide), and myosin II filaments (labeled with Atto-565 maleimide) upon addition of myosin II to an SLB containing HYE and F-actin. Time is indicated on the top: 0 min is immediately before fluorescent myofilaments started appearing in the TIRF field. HYE and F-actin are homogeneously distributed over the lipid bilayer prior to myosin addition (0 min). Myosin activity induces contractile actomyosin flows, which emerge into aster-like structures at the steady state (15 min), driving local clustering of the coupled membrane component (HYE). The lowest row is a merge of actin (yellow) and myosin II (magenta) images showing the organization of actin and myosin at different time points. The images used in making these montages were corrected in Fiji for background signal, non-uniform intensity patterns, and translational movement. Scale bar = 10 &#181;m. For details, see Supplementary Video S1. <a href="https://www.jove.com/files/ftp_upload/63968/63968fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Buffer Name</td>
			<td colspan="2">Composition</td>
		</tr>
		<tr>
			<td>Lipid Rehydration buffer</td>
			<td colspan="2">50 mM HEPES, 150 mM NaCl, 5% sucrose, pH 7.5</td>
		</tr>
		<tr>
			<td>SLB Formation buffer</td>
			<td colspan="2">50 mM HEPES, 150 mM NaCl, pH 5-6</td>
		</tr>
		<tr>
			<td>SLB Storage buffer</td>
			<td colspan="2">50 mM HEPES, 150 mM NaCl, pH 7.2</td>
		</tr>
		<tr>
			<td>Protein Dilution buffer</td>
			<td colspan="2">20 mM HEPES, 100 mM KCl, 1mM TCEP or DTT, pH 7.2</td>
		</tr>
		<tr>
			<td>1X ME or Actin ion-exchange buffer&#160;</td>
			<td colspan="2">50 mM MgCl2, 0.2 mM EGTA, 10 mM HEPES, pH 7.2 (store at 4&#176;C)</td>
		</tr>
		<tr>
			<td>1X KMEH or Actin polymerization buffer&#160;</td>
			<td colspan="2">50 mM KCl, 1 mM MgCl2, 1 mM EGTA, 50 mM HEPES, pH 7.2</td>
		</tr>
		<tr>
			<td>100 mM ATP stock</td>
			<td colspan="2">100 mM ATP disodium salt, 50 mM Tris, 50 mM NaCl, 5 mM MgCl2, 2 mM EGTA, pH 7.5 (store at -20&#176;C)&#160;</td>
		</tr>
		<tr>
			<td>2x Target buffer</td>
			<td colspan="2">2x KMEH, 2 mg/ml BSA, 2mM ATP, 5mM TCEP (stored at 4&#176;C)</td>
		</tr>
		<tr>
			<td>G-buffer</td>
			<td colspan="2">2 mM Tris, 0.1 mM CaCl2, 0.2 mM ATP, 0.5 mM TCEP, 0.04 % NaN3, pH 8 (store at 4&#176;C)</td>
		</tr>
		<tr>
			<td>Myosin II buffer</td>
			<td colspan="2">500 mM KCl, 1 mM EDTA, 10-20 mM Hepes, pH 7.0</td>
		</tr>
		<tr>
			<td>Gel-filtration chromatography buffer</td>
			<td colspan="2">50 mM Tris-HCl, 150-300 mM NaCl, 5 mM TCEP, 0.1% Tween-20, pH 7.5</td>
		</tr>
		<tr>
			<td>Capping protein Storage buffer&#160;</td>
			<td colspan="2">10 mM Tris&#183;Cl, 50 mM NaCl, 1 mM TCEP, pH 7.5, 20% glycerol&#160;</td>
		</tr>
	</tbody>
</table>
Table 1: List of buffer compositions used in this protocol.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Common problems and their trouble&#160; shooting&#160;</td>
			<td>Problem</td>
			<td colspan="4">Cause</td>
			<td colspan="4">Possible solutions</td>
		</tr>
		<tr>
			<td>1</td>
			<td>Lipid bilayer shows no diffusion</td>
