This work was funded by the European Research Council ERC Starting Grant ARTIST (# 757593 S.V.W.). DDI thanks the Alexander von Humboldt Foundation for a postdoctoral fellowship.

1. Experimental preparation
<ol>
	<li>Express and purify biotinylated-disiLID (b-disiLID) and mOrange-Nano (see Table of Materials) by following previously reported procedures7,8.</li>
	<li>Prepare lipid mixtures in glass vials with the selected lipid composition and concentrations. First, dissolve lipids in chloroform to obtain a final lipid solution with 1 mg/mL concentration.
	<ol>
		<li>Mix lipids in order to obtain a composition of 94.9 mol% 2-dioleoyl-sn-glycero-3-phosphocholine (DOPC), 5 mol% 1,2-dioleoyl-sn-glycero-3-phosphoethanolamineN-(cap biotinyl) sodium salt (DOPE-biotin) and 0.1 mol% 1,1&#39;-Dioctadecyl-3,3,3&#39;,3&#39;-Tetramethylindodicarbocyanine (DiD) (see Table of Materials). 
		NOTE: The lipid mixture can be adjusted with varying ratios and DOPE-biotin concentration and/or different membrane dyes. The recommended DOPE-biotin concentration (5 mol%) allows the formation of a high-density Streptavidin (SAv) layer in the following steps.</li>
	</ol>
	</li>
	<li>Prepare small unilamellar vesicles (SUVs) following previously reported methods7,8,12. For this step, it is recommended to prepare SUVs with a diameter &#8804;100 nm.</li>
	<li>For this study, sonication method is employed. First, evaporate the chloroform solution in the glass vial with a nitrogen stream while rotating the vials in order to form a thin lipid film. Next, remove the residual chloroform for at least 1 h under vacuum.
	<ol>
		<li>Rehydrate the dried film in ultrapure water with a final concentration of 1 mg/mL lipids by vortexing. Finally, sonicate the obtained solution for 10 min until the opaque solution turns clear. 
		NOTE: Store the lipid mixture in a microcentrifuge tube in the refrigerator for a maximum of 2 weeks. Different SUV preparation methods (e.g., extrusion method) can also be used as long as the size of the final SUVs is &#8804; 100 nm.</li>
	</ol>
	</li>
</ol>
2. mOrange-Nano recruitment to disiLID functionalized SLBs
<ol>
	<li>Add 150 &#181;L of 2 M NaOH in each well of the &#181;-slide 18 well glass-bottom chamber (see Table of Materials) and incubate for 1 h at room temperature. Subsequently, remove the NaOH and wash the wells 3-5 times first with 150 &#181;L ultrapure&#160;water, and then 3 times with 150 &#181;L buffer (10 mM Tris pH 7.4, 100 mM NaCl) containing 10 mM CaCl2.</li>
	<li>Add 15 &#181;L of freshly prepared SUVs (stock concentration 1 mg/mL in water) into the wells containing 150 &#181;L buffer with 10 mM CaCl2 in order to have approximately a factor 10 dilution of SUVs in the buffer. Let the SUVs incubate for 30 min at room temperature. After incubation time, biotinylated-SLBs will be formed.</li>
	<li>Wash the SLBs at least 7 times with buffer (10 mM Tris pH 7.4, 100 mM NaCl) without CaCl2 by first removing the solution and subsequently adding fresh buffer in each step. It is recommended to use 80 &#181;L of buffer for each washing step. 
	NOTE: An optimal washing of the SLBs is obtained by pipetting fresh solution up and down several times without touching the surface. Pipetting of the solution in the wells containing the newly formed SLBs should be gentle in order to reduce the formation of small air bubbles that would damage the formed SLBs. From this moment onwards, wells should contain a sufficient volume of buffer in order to prevent the SLBs from drying out.</li>
	<li>For the further functionalization of the biotinylated SLBs with SAv, add a solution of SAv to a final concentration of 250 nM and incubate for 30 min at room temperature. Subsequently, remove the excess of SAv by washing it with a buffer (10 mM Tris pH 7.4, 100 mM NaCl) at least 5 times.</li>
