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.

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<li>Shrivastava, R., Köster, D., Kalme, S., Mayor, S., Neerathilingam, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Tailor-made+ezrin+actin+binding+domain+to+probe+its+interaction+with+actin+in-vitro%5BTitle%5D)+AND+%22PLoS+One%22%5BJournal%5D)">Tailor-made ezrin actin binding domain to probe its interaction with actin in-vitro.</a> PLoS One. 10 (4), 0123428(2015).</li>
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</ol>

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><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-Ni&lt;sup&gt;2+&lt;/sup&gt;</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>CaCl&lt;sub&gt;2&lt;/sub&gt;</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 &amp;micro;L, 100 &amp;micro;L, 500 &amp;micro;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>KaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</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>MgCl&lt;sub&gt;2&lt;/sub&gt;</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>Na&lt;sub&gt;2&lt;/sub&gt;CO&lt;sub&gt;3&lt;/sub&gt;</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>NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</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 O&lt;sub&gt;2&lt;/sub&gt; 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

Research

JoVE Journal

Biology

2.1K Views.  University of Warwick. Hier demonstreren we een relatief eenvoudige en ongecompliceerde procedure om membraangebonden actomyosinenetwerken samen te stellen en de dynamiek te bestuderen met behulp van geavanceerde lichtmicroscopische technieken. Het belangrijkste voordeel is dat onze test de sequentiële toevoeging van eiwitten en kleine moleculen mogelijk maakt zonder de membraangebonden netwerken te verstoren. Deze methode kan eenvoudig worden aangepast om andere cytoskeletale eiwitten of samengestelde netwerken t...

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

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 ...

Biochemistry

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 ...

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.

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 ...

Visualizing Actin and Microtubule Coupling Dynamics In Vitro by Total Internal Reflection Fluorescence (TIRF) Microscopy

Traditionally, the actin and microtubule cytoskeletons have been studied as separate entities, restricted to specific cellular regions or processes, and regulated by different suites of binding proteins unique for each polymer. Many studies now demonstrate that the dynamics of both cytoskeletal polymers are intertwined and that this crosstalk is required for most cellular behaviors. A number of proteins involved in actin-microtubule interactions have already been identified (i.e., Tau, MACF, GAS, formins, and more) and are well characterized with regard to either actin or microtubules alone. However, relatively few studies showed assays of actin-microtubule coordination with dynamic versions of both polymers. This may occlude emergent linking mechanisms between actin and microtubules. Here, a total internal reflection fluorescence (TIRF) microscopy-based in vitro reconstitution technique permits the visualization of both actin and microtubule dynamics from the one biochemical reaction. This technique preserves the polymerization dynamics of either actin filament or microtubules individually or in the presence of the other polymer. Commercially available Tau protein is used to demonstrate how actin-microtubule behaviors change in the presence of a classic cytoskeletal crosslinking protein. This method can provide reliable functional and mechanistic insights into how individual regulatory proteins coordinate actin-microtubule dynamics at a resolution of single filaments or higher-order complexes.

Visualizing Actin and Microtubule Coupling Dynamics In Vitro by Total Internal Reflection Fluorescence TIRF Microscopy

This protocol is a guide for visualizing dynamic actin and microtubules using an in vitro total internal fluorescence (TIRF) microscopy assay.

Traditionally, the actin and microtubule cytoskeletons have been studied as separate entities, restricted to specific cellular regions or processes, and ...

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 ...

SNARE-mediated Fusion of Single Proteoliposomes with Tethered Supported Bilayers in a Microfluidic Flow Cell Monitored by Polarized TIRF Microscopy