			<td colspan="4">The most probable cause for this problem is dirty coverglass that can happen when the cleaning solution is aged or the heating did not take place during bath sonication. These bilayers have a &#8216;vesicular&#8217; appearance because the bursted vesicles stick to the coverglass but do not fuse with each other. Using MLVs older than 6 weeks or SUVs older than 6 days, or adding low amounts of SUVs can also lead to vesicular bilayer formation.</td>
			<td colspan="4">Use fresh cleaning solution. Make sure the heater is turned on and the temperature is between 45-65&#176;C. Use fresh lipid mixes.(Using a fluorescent lipid probe versus a fluorescent protein probe can sometimes manifest differently. E.g., if the bilayer has sub-diffraction defects and surface passivation step is skipped (or does not work), the lipid probe will show a uniform intensity distribution but the fluorescent protein probe may display bright fluorescent spots. )</td>
		</tr>
		<tr>
			<td>2</td>
			<td>Lipid bilayer has bright patches</td>
			<td colspan="4">Long incubation of SUVs for bilayer formation can create a lipid bilayer that is overall diffusing but with occasional bright patches. These patches can be multi-layered bilayers that can attract large amounts of fluorescent probe.</td>
			<td colspan="4">15-20 min incubation with SUVs is enough. Make sure the probe is not aggregating: a quick hard spin of the linker protein (300 x g for 15 min at 4 &#176;C) can remove the aggregates</td>
		</tr>
		<tr>
			<td>3</td>
			<td>Lipid bilayer has dark holes</td>
			<td colspan="4">This happens when the bilayer is made from old SUVs and imaged for prolonged hours (&#62; 4 hours post formation), or the pH of the solution changes drastically due to prolonged imaging (e.g., in the high ATP state and in the presence of certain oxygen scavengers), or when the surface is over-passivated with beta-Casein (adding too much beta-Casein for more than 10-15 min and or not washing it out).</td>
			<td colspan="4">Use fresh lipids. Reduce the imaging frame rate or the effective laser illumination time. Use buffers with higher buffering capacity.</td>
		</tr>
		<tr>
			<td>4</td>
			<td>Lipid bilayer shows slow diffusion&#160;</td>
			<td colspan="4">Lipid bilayers with high percentage of cholesterol, long saturated lipids or charged lipids diffuse slower.</td>
			<td colspan="4">In such cases, prepare your sample at a high temperature. One can also use a simple, tested lipid composition as a control along with complex and untested lipid compositions. Make sure the glass is clean.</td>
		</tr>
		<tr>
			<td>5</td>
			<td>Actin does not polymerize</td>
			<td colspan="4">&#160;Target buffer is old, G-actin stock is too old, old and new G-actin were co-polymerized.&#160;</td>
			<td colspan="4">Make sure the Ca2+ is replaced by Mg2+ before polymerization (using ME buffer). Use fresh ATP-Mg2+ stock. Use freshly recycled G-actin. Make sure the concentration of F-actin (in terms of G-actin) added to the bilayer is higher than 0.2 &#181;M. For lower concentrations, use phalloidin stabilized F-actin.</td>
		</tr>
		<tr>
			<td>6</td>
			<td>Actin does not bind to the bilayer</td>