	<li>From this moment onwards, keep the samples under a protective red light in order to avoid unwanted photoactivation of photoswitchable proteins. Add b-disiILD (see Table of Materials) to a final concentration of 1 &#181;M in the well. After 30 min incubation at room temperature, remove the excess protein by washing with buffer at least 5 times.</li>
	<li>Add mOrange-Nano (see Table of Materials) to a final concentration of 200 nM and keep the sample in the dark by covering it with aluminum foil.</li>
	<li>Place the &#181;-slide under the fluorescence microscope and adjust the imaging settings. Set the 552 nm laser for excitation of mOrange-Nano. Adjust the emission range to optimize the mOrange signal. The photoactivation of disiLID is achieved with a 488 nm laser, using light pulses with 2.58 s intervals.</li>
</ol>
3. Preparation of GUVs
<ol>
	<li>Prepare a 5% (w/v) solution of polyvinyl alcohol (PVA, see Table of Materials) (MW: 145 000 g/ mol) with 100 mM sucrose in ultrapure water, mixing overnight at 80 &#176;C at 400 rpm.</li>
	<li>Prepare a lipid solution in chloroform with a desired composition (final concentration 10 mg/mL). For this method, it is recommended a composition consisting of 10 mg/mL POPC, 10 mol% 1-palmitoyl-2-oleoyl-sn-glycero-3-phospho-(1&#39;-rac-glycerol) (POPG), 2 mol% DOPE-biotin and 1 mol% DiD (see Table of Materials).</li>
	<li>Prepare GUVs with the hydration technique7,8. First, spread 40 &#181;L of the prepared PVA solution as a homogeneous thin layer on top of a 60 mm x 24 mm glass slide, preferably with a pipette tip. Next, dry the thin layer at 50 &#176;C for 30 min.</li>
	<li>Spread 5 &#181;L of the lipid solution with a needle on the PVA layer and let dry at 30 &#176;C for 1 h.</li>
	<li>Assemble a chamber on the functionalized glass slide using a spacer (~40 mm &#215; 24 mm &#215; 2 mm, see Table of Materials) and a second glass slide.</li>
	<li>Add 1 mL of rehydration buffer (10 mM Tris pH 7.4, 100 mM NaCl) into the chamber for 1 h at room temperature to form GUVs. After 1 h, invert the chamber and gently tap on the glass surfaces with a pipette tip.</li>
	<li>Carefully remove the glass slide on one side to open the built chamber and harvest the GUVs with a pipette.</li>
	<li>Place the solution in a plastic tube and let the GUVs settle for 2 h.</li>
</ol>
4. mOrange-Nano Recruitment to disiLID functionalized GUVs
<ol>
	<li>Add a solution of SAv to the freshly harvested GUVs and let sit for 30 min at room temperature. 
	NOTE: The following steps should be performed with protective red light in order to avoid photoactivation of disiLID.</li>
	<li>Add 1 &#181;M of b-disiLID to the GUVs solution and place the sample for 30 min in the dark, covering it with aluminum foil.</li>
	<li>Pretreat a &#181;-slide 18-well glass bottom chamber with 150 &#181;L BSA solution (3% w/v in water) for 10 min. Next, remove the BSA solution and wash the wells with 150 &#181;L ultrapure water 3 times.</li>
	<li>Add 145 &#181;L of 200 nM mOrange-Nano in buffer (10 mM Tris pH 7.4, 100 mM NaCl) to the well.</li>
	<li>Next, add 5 &#181;L of GUVs decorated with b-disiLID to the solution and wait ~15 min for the GUVs to settle.</li>
	<li>Place the &#181;-slide under the confocal microscope. Excite the sample at 552 nm to visualize mOrange (&#955;ex = 557 nm; &#955;em = 576 nm) fluorescence and at 638 nm to visualize DiD (&#955;ex = 644 nm; &#955;em = 665 nm) in the GUV membranes. The mOrange-Nano recruitment is triggered with blue light pulses (488 nm, intensity 1%) every 5.3 s in order to minimize undesired photobleaching effects. 
	NOTE: The excitation wavelengths can be adapted based on the type of microscope in use. Other common excitation wavelengths available for microscopes are also 532 nm or 561 nm and 633 nm, 647 nm, 639 nm, or 640 nm lasers.</li>
</ol>