In the ubiquitous process of membrane fusion the opening of a fusion pore establishes the first connection between two formerly separate compartments. During neurotransmitter or hormone release via exocytosis, the fusion pore can transiently open and close repeatedly, regulating cargo release kinetics. Pore dynamics also determine the mode of vesicle recycling; irreversible resealing results in transient, &#34;kiss-and-run&#34; fusion, whereas dilation leads to full fusion. To better understand what factors govern pore dynamics, we developed an assay to monitor membrane fusion using polarized total internal reflection fluorescence (TIRF) microscopy with single molecule sensitivity and ~15 msec time resolution in a biochemically well-defined in vitro system. Fusion of fluorescently labeled small unilamellar vesicles containing v-SNARE proteins (v-SUVs) with a planar bilayer bearing t-SNAREs, supported on a soft polymer cushion (t-SBL, t-supported bilayer), is monitored. The assay uses microfluidic flow channels that ensure minimal sample consumption while supplying a constant density of SUVs. Exploiting the rapid signal enhancement upon transfer of lipid labels from the SUV to the SBL during fusion, kinetics of lipid dye transfer is monitored. The sensitivity of TIRF microscopy allows tracking single fluorescent lipid labels, from which lipid diffusivity and SUV size can be deduced for every fusion event. Lipid dye release times can be much longer than expected for unimpeded passage through permanently open pores. Using a model that assumes retardation of lipid release is due to pore flickering, a pore &#34;openness&#34;, the fraction of time the pore remains open during fusion, can be estimated. A soluble marker can be encapsulated in the SUVs for simultaneous monitoring of lipid and soluble cargo release. Such measurements indicate some pores may reseal after losing a fraction of the soluble cargo.

Here, we present a protocol to detect single, SNARE-mediated fusion events between liposomes and supported bilayers in microfluidic channels using polarized TIRFM, with single molecule sensitivity and ~15 msec time resolution. Lipid and soluble cargo release can be detected simultaneously. Liposome size, lipid diffusivity, and fusion pore properties are measured.

In the ubiquitous process of membrane fusion the opening of a fusion pore establishes the first connection between two formerly separate compartments. ...

Neuroscience

Spontaneous Formation and Rearrangement of Artificial Lipid Nanotube Networks as a Bottom-Up Model for Endoplasmic Reticulum

We present a convenient method to form a bottom-up structural organelle model for the endoplasmic reticulum (ER). The model consists of highly dense lipidic nanotubes that are, in terms of morphology and dynamics, reminiscent of ER. The networks are derived from phospholipid double bilayer membrane patches adhering to a transparent Al2O3 substrate. The adhesion is mediated by Ca2+ in the ambient buffer. Subsequent depletion of Ca2+ by means of BAPTA/EDTA causes retraction of the membrane, resulting in spontaneous lipid nanotube network formation. The method only comprises phospholipids and microfabricated surfaces for simple formation of an ER model and does not require the addition of proteins or chemical energy (e.g., GTP or ATP). In contrast to the 3D morphology of the cellular endoplasmic reticulum, the model is two-dimensional (albeit the nanotube dimensions, geometry, structure, and dynamics are maintained). This unique in vitro ER model consists of only a few components, is easy to construct, and can be observed under a light microscope. The resulting structure can be further decorated for additional functionality, such as the addition of ER-associated proteins or particles to study transport phenomena among the tubes. The artificial networks described here are suitable structural models for the cellular ER, whose unique characteristic morphology has been shown to be related to its biological function, whereas details regarding formation of the tubular domain and rearrangements within are still not completely understood. We note that this method uses Al2O3 thin-film-coated microscopy coverslips, which are commercially available but require special orders. Therefore, it is advisable to have access to a microfabrication facility for preparation.

Solid-supported, protein-free, double phospholipid bilayer membranes (DLBM) can be transformed into complex and dynamic lipid nanotube networks and can serve as 2D bottom-up models of the endoplasmic reticulum.

We present a convenient method to form a bottom-up structural organelle model for the endoplasmic reticulum (ER). The model consists of highly dense ...

Bioengineering

Assembly of Cell Mimicking Supported and Suspended Lipid Bilayer Models for the Study of Molecular Interactions

Model cell membranes are a useful screening tool with applications ranging from early drug discovery to toxicity studies. The cell membrane is a crucial protective barrier for all cell types, separating the internal cellular components from the extracellular environment. These membranes are composed largely of a lipid bilayer, which contains outer hydrophilic head groups and inner hydrophobic tail groups, along with various proteins and cholesterol. The composition and structure of the lipids themselves play a crucial role in regulating biological function, including interactions between cells and the cellular microenvironment, which may contain pharmaceuticals, biological toxins, and environmental toxicants. In this study, methods to formulate uni-lipid and multi-lipid supported and suspended cell mimicking lipid bilayers are described. Previously, uni-lipid phosphatidylcholine (PC) lipid bilayers as well as multi-lipid placental trophoblast-inspired lipid bilayers were developed for use in understanding molecular interactions. Here, methods for achieving both types of bilayer models will be presented. For cell mimicking multi-lipid bilayers, the desired lipid composition is first determined via lipid extraction from primary cells or cell lines followed by liquid chromatography-mass spectrometry (LC-MS). Using this composition, lipid vesicles are fabricated using a thin-film hydration and extrusion method and their hydrodynamic diameter and zeta potential are characterized. Supported and suspended lipid bilayers can then be formed using quartz crystal microbalance with dissipation monitoring (QCM-D) and on a porous membrane for use in a parallel artificial membrane permeability assay (PAMPA), respectively. The representative results highlight the reproducibility and versatility of in vitro cell membrane lipid bilayer models. The methods presented can aid in rapid, facile assessment of the interaction mechanisms, such as permeation, adsorption, and embedment, of various molecules and macromolecules with a cell membrane, helping in the screening of drug candidates and prediction of potential cellular toxicity.