			<td colspan="4">Membrane-actin linker is not added or added at very low concentration&#8212;this can be inferred form the fluorescence of the linker protein. If the fluorescence is decent, the membrane-actin linker has lost actin-binding capacity. Also, if the linker protein is non-specifically bound to the glass surface (when the bilayer is bad), it might not recruit actin filaments.</td>
			<td colspan="4">Make sure the bilayer is diffusing. Use fresh linker protein</td>
		</tr>
		<tr>
			<td>7</td>
			<td>Fluorescent F-actin signal is weak</td>
			<td colspan="4">The ratio of labelled to dark actin is too low. Either the labelled actin or unlabelled actin is too old and they are not copolymerizing with each other.</td>
			<td colspan="4">Recycle actin again, and retry the ploymerization with freshly recycled actin. Photodamage can destroy or depolymerize F-actin; if possible, use red or far-red dyes for actin (and myosin).&#160;</td>
		</tr>
		<tr>
			<td>8</td>
			<td>Myosin does not show contractility</td>
			<td colspan="4">it may be observed that after adding ATP to the myosin-infused system, there is no contractility of the acto-myosin.&#160;</td>
			<td colspan="4">Check if myosin concentration or purity level is good. Use freshly recycled myosin (use within 6 weeks after recycling). Adding fresh ATP to the myosin mix can help. De-gassing buffers and using oxygen scavengers etc. can reduce photodamaging of the motors.&#160; Further information might be found in the protocols by Plastino et al. or Stam et al. of the same methods collection</td>
		</tr>
		<tr>
			<td>9</td>
			<td>Coverglass is not hydrophilic</td>
			<td colspan="4">Coverglass is not cleaned properly.</td>
			<td colspan="4">Clean hydrophilllic coverglass is&#160; crucial for the lipid bilayer formation. A useful, visual readout of the hydrophilicity of the coverglass after the cleaning protocol is to observe the wetting of the glass by water. Add a small volume of water to a flat-lying coverslip. The water will remain in the shape of a round droplet if the coverslip is not cleaned properly. However, the same volume of water will spread out and form a thin layer, on a treated hydrophilic coverglass. This wetting behaviour of the water on the cover glass surface can be used to ascertain if the cleaning steps with the cleaning solution/NaOH have worked&#160;.&#160;&#160;&#160;</td>
		</tr>
	</tbody>
</table>
Table 2: Troubleshooting guide summarizing common problems and corresponding solutions. 
Supplementary Video S1: Contractile actomyosin flows drive local clustering of the membrane-actin linker protein HYE. TIRF timelapse of HYE (YFP-tagged), actin filaments (labeled with Atto-635 maleimide), and myosin II filaments (labeled with Atto-565 maleimide) upon addition of myosin II to an SLB containing HYE and F-actin. Time is indicated on the top: 0 min is immediately before fluorescent myofilaments started appearing in the TIRF field. Scale bar = 10 &#181;m. <a href="https://www.jove.com/files/ftp_upload/63968/S1Movie_Act_myo_Hye_merge_timestamp_15fps.zip" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Reconstitution of Membrane-Tethered Minimal Actin Cortices on Supported Lipid Bilayers at JoVE.com