Light-Controlled Reversible Protein Patterning on Lipid Membranes

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	<li>Kretschmer, S., 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=Pattern+formation+on+membranes+and+its+role+in+bacterial+cell+division.">Pattern formation on membranes and its role in bacterial cell division.</a> Curr Opin Cell Biol. 38, 52-59 (2016).</li><li>Yang, H. W., Collins, S. R., Meyer, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Locally+excitable+Cdc42+signals+steer+cells+during+chemotaxis.">Locally excitable Cdc42 signals steer cells during chemotaxis.</a> Nat Cell Biol. 18 (2), 191-201 (2016).</li><li>Caldwell, R. M., 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=Optochemical+control+of+protein+localization+and+activity+within+cell-like+compartments.">Optochemical control of protein localization and activity within cell-like compartments.</a> Biochem. 57 (18), 2590-2596 (2018).</li><li>Kennedy, M. J., 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=Rapid+blue-light-mediated+induction+of+protein+interactions+in+living+cells.">Rapid blue-light-mediated induction of protein interactions in living cells.</a> Nat Methods. 7 (12), 973-975 (2010).</li><li>Guntas, G., 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=Engineering+an+improved+light-induced+dimer+(iLID)+for+controlling+the+localization+and+activity+of+signaling+proteins.">Engineering an improved light-induced dimer (iLID) for controlling the localization and activity of signaling proteins.</a> Proc Natl Acad Sci. 112 (1), 112-117 (2015).</li><li>Levskaya, A., Weiner, O. D., Lim, W. A., Voigt, C. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spatiotemporal+control+of+cell+signalling+using+a+light-switchable+protein+interaction.">Spatiotemporal control of cell signalling using a light-switchable protein interaction.</a> Nature. 461 (7266), 997-1001 (2009).</li><li>Bartelt, S. M., 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=Dynamic+blue+light-switchable+protein+patterns+on+giant+unilamellar+vesicles.">Dynamic blue light-switchable protein patterns on giant unilamellar vesicles.</a> Chem Commun. 54 (8), 948-951 (2018).</li><li>Di Iorio, D., Bergmann, J., Higashi, S. L., Hoffmann, A., Wegner, S. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+disordered+tether+to+iLID+improves+photoswitchable+protein+patterning+on+model+membranes.">A disordered tether to iLID improves photoswitchable protein patterning on model membranes.</a> Chem. Commun. 59 (29), 4380-4383 (2023).</li><li>Hartzell, E. J., Terr, J., Chen, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Engineering+a+blue+light+inducible+Spytag+system+(BLISS).">Engineering a blue light inducible Spytag system (BLISS).</a> J Am Chem Soc. 143 (23), 8572-8577 (2021).</li><li>Xu, D., Bartelt, S. M., Rasoulinejad, S., Chen, F., Wegner, S. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Green+light+lithography:+a+general+strategy+to+create+active+protein+and+cell+micropatterns.">Green light lithography: a general strategy to create active protein and cell micropatterns.</a> Mater Horiz. 6 (6), 1222-1229 (2019).</li><li>Jia, H., 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=Light-induced+printing+of+protein+structures+on+membranes+in+vitro.">Light-induced printing of protein structures on membranes in vitro.</a> Nano Lett. 18 (11), 7133-7140 (2018).</li><li>Di Iorio, D., Verheijden, M. L., vander Vries, E., Jonkheijm, P., Huskens, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Weak+Multivalent+Binding+of+influenza+hemagglutinin+nanoparticles+at+a+sialoglycan-functionalized+supported+lipid+bilayer.">Weak Multivalent Binding of influenza hemagglutinin nanoparticles at a sialoglycan-functionalized supported lipid bilayer.</a> ACS Nano. 13 (3), 3413-3423 (2019).</li><li>Kasahara, M., Torii, M., Fujita, A., Tainaka, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=FMN+binding+and+photochemical+properties+of+plant+putative+photoreceptors+containing+two+LOV+domains,+LOV/LOV+proteins.">FMN binding and photochemical properties of plant putative photoreceptors containing two LOV domains, LOV/LOV proteins.</a> J Biol Chem. 285 (45), 34765-34772 (2010).</li></ol>

The authors have nothing to disclose.

We have described a method for the localized recruitment of mOrange-Nano proteins on model membranes, such as supported lipid bilayer and giant unilamellar vesicles by using the photoswitchable protein disiLID8. Aspects that contribute to the quality of the pattern include the quality of the proteins as well as the good quality of SLBs and GUVs.
To ensure a good protein quality after expression and purification, it is important to first evaluate the photoswitchable properties of disiLID. To this purpose, the absorption of the FMN cofactor has to be measured in the dark and after blue light illumination. UV-Vis spectra of disiLID are expected to show the characteristic triple peak of the cofactor FMN in the dark, which significantly decreases upon blue light illumination and recovers in the dark13. This photoswitchable behavior is crucial to obtaining reproducible and reversible recruitment in the following steps. Working with protective red light and exposing disiLID to minimum external illumination during the preparation of the samples increases the performance of the experiments.
Another critical step, and perhaps the most crucial, is the formation of proper SLBs. Defects in the membranes and/or the formation of inhomogeneous SLBs (i.e., the presence of multilayers or patched SLBs) will affect the quality of the protein patterning. Therefore, for inexperienced users, it is recommended to reproduce the protocol by labeling the SUVs with some membrane dyes, such as DiD and DiO, in order to form fluorescently labeled SLBs. In this way, the properties and quality of SLBs can be well characterized with fluorescence microscopy. FRAP measurements represent a typical approach to assessing the quality of an SLB by evaluating the fluidity of the membranes. Alternatively, in the case of biotinylated SLBs such as the ones described in this protocol, fluorescently labeled SAv (e.g., Atto 488-SAv) can be used to visualize and assess the quality of SLBs.
The first part of the protocol describes the formation of patterns on SLBs. To ensure an optimal result, it is important to add mOrange-Nano onto the SLBs and let the sample incubate in the dark for 15 min. During the photoactivation, the selection of the ROI is not restricted to a specific size. However, laser intensity and exposure time need to be regulated in order to reduce undesired photobleaching of the fluorescent proteins.
This method is not restricted to biotinylated proteins, and other approaches can be used to anchor disiLID to SLBs. For example, His-tagged disiLID can be expressed and anchored onto Ni- NTA-containing SLBs. However, it is crucial to express Nano and disiLID with different tags in order to avoid replacement of the proteins on the SLBs. This method also allows the possibility to invert the order of the proteins, thus functionalizing SLBs with Nano and recruiting disiLID (or disiLID-fused proteins) upon blue light illumination.
For dynamic control of protein localization, the reversible localization of the protein to the selected region should be possible repeatedly. To achieve this, the concentration of Nano (200 nM) in the solution is a critical parameter to obtain high reversibility.
Another concern is the recruitment of Nano to the disiLID-functionalized GUV surface. As in the case of protein patterning on SLBs, this method can be extended to different membrane functionalization strategies. In this protocol, the entire GUV was illuminated with blue light to recruit mOrange-Nano on the entire GUV surface. However, the selection of small ROIs localized on the GUV membrane should lead to the precise localization of proteins in a more restricted area.
This method presents only a limitation related to the choice of fluorophore employed for imaging of the Nano recruitment at the SUVs&#39; or GUVs&#39; membrane. In particular, fluorophores with an excitation spectrum in the blue-light range must be avoided, as their usage will interfere with the photoactivation of (dis)iLID. Therefore, the choice of fluorophores in the green or red-light range (e.g., mOrange or Cy5) is recommended for this type of experiment.
The disiLID design offers a simple and adaptable way to improve local protein recruitment to membranes and broadens the dynamic range of iLID and Nano from optogenetics4. These methods focus on the recruitment of Nano onto mimic membranes such as lipid bilayers and GUVs. Nevertheless, this approach is extendable to the numerous optogenetic tools in cells where (dis)iLID or Nano are linked to a membrane.