This protocol describes the formation of cell mimicking uni-lipid and multi-lipid vesicles, supported lipid bilayers, and suspended lipid bilayers. These in vitro models can be adapted to incorporate a variety of lipid types and can be used to investigate various molecule and macromolecule interactions.

Model cell membranes are a useful screening tool with applications ranging from early drug discovery to toxicity studies. The cell membrane is a crucial ...

Rapid Encapsulation of Reconstituted Cytoskeleton Inside Giant Unilamellar Vesicles

Giant unilamellar vesicles (GUVs) are frequently used as models of biological membranes and thus are a great tool to study membrane-related cellular processes in vitro. In recent years, encapsulation within GUVs has proven to be a helpful approach for reconstitution experiments in cell biology and related fields. It better mimics confinement conditions inside living cells, as opposed to conventional biochemical reconstitution. Methods for encapsulation inside GUVs are often not easy to implement, and success rates can differ significantly from lab to lab. One technique that has proven to be successful for encapsulating more complex protein systems is called continuous droplet interface crossing encapsulation (cDICE). Here, a cDICE-based method is presented for rapidly encapsulating cytoskeletal proteins in GUVs with high encapsulation efficiency. In this method, first, lipid-monolayer droplets are generated by emulsifying a protein solution of interest in a lipid/oil mixture. After being added into a rotating 3D-printed chamber, these lipid-monolayered droplets then pass through a second lipid monolayer at a water/oil interface inside the chamber to form GUVs that contain the protein system. This method simplifies the overall procedure of encapsulation within GUVs and speeds up the process, and thus allows us to confine and observe the dynamic evolution of network assembly inside lipid bilayer vesicles. This platform is handy for studying the mechanics of cytoskeleton-membrane interactions in confinement.

This article introduces a simple method for expeditious production of giant unilamellar vesicles with encapsulated cytoskeletal proteins. The method proves to be useful for bottom-up reconstitution of cytoskeletal structures in confinement and cytoskeleton-membrane interactions.

Giant unilamellar vesicles (GUVs) are frequently used as models of biological membranes and thus are a great tool to study membrane-related cellular ...

In Vitro Reconstitution of the Actin Cytoskeleton Inside Giant Unilamellar Vesicles

The actin cytoskeleton, the principal mechanical machinery in the cell, mediates numerous essential physical cellular activities, including cell deformation, division, migration, and adhesion. However, studying the dynamics and structure of the actin network in vivo is complicated by the biochemical and genetic regulation within live cells. To build a minimal model devoid of intracellular biochemical regulation, actin is encapsulated inside giant unilamellar vesicles (GUVs, also called liposomes). The biomimetic liposomes are cell-sized and facilitate a quantitative insight into the mechanical and dynamical properties of the cytoskeleton network, opening a viable route for bottom-up synthetic biology. To generate liposomes for encapsulation, the inverted emulsion method (also referred to as the emulsion transfer method) is utilized, which is one of the most successful techniques for encapsulating complex solutions into liposomes to prepare various cell-mimicking systems. With this method, a mixture of proteins of interest is added to the inner buffer, which is later emulsified in a phospholipid-containing mineral oil solution to form monolayer lipid droplets. The desired liposomes are generated from monolayer lipid droplets crossing a lipid/oil-water interface. This method enables the encapsulation of concentrated actin polymers into the liposomes with desired lipid components, paving the way for in vitro reconstitution of a biomimicking cytoskeleton network.

In Vitro Reconstitution of the Actin Cytoskeleton Inside Giant Unilamellar Vesicles

In this manuscript, we demonstrate the experimental techniques to encapsulate the F-actin cytoskeleton into giant unilamellar lipid vesicles (also called liposomes), and the method to form a cortex-biomimicking F-actin layer at the inner leaflet of the liposome membrane.

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

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