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

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 ...

reconstitution-membrane-tethered-minimal-actin-cortices-on-supported

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2.0K Views.  University of Warwick. 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.

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

Monitoring Actin Disassembly with Time-lapse Microscopy

Okay, so what I'd like to do today is to show you how to construct a flow chamber, a fusion chamber for imaging on an upright microscope. What then I then do is I create two spaces using film. So what I do is I take the para film, I cut it into small strips.Next I'm going to, this will basically form the channels for the, the walls, and I dry them with a tilted air. I put the cover slip on top of the para, And here's the finished. Then I heat it on a heat block to melt the param and basically create a seal.So what this is, is the, the cover glass here, and then on top of the cover glass is two layers of phim as you see right here. And then on top of this, again, there's a, it's a 22 by two two mil meter cover slip. This one right here, which I pressed against the param to fall the seal.Okay, so the liquid goes in in one direction here and I can suck it out with a filter paper here in this direction. And I'll show you how to do that in a bit. So what I like to do is I like to put it in a humidified chamber, which is very simple.Take two kind of elevated strips and have a filter of paper underneath. And to keep it noisy as I, I put the cover, cover slip on the flow chamber on. Now the way you introduce a solution to the chamber is that you just put the pipet tip at the entrance of the chamber and you just split.Now what I do is I incubate the chamber with the lid closed for 10 minutes. What I usually do to suck the liquid out from the other side is to use it, get a filter paper and cut it into thin strips like this. And I take one of these thin strips and I simultaneously apply the solution into the entrance while putting the filter paper at the exit to suck the exchange, the solution.So you'll see the liquid flowing through the solution. And then I wait for five minutes for the mid close. Now what I'm ready to do is to actually put that the actin it and the actin, I actually have to polymerize, I polymerize inside the flow chamber and it's just the same procedures before.So because like this, and then now that the action is polymerized, I wash out all the un excess un polymerized action using a large volume of just buffer. So here I have about a hundred microliters and it's a few reaction chamber volumes worth of buffer. Just, you know, when the filter paper becomes saturated, I have to change the direction.Okay?And so I'm ready to do imaging with this. So perfusion chamber with actin attached to it. Now I take the slide, mount it under the microscope, and since I'm imaging with an oil objective, I'm gonna put some immersion oil on the top surface of the cover, grass touch, such so assess solution or protein or anything on acting inside.So what I do is I can actually exchange the solution side while I'm doing the time lapse imaging. As such.

Monitoring Actin Demontaż z poklatkowy Mikroskopia

Actin Co-Sedimentation Assay; for the Analysis of Protein Binding to F-Actin

The actin cytoskeleton within the cell is a network of actin filaments that allows the movement of cells and cellular processes, and that generates tension and helps maintains cellular shape.  Although the actin cytoskeleton is a rigid structure, it is a dynamic structure that is constantly remodeling.  A number of proteins can bind to the actin cytoskeleton.  The binding of a particular protein to F-actin is often desired to support cell biological observations or to further understand dynamic processes due to remodeling of the actin cytoskeleton. The actin co-sedimentation assay is an in vitro assay routinely used to analyze the binding of specific proteins or protein domains with F-actin.  The basic principles of the assay involve an incubation of the protein of interest (full length or domain of) with F-actin, ultracentrifugation step to pellet F-actin and analysis of the protein co-sedimenting with F-actin.  Actin co-sedimentation assays can be designed accordingly to measure actin binding affinities and in competition assays.

Proteins bind to filamentous actin (F-actin) through distinct actin binding modules. In this video we demonstrate the procedure of actin co-sedimentation, which is an in vitro assay routinely used to analyze proteins or specific domains that bind F-actin.

The actin cytoskeleton within the cell is a network of actin filaments that allows the movement of cells and cellular processes, and that generates ...

Selective Labelling of Cell-surface Proteins using CyDye DIGE Fluor Minimal Dyes

Surface  proteins are central to the cell's ability to react to its environment and to interact with neighboring cells. They are known to be inducers of almost all intracellular signaling. Moreover, they play an important role in environmental adaptation and drug treatment, and are often involved in disease pathogenesis and pathology (1). Protein-protein interactions are intrinsic to signaling pathways, and to gain more insight in these complex biological processes, sensitive and reliable methods are needed for studying cell surface proteins. Two-dimensional (2-D) electrophoresis is used extensively for detection of biomarkers and other targets in complex protein samples to study differential changes. Cell surface proteins, partly due to their low abundance (1 2% of cellular proteins), are difficult to detect in a 2-D gel without fractionation or some other type of enrichment. They are also often poorly represented in 2-D gels due to their hydrophobic nature and high molecular weight (2). In this study, we present a new protocol for intact cells using CyDye  DIGE Fluor minimal dyes for specific labeling and detection of this important group of proteins. The results showed specific labeling of a large number of cell surface proteins with minimal labeling of intracellular proteins. This protocol is rapid, simple to use, and all three CyDye DIGE Fluor minimal dyes (Cy 2, Cy 3 and Cy 5) can be used to label cell-surface proteins. These features allow for multiplexing using the 2-D Fluorescence Difference Gel Electrophoresis (2-D DIGE) with Ettan  DIGE technology and analysis of protein expression changes using DeCyder  2-D Differential Analysis Software. The level of cell-surface proteins was followed during serum starvation of CHO cells for various lengths of time (see Table 1). Small changes in abundance were detected with high accuracy, and results are supported by defined statistical methods.