The formation of protein patterns on cell membranes within subcellular regions gives rise to numerous biological processes, including migration, division, and localized cell-to-cell communication1,2. These protein patterns are regulated in space and time, and are highly dynamic. Replicating such protein patterns in synthetic cells is essential for mimicking cellular processes that arise from them and for gaining a better understanding of how such regulation works at a molecular level. Analogous to what is observed for membranes in living cells, methods for generating protein patterns on artificial membranes must capture their dynamics and provide precise spatiotemporal control.
Among various stimuli, light stands out for providing the highest spatiotemporal control and several additional advantages3. Through regulation with light, it is straightforward to illuminate a desired area at any desired time with unmatched precision. In addition, light provides high tunability as both the light intensity and the pulse durations can be adjusted. Furthermore, visible light is harmless to biomolecules, including proteins, and it is possible to even address multiple functionalities with different wavelengths. Hence, light-responsive approaches based on visible light emerge as promising avenues for the controlled and biorthogonal regulation of protein patterns in space and time4,5,6. Utilizing photoswitchable protein pairs from optogenetics, acting as light-inducible dimerizers, provides a straightforward method for recruiting&#160;specific proteins to membranes. In particular, protein patterns have successfully been formed on artificial membranes using the blue light-triggered interaction between iLID (improved light-inducible dimer, based on the photoswitchable LOV2 domain from&#160;Avena sativa) and Nano (wild-type SspB)7,8, the blue light-inducible SpyTag system (BLISS)9, the green light-responsive&#160;protein tetramer CarH10 and the red light-inducible interaction between PhyB and PIF611.
It has been demonstrated that the photoswitchable interaction between iLID&#160;and Nano5 can be used to photo-pattern proteins on model membranes using blue light7. The iLID/Nano interaction is reversible in the dark, highly specific, and operates under physiological conditions. Anchoring iLID onto lipid membrane models, such as giant unilamellar vesicles (GUVs) or supported lipid bilayers (SLBs), enables light-regulated recruitment of Nano to these membranes, which is reversible in the dark. Notably, we observed that introducing a disordered domain to the N-terminus of iLID (resulting in a protein named disiLID) as a tether to a model lipid membrane enhances Nano recruitment efficiency and reversion dynamics8.
By employing the disiLID/Nano interaction, we have developed a method for generating high contrast patterns of Nano-fused proteins of interest (POI) on SLBs and the external membranes of GUVs. This method allows for the creation of protein patterns with remarkable spatial and temporal resolution and high reversibility within minutes. The detailed protocol outlines the process for locally recruiting proteins onto artificial membranes. Specifically, this is achieved by immobilizing a biotinylated version of disiLID on SLBs and GUVs through the biotin-streptavidin (SAv) interaction. Subsequently, fluorescently labeled Nano (mOrange-Nano) is recruited to these disiLID functionalized membranes under blue light illumination. Our experimental protocol offers a straightforward and adaptable approach for achieving localized protein recruitment to membranes. Importantly, this methodology is not confined to the reported SLB and GUV interfaces or mOrange-Nano; it can be extended to other disiLID-functionalized materials and proteins fused to Nano.

<table><tbody><tr><td>&amp;micro;-Slide 18 Well</td><td>Ibidi</td><td>81817</td><td>For SLB preparation</td></tr><tr><td>25 &amp;micro;L Microliter Syringe&amp;nbsp;</td><td>Hamilton</td><td>Model 702 N</td><td>For the preparation of lipid mixture and spreading the lipid solution on the PVA layer</td></tr><tr><td>Biotinyl Cap PE (1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-(cap biotinyl)) (sodium salt)</td><td>Avanti Polar Lipids</td><td>870273C</td><td>For SUVs preparation</td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt; (Calcium chloride)</td><td>Sigma-Aldrich</td><td>C5670</td><td>For SLB formation</td></tr><tr><td>Cover Slips 24 mm x 60 mm</td><td>Engelbrecht</td><td>K12460</td><td>For GUVs formation</td></tr><tr><td>DiD (1,1&#039;-Dioctadecyl-3,3,3&#039;,3&#039;-&lt;br/&gt; Tetramethylindodicarbocyanine)</td><td>Thermo Fisher Scientific&amp;nbsp;</td><td>D7757</td><td>Membrane dye</td></tr><tr><td>disiLID</td><td></td><td></td><td>Sequence: MGGSGLNDIFEAQKIEWHEGGSH&lt;br/&gt; HHHHHGSMAATELRGVVGPGPAA&lt;br/&gt; IAALGGGGAGPPVGGGGGRGDA&lt;br/&gt; GPGSGAASGTVVAAAAGGPGPG&lt;br/&gt; AGGVAAAGPPAPPTGGSGGSGA&lt;br/&gt; GGSGSAGEFLATTLERIEKNFVIT&lt;br/&gt; DPRLPDNPIIFASDSFLQLTEYSR&lt;br/&gt; EEILGRNCRFLQGPETDRATVRK&lt;br/&gt; IRDAIDNQTEVTVQLINYTKSGKK&lt;br/&gt; FWNVFHLQPMRDYKGDVQYFIG&lt;br/&gt; VQLDGTERLHGAAEREAVMLIKK&lt;br/&gt; TAFQIAEAANDENYF</td></tr><tr><td>DOPC (1,2-di-(9Z-octadecenoyl)-sn-glycero-3-phosphocholine)</td><td>Avanti Polar Lipids</td><td>850375P</td><td>For SUVs lipid composition</td></tr><tr><td>Eppendorf Protein LoBind&lt;br/&gt; microcentrifuge tubes</td><td>Merk</td><td>EP0030108116-100EA</td><td>For collecting freshly made GUVs</td></tr><tr><td>mOrange-Nano</td><td></td><td></td><td>Sequence: MRGSHHHHHHGSKIEEGKLVI&lt;br/&gt; WINGDKGYNGLAEVGKKFEKDT&lt;br/&gt; GIKVTVEHPDKLEEKFPQVAATG&lt;br/&gt; DGPDIIFWAHDRFGGYAQSGLLA&lt;br/&gt; EITPDKAFQDKLYPFTWDAVRYN&lt;br/&gt; GKLIAYPIAVEALSLIYNKDLLPNP&lt;br/&gt; PKTWEEIPALDKELKAKGKSALM&lt;br/&gt; FNLQEPYFTWPLIAADGGYAFKY&lt;br/&gt; ENGKYDIKDVGVDNAGAKAGLTF&lt;br/&gt; LVDLIKNKHMNADTDYSIAEAAFN&lt;br/&gt; KGETAMTINGPWAWSNIDTSKVN&lt;br/&gt; YGVTVLPTFKGQPSKPFVGVLSA&lt;br/&gt; GINAASPNKELAKEFLENYLLTDE&lt;br/&gt; GLEAVNKDKPLGAVALKSYEEELA&lt;br/&gt; KDPRIAATMENAQKGEIMPNIPQM&lt;br/&gt; SAFWYAVRTAVINAASGRQTVDEA&lt;br/&gt; LKDAQTNSSSNNNNNNNNNNLGI&lt;br/&gt; EGTTENLYFQGSVSKGEENNMAI&lt;br/&gt; IKEFMRFKVRMEGSVNGHEFEIE&lt;br/&gt; GEGEGRPYEGFQTAKLKVTKGG&lt;br/&gt; PLPFAWDILSPQFTYGSKAYVKH&lt;br/&gt; PADIPDYFKLSFPEGFKWERVMN&lt;br/&gt; FEDGGVVTVTQDSSLQDGEFIYK&lt;br/&gt; VKLRGTNFPSDGPVMQKKTMG&lt;br/&gt; WEASSERMYPEDGALKGEIKMR&lt;br/&gt; LKLKDGGHYTSEVKTTYKAKKPV&lt;br/&gt; QLPGAYIVGIKLDITSHNEDYTIVE&lt;br/&gt; QYERAEGRHSTGGMDELYKGG&lt;br/&gt; SGTSSPKRPKLLREYYDWLVDN&lt;br/&gt; SFTPYLVVDATYLGVNVPVEYVK&lt;br/&gt; DGQIVLNLSASATGNLQLTNDFIQ&lt;br/&gt; FNARFKGVSRELYIPMGAALAIYA&lt;br/&gt; RENGDGVMFEPEEIYDELNIG</td></tr><tr><td>NaCl (Sodium chloride)</td><td>Sigma-Aldrich</td><td>S9888</td><td>For buffer</td></tr><tr><td>NaOH (Sodium hydroxide)</td><td>Sigma-Aldrich</td><td>1064980500</td><td>For surface activation in SLB formation</td></tr><tr><td>POPC (1-Palmitoyl-2-&lt;br/&gt; oleoylphosphatidylcholine)</td><td>Avanti Polar Lipids</td><td>850457C</td><td>For GUVs lipid composition</td></tr><tr><td>PVA (Polyvinyl alcohol) fully hydrolyzed</td><td>Sigma-Aldrich</td><td>8148940101</td><td>For GUVs formation</td></tr><tr><td>SP8 confocal laser scanning microscope</td><td>Leica</td><td></td><td></td></tr><tr><td>Streptavidin</td><td>TermoFisher</td><td>434301</td><td></td></tr><tr><td>Sucrose&amp;nbsp;</td><td>Sigma-Aldrich&amp;nbsp;</td><td>84097</td><td>For GUVs formation</td></tr><tr><td>Tris hydrochloride</td><td>Sigma-Aldrich</td><td>10812846001</td><td>For buffer</td></tr></tbody></table>