A simple and specific method was demonstrated for fluorescent labeling and enhanced detection of cell surface proteins without a fractionation step. Differential abundance in cell surface proteins was analyzed using two-dimensional (2-D) electrophoresis and Ettan&#x2122; DIGE technology.

Surface  proteins are central to the cell's ability to react to its environment and to interact with neighboring cells. They are known to be inducers of ...

Supported Planar Bilayers for the Formation of Study of Immunological Synapses and Kinapse

Supported planar bilayers are powerful tools that can be used to model the molecular interactions in an immunological synapse.   To mimic the interactions between lymphocytes and antigen presenting cells, we use Ni2+-chelating lipids to anchor recombinant cell adhesion and MHC proteins to the upper leaflet of a bilayer with poly-histidine tags. Planar bilayers are generated by preparing lipid, treating the glass surfaces where the bilayer will form, and then forming the bilayer in a specialized chamber containing a flow-cell where the lymphocytes will be added.  Then, bilayers are charged with Ni and his-tagged recombinant proteins are added.  Finally,  lymphocytes are injected into the flow cell and TIRF microscopy can be used to image synapse formation and the mechanisms that control T cell locomotion, sites of receptor sorting,  and sites of receptor degradation.

Supported planar bilayers are powerful tools that can be used to model the molecular interactions in an immunological synapse. Here, we show methods for anchoring cell adhesion proteins known to modulate synapse formation to the upper leaflet of the lipid bilyer and visualize synapse formation using TIRF microscopy.

Supported planar bilayers are powerful tools that can be used to model the molecular interactions in an immunological synapse.   To mimic the interactions ...

Preparation of Artificial Bilayers for Electrophysiology Experiments

Planar lipid bilayers, also called artificial lipid bilayers, allow you to study ion-conducting channels in a well-defined environment. These bilayers can be used for many different studies, such as the characterization of membrane-active peptides, the reconstitution of ion channels or investigations on how changes in lipid bilayer properties alter the function of bilayer-spanning channels.  Here, we show how to form a planar bilayer and how to isolate small patches from the bilayer, and in a second video  will also demonstrate a procedure for using gramicidin channels to determine changes in lipid bilayer elastic properties.   We also demonstrate the individual steps needed to prepare the bilayer chamber, the electrodes and how to test that the bilayer is suitable for single-channel measurements.

Planar lipid bilayers, also called artificial lipid bilayers, allow you to study ion-conducting channels in a well-defined environment. Here, we demonstrate the individual steps needed to prepare the bilayer chamber, the electrodes and how to test that the bilayer is suitable for single-channel measurements.

Planar lipid bilayers, also called artificial lipid bilayers, allow you to study ion-conducting channels in a well-defined environment. These bilayers can ...