dynamic light-induced protein patterns at model membranes

The precise localization and activation of proteins at the cell membrane at a certain time gives rise to many cellular processes, including cell polarization, migration, and division. Thus, methods to recruit proteins to model membranes with subcellular resolution and high temporal control are essential when reproducing and controlling such processes in synthetic cells. Here, a method is described for fabricating light-regulated reversible protein patterns at lipid membranes with high spatiotemporal precision. For this purpose, we immobilize the photoswitchable protein iLID (improved light-inducible dimer) on supported lipid bilayers (SLBs) and on the outer membrane of giant unilamellar vesicles (GUVs). Upon local blue light illumination, iLID binds to its partner Nano (wild-type SspB) and allows the recruitment of any protein of interest (POI) fused to Nano from the solution to the illuminated area on the membrane. This binding is reversible in the dark, which provides dynamic binding and release of the POI. Overall, this is a flexible and versatile method for regulating the localization of proteins with high precision in space and time using blue light.

The described procedures allow the formation of SLBs to recruit mOrange-Nano on the synthetic membranes. The formation of defined mOrange-Nano patterned on the SLBs functionalized with b-disiLID is shown in Figure 1A. As a squared (24 &#181;m &#215; 24 &#181;m) region of interest (ROI) on the SLB is illuminated with 488 nm blue light, a rapid increase of fluorescence signal is observed in the mOrange channel (shown in red) in the ROI within 200 s. The pattern shows very defined and sharp edges (Figure 1B), indicating high spatial control over the photoactivated area. The interaction is fast and fully reversible as the illumination with blue light is interrupted. This method also allows the formation of patterns over several illumination cycles (Figure 2). Alternate cycles of ~200 s of blue light and 200 s of dark lead to reversible recruitment of mOrange-Nano in the selected area for multiple times with comparable values of &#916; intensity of fluorescence in the patterns.
Figure 3 shows the schematic representation of the GUVs preparation. The recruitment of mOrange-Nano is also observed on GUVs. It is shown that GUVs placed in the dark do not exhibit mOrange fluorescence (Figure 4A). As the GUVs are globally illuminated with blue light, mOrange fluorescence is observed, colocalizing with the GUV membrane dye (DiD). The interaction is highly reversible as the illumination is terminated. The quantification of the mOrange intensity at the GUV membrane over time shows the fast and effective recruitment of proteins as well as the full reversibility (Figure 4B).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66531/66531fig01.jpg" /> 
Figure 1: Fluorescence microscopy images of SLBs functionalized with b-disiLID. The fluorescence images in the presence of mOrange-Nano before (A) and during (B) local blue light (488 nm) illumination in the ROI. Scale bar = 20 &#181;m. (C) Fluorescence intensity of mOrange measured in the ROI for SLBs functionalized with b-disiLID (at = 200 s). The figure is adapted from Di Iorio et al.8. <a href="https://www.jove.com/files/ftp_upload/66531/66531fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66531/66531fig02v3.jpg" /> 
Figure 2: Fluorescence intensity of mOrange recruited in the ROI on SLBs decorated with b-disiLID during three recruitment cycles. After each photoactivation step, the mOrange-Nano fluorescence increased within the ROI. The pattern reaches saturation within 120 s, and the fluorescence decreases within 120 s, almost to background levels. No loss of pattern quality is observed over different blue light/dark cycles. The figure is adapted from Di Iorio et al.8. <a href="https://www.jove.com/files/ftp_upload/66531/66531fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66531/66531fig03v2.jpg" /> 
Figure 3: Schematic representation of the GUVs preparation using the gentle hydration method. The scheme offers a visual representation of the several steps and of the chamber built using two glass slides and a spacer. <a href="https://www.jove.com/files/ftp_upload/66531/66531fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/66531/66531fig04.jpg" /> 
Figure 4: Fluorescence microscopy measurements of the light-dependent recruitment of mOrange-Nano on GUVs membranes. (A) Fluorescence images of a disiLID-functionalized GUV in the presence of mOrange-Nano. In green is the membrane dye of the GUVs, and in red is the mOrange fluorescence before, during, and after blue light illumination. Scale bars = 10 &#181;m. (B) Fluorescence intensity of mOrange localized on GUV over time. Upon illumination, the mOrange fluorescence (shown in red) on the lipid membrane reaches maximum intensity within 60 s, with a 5.9-fold fluorescence intensity increase. As the illumination is stopped, the mOrange fluorescence decreases to almost pre-illumination values within 60 s (with a 90% recovery). The figure is adapted from Di Iorio et al.8. <a href="https://www.jove.com/files/ftp_upload/66531/66531fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Read this Scientific Article Protocol about Author Spotlight: Photo Switchable Protein Recruitment for Reversible Patterning in Artificial Cellular Systems at JoVE.com

Author Spotlight: Photo Switchable Protein Recruitment for Reversible Patterning in Artificial Cellular Systems

Here, a protocol is described for generating light-regulated and reversible protein patterns with high spatiotemporal precision at artificial lipid membranes. The method consists of the localized photoactivation of the protein iLID (improved light-inducible dimer) immobilized on model membranes that, under blue light, binds to its partner protein Nano (wild-type SspB).

Dynamic Light-Induced Protein Patterns at Model Membranes

The precise localization and activation of proteins at the cell membrane at a certain time gives rise to many cellular processes, including cell ...

dynamic-light-induced-protein-patterns-at-model-membranes

Research

JoVE Journal

Bioengineering

1.0K Views.  University of Münster. Here, a protocol is described for generating light-regulated and reversible protein patterns with high spatiotemporal precision at artificial lipid membranes. The method consists of the localized photoactivation of the protein iLID (improved light-inducible dimer) immobilized on model membranes that, under blue light, binds to its partner protein Nano (wild-type SspB).

Video: Author Spotlight: Photo Switchable Protein Recruitment for Reversible Patterning in Artificial Cellular Systems

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

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

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

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

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 ...

Single Molecule Fluorescence Microscopy on Planar Supported Bilayers

In the course of a single decade single molecule microscopy has changed from being a secluded domain shared merely by physicists with a strong background in optics and laser physics to a discipline that is now enjoying vivid attention by life-scientists of all venues 1. This is because single molecule imaging has the unique potential to reveal protein behavior in situ in living cells and uncover cellular organization with unprecedented resolution below the diffraction limit of visible light 2. Glass-supported planar lipid bilayers (SLBs) are a powerful tool to bring cells otherwise growing in suspension in close enough proximity to the glass slide so that they can be readily imaged in noise-reduced Total Internal Reflection illumination mode 3,4. They are very useful to study the protein dynamics in plasma membrane-associated events as diverse as cell-cell contact formation, endocytosis, exocytosis and immune recognition. Simple procedures are presented how to generate highly mobile protein-functionalized SLBs in a reproducible manner, how to determine protein mobility within and how to measure protein densities with the use of single molecule detection. It is shown how to construct a cost-efficient single molecule microscopy system with TIRF illumination capabilities and how to operate it in the experiment.

Preparation of protein-functionalized planar glass-supported lipid bilayers, determination of protein mobility within and measurement of protein densities is shown here. A roadmap to building a noise-reduced Total Internal Reflection microscope is outlined, which allows visualizing single bilayer-resident fluorochromes with high spatiotemporal resolution.

In the course of a single decade single molecule microscopy has changed from being a secluded domain shared merely by physicists with a strong background ...

Ligand Nano-cluster Arrays in a Supported Lipid Bilayer

Currently there is considerable interest in creating ordered arrays of adhesive protein islands in a sea of passivated surface for cell biological studies. In the past years, it has become increasingly clear that living cells respond, not only to the biochemical nature of the molecules presented to them but also to the way these molecules are presented. Creating protein micro-patterns is therefore now standard in many biology laboratories; nano-patterns are also more accessible. However, in the context of cell-cell interactions, there is a need to pattern not only proteins but also lipid bilayers. Such dual proteo-lipidic patterning has so far not been easily accessible. We offer a facile technique to create protein nano-dots supported on glass and propose a method to backfill the inter-dot space with a supported lipid bilayer (SLB). From photo-bleaching of tracer fluorescent lipids included in the SLB, we demonstrate that the bilayer exhibits considerable in-plane fluidity. Functionalizing the protein dots with fluorescent groups allows us to image them and to show that they are ordered in a regular hexagonal lattice. The typical dot size is about 800 nm and the spacing demonstrated here is 2 microns. These substrates are expected to serve as useful platforms for cell adhesion, migration and mechano-sensing studies.

We present a protocol to functionalize glass with nanometric protein patches surrounded by a fluid lipid bilayer. These substrates are compatible with advanced optical microscopy and are expected to serve as platform for cell adhesion and migration studies.

Currently there is considerable interest in creating ordered arrays of adhesive protein islands in a sea of passivated surface for cell biological ...

Mass-Sensitive Particle Tracking to Characterize Membrane-Associated Macromolecule Dynamics

Short-lived or transient interactions of macromolecules at and with lipid membranes, an interface where a multitude of essential biological reactions take place, are inherently difficult to assess with standard biophysical methods. The introduction of mass-sensitive particle tracking (MSPT) constitutes an important step toward a thorough quantitative characterization of such processes. Technically, this was made possible through the advent of interferometric scattering microscopy (iSCAT)-based mass photometry (MP). When the background removal strategy is optimized to reveal the two-dimensional motion of membrane-associated particles, this technique allows the real-time analysis of both diffusion and molecular mass of unlabeled macromolecules on biological membranes. Here, a detailed protocol to perform and analyze mass-sensitive particle tracking of membrane-associated systems is described. Measurements performed on a commercial mass photometer achieve time resolution in the millisecond regime and, depending on the MP system, a mass detection limit down to 50 kDa. To showcase the potential of MSPT for the in-depth analysis of membrane-catalyzed macromolecule dynamics in general, results obtained for exemplary protein systems such as the native membrane interactor annexin V are presented.

This protocol describes an iSCAT-based image processing and single-particle tracking approach that enables the simultaneous investigation of the molecular mass and the diffusive behavior of macromolecules interacting with lipid membranes. Step-by-step instructions for sample preparation, mass-to-contrast conversion, movie acquisition, and post-processing are provided alongside directions to prevent potential pitfalls.

Short-lived or transient interactions of macromolecules at and with lipid membranes, an interface where a multitude of essential biological reactions take ...