Study of the Actin Cytoskeleton in Live Endothelial Cells Expressing GFP-Actin

The microvascular endothelium plays an important role as a selectively permeable barrier to fluids and solutes. The adhesive junctions between endothelial cells regulate permeability of the endothelium, and many studies have indicated the important contribution of the actin cytoskeleton to determining junctional integrity1-5. A cortical actin belt is thought to be important for the maintenance of stable junctions1, 2, 4, 5. In contrast, actin stress fibers are thought to generate centripetal tension within endothelial cells that weakens junctions2-5. Much of this theory has been based on studies in which endothelial cells are treated with inflammatory mediators known to increase endothelial permeability, and then fixing the cells and labeling F-actin for microscopic observation. However, these studies provide a very limited understanding of the role of the actin cytoskeleton because images of fixed cells provide only snapshots in time with no information about the dynamics of actin structures5. 
Live-cell imaging allows incorporation of the dynamic nature of the actin cytoskeleton into the studies of the mechanisms determining endothelial barrier integrity. A major advantage of this method is that the impact of various inflammatory stimuli on actin structures in endothelial cells can be assessed in the same set of living cells before and after treatment, removing potential bias that may occur when observing fixed specimens. Human umbilical vein endothelial cells (HUVEC) are transfected with a GFP-&beta;-actin plasmid and grown to confluence on glass coverslips. Time-lapse images of GFP-actin in confluent HUVEC are captured before and after the addition of inflammatory mediators that elicit time-dependent changes in endothelial barrier integrity. These studies enable visual observation of the fluid sequence of changes in the actin cytoskeleton that contribute to endothelial barrier disruption and restoration. 
Our results consistently show local, actin-rich lamellipodia formation and turnover in endothelial cells. The formation and movement of actin stress fibers can also be observed. An analysis of the frequency of formation and turnover of the local lamellipodia, before and after treatment with inflammatory stimuli can be documented by kymograph analyses. These studies provide important information on the dynamic nature of the actin cytoskeleton in endothelial cells that can used to discover previously unidentified molecular mechanisms important for the maintenance of endothelial barrier integrity.

Microscopic imaging of live endothelial cells expressing GFP-actin allows characterization of dynamic changes in cytoskeletal structures. Unlike techniques that use fixed specimens, this method provides a detailed assessment of temporal changes in the actin cytoskeleton in the same cells before, during, and after various physical, pharmacological, or inflammatory stimuli.

The microvascular endothelium plays an important role as a selectively permeable barrier to fluids and solutes. The adhesive junctions between endothelial ...

A Step-by-step Method for the Reconstitution of an ABC Transporter into Nanodisc Lipid Particles

The nanodisc is a discoidal particle (~ 10-12 nm large) that trap membrane proteins into a small patch of phospholipid bilayer. The nanodisc is a particularly attractive option for studying membrane proteins, especially in the context of ligand-receptor interactions. The method pioneered by Sligar and colleagues is based on the amphipathic properties of an engineered highly a-helical scaffold protein derived from the apolipoprotein A1. The hydrophobic faces of the scaffold protein interact with the fatty acyl side-chains of the lipid bilayer whereas the polar regions face the aqueous environment. Analyses of membrane proteins in nanodiscs have significant advantages over liposome because the particles are small, homogeneous and water-soluble. In addition, biochemical and biophysical methods normally reserved to soluble proteins can be applied, and from either side of the membrane. In this visual protocol, we present a step-by-step reconstitution of a well characterized bacterial ABC transporter, the MalE-MalFGK2 complex. The formation of the disc is a self-assembly process that depends on hydrophobic interactions taking place during the progressive removal of the detergent. We describe the essential steps and we highlight the importance of choosing a correct protein-to-lipid ratio in order to limit the formation of aggregates and larger polydisperse liposome-like particles. Simple quality controls such as gel filtration chromatography, native gel electrophoresis and dynamic light scattering spectroscopy ensure that the discs have been properly reconstituted.

Nanodiscs are small discoid particles that incorporate membrane proteins into a small patch of phospholipid bilayer. We provide a visual protocol that shows the step-by-step incorporation of the MalFGK2 transporter into a disc.

The nanodisc is a discoidal particle (~ 10-12 nm large) that trap membrane proteins into a small patch of phospholipid bilayer. The nanodisc is a ...

Introduction to Solid Supported Membrane Based Electrophysiology

The electrophysiological method we present is based on a solid supported membrane (SSM) composed of an octadecanethiol layer chemisorbed on a gold coated sensor chip and a phosphatidylcholine monolayer on top. This assembly is mounted into a cuvette system containing the reference electrode, a chlorinated silver wire.
After adsorption of membrane fragments or proteoliposomes containing the membrane protein of interest, a fast solution exchange is used to induce the transport activity of the membrane protein. In the single solution exchange protocol two solutions, one non-activating and one activating solution, are needed. The flow is controlled by pressurized air and a valve and tubing system within a faraday cage.
The kinetics of the electrogenic transport activity is obtained via capacitive coupling between the SSM and the proteoliposomes or membrane fragments. The method, therefore, yields only transient currents. The peak current represents the stationary transport activity. The time dependent transporter currents can be reconstructed by circuit analysis.
This method is especially suited for prokaryotic transporters or eukaryotic transporters from intracellular membranes, which cannot be investigated by patch clamp or voltage clamp methods.

Here we present an electrophysiological method based on solid supported membranes with focus on its applications for the characterization of electrogenic membrane transporters.

The electrophysiological method we present is based on a solid supported membrane (SSM) composed of an octadecanethiol layer chemisorbed on a gold coated ...

Reconstitution of a Kv Channel into Lipid Membranes for Structural and Functional Studies

To study the lipid-protein interaction in a reductionistic fashion, it is necessary to incorporate the membrane proteins into membranes of well-defined lipid composition. We are studying the lipid-dependent gating effects in a prototype voltage-gated potassium (Kv) channel, and have worked out detailed procedures to reconstitute the channels into different membrane systems. Our reconstitution procedures take consideration of both detergent-induced fusion of vesicles and the fusion of protein/detergent micelles with the lipid/detergent mixed micelles as well as the importance of reaching an equilibrium distribution of lipids among the protein/detergent/lipid and the detergent/lipid mixed micelles. Our data suggested that the insertion of the channels in the lipid vesicles is relatively random in orientations, and the reconstitution efficiency is so high that no detectable protein aggregates were seen in fractionation experiments. We have utilized the reconstituted channels to determine the conformational states of the channels in different lipids, record electrical activities of a small number of channels incorporated in planar lipid bilayers, screen for conformation-specific ligands from a phage-displayed peptide library, and support the growth of 2D crystals of the channels in membranes. The reconstitution procedures described here may be adapted for studying other membrane proteins in lipid bilayers, especially for the investigation of the lipid effects on the eukaryotic voltage-gated ion channels.

Procedures for complete reconstitution of a prototype voltage-gated potassium channel into lipid membranes are described. The reconstituted channels are suitable for biochemical assays, electrical recordings, ligand screening and electron crystallographic studies. These methods may have general applications to the structural and functional studies of other membrane proteins.

To study the lipid-protein interaction in a reductionistic fashion, it is necessary to incorporate the membrane proteins into membranes of well-defined ...

Reconstitution Of &#946;-catenin Degradation In Xenopus Egg Extract

Xenopus laevis egg extract is a well-characterized, robust system for studying the biochemistry of diverse cellular processes. Xenopus egg extract has been used to study protein turnover in many cellular contexts, including the cell cycle and signal transduction pathways1-3. Herein, a method is described for isolating Xenopus egg extract that has been optimized to promote the degradation of the critical Wnt pathway component, &beta;-catenin. Two different methods are described to assess &beta;-catenin protein degradation in Xenopus egg extract. One method is visually informative ([35S]-radiolabeled proteins), while the other is more readily scaled for high-throughput assays (firefly luciferase-tagged fusion proteins). The techniques described can be used to, but are not limited to, assess &beta;-catenin protein turnover and identify molecular components contributing to its turnover. Additionally, the ability to purify large volumes of homogenous Xenopus egg extract combined with the quantitative and facile readout of luciferase-tagged proteins allows this system to be easily adapted for high-throughput screening for modulators of &beta;-catenin degradation.

Reconstitution Of &amp;#946;-catenin Degradation In Xenopus Egg Extract

A method is described for analyzing protein degradation using radiolabeled and luciferase-fusion proteins in Xenopus egg extract and its adaptation for high-throughput screening for small molecule modulators of protein degradation.

Xenopus laevis egg extract is a well-characterized, robust system for studying the biochemistry of diverse cellular processes. Xenopus egg extract has ...