Single-Molecule Diffusion and Assembly on Polymer-Crowded Lipid Membranes

Cellular membranes are highly crowded environments for biomolecular reactions and signaling. Yet, most in vitro experiments probing protein interaction with lipids employ naked bilayer membranes. Such systems lack the complexities of crowding by membrane-embedded proteins and glycans and exclude the associated volume effects encountered on cellular membrane surfaces. Also, the negatively charged glass surface onto which the lipid bilayers are formed prevents the free diffusion of transmembrane biomolecules. Here, we present a well-characterized polymer-lipid membrane as a mimic for crowded lipid membranes. This protocol utilizes polyethylene glycol (PEG)-conjugated lipids as a generalized approach for incorporating crowders into the supported lipid bilayer&#160;(SLB). First, a cleaning procedure of the microscopic slides and coverslips for performing single-molecule experiments is presented. Next, methods for characterizing the PEG-SLBs and performing single-molecule experiments of the binding, diffusion, and assembly of biomolecules using single-molecule tracking and photobleaching are discussed. Finally, this protocol demonstrates how to monitor the nanopore assembly of bacterial pore-forming toxin Cytolysin A (ClyA) on crowded lipid membranes with single-molecule photobleaching analysis. MATLAB codes with example datasets are also included to perform some of the common analyses such as particle tracking, extracting diffusive behavior, and subunit counting.

Here, a protocol to perform and analyze the binding, mobility, and assembly of single molecules on artificial crowded lipid membranes using single-molecule total internal reflection fluorescence (smTIRF) microscopy is presented.

Cellular membranes are highly crowded environments for biomolecular reactions and signaling. Yet, most in vitro experiments probing protein interaction ...

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

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

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

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

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

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane proteins including ion channels2, ion pumps3, and transporters4. Recent developments have combined site-specific fluorophore labeling alongside TEVC to 
concurrently examine the conformational dynamics at specific residues and function of these proteins on the surface of single cells.
We will describe a method to study the conformational dynamics of membrane proteins by simultaneously monitoring fluorescence and current changes using voltage-clamp fluorometry. This approach can be used to examine the molecular motion of membrane proteins site-specifically following cysteine replacement and site-directed fluorophore labeling5,6. Furthermore, this method provides an approach to determine distance constraints between specific residues7,8. 
This is achieved by selectively attaching donor and acceptor fluorophores to two mutated cysteine residues of interest.
In brief, these experiments are performed following functional expression of the desired protein on the surface of Xenopus leavis oocytes. The large surface area of these oocytes enables facile functional measurements and a robust fluorescence signal5. It is also possible to readily change the extracellular conditions such as pH, ligand or cations/anions, which can provide further information on the mechanism of membrane proteins4. Finally, recent developments 
have also enabled the manipulation of select internal ions following co-expression with a second protein9.
Our protocol is described in multiple parts. First, cysteine scanning mutagenesis proceeded by fluorophore labeling is completed at residues located at the interface of the transmembrane and extracellular domains. Subsequent experiments are designed to identify residues which demonstrate large changes in fluorescence intensity (&lt;5%)3 upon a conformational change of the protein. Second, these changes in fluorescence intensity are compared to the kinetic parameters of 
the membrane protein in order to correlate the conformational dynamics to the function of the protein10. This enables a rigorous biophysical analysis of the molecular motion of the target protein. Lastly, two residues of the holoenzyme can be labeled with a donor and acceptor fluorophore in order to determine distance constraints using donor photodestruction methods. It is also possible to monitor the relative movement of protein subunits following labeling with a donor and acceptor fluorophore.

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

We will describe a method which measures the kinetics of ion transport of membrane proteins alongside site-specific analysis of conformational changes using fluorescence on single cells. This technique is adaptable to ion channels, transporters and ion pumps and can be utilized to determine distance constraints between protein subunits.

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane ...

Biology

Light-Induced Molecular Adsorption of Proteins Using the PRIMO System for Micro-Patterning to Study Cell Responses to Extracellular Matrix Proteins

Cells sense a variety of extracellular cues, including the composition and geometry of the extracellular matrix, which is synthesized and remodeled by the cells themselves. Here, we present the method of Light-Induced Molecular Adsorption of Proteins (LIMAP) using the PRIMO system as a patterning technique to produce micro-patterned extracellular matrix (ECM) substrates using a single or combination of proteins. The method enables printing of ECM patterns in micron resolution with excellent reproducibility. We provide a step-by-step protocol and demonstrate how this can be applied to study the processes of neuronal pathfinding. LIMAP has significant advantages over existing micro-printing methods in terms of the ease of patterning more than one component and the ability to generate a pattern with any geometry or gradient. The protocol can easily be adapted to study the contribution of almost any chemical component towards cell fate and cell behavior. Finally, we discuss common issues that can arise and how these can be avoided.

Our overall aim is to understand how cells sense extracellular cues that lead to directed axonal growth. Here, we describe the methodology of Light-Induced Molecular Adsorption of Proteins, used to produce defined micro-patterns of extracellular matrix components in order to study specific events that govern axon outgrowth and pathfinding.

Cells sense a variety of extracellular cues, including the composition and geometry of the extracellular matrix, which is synthesized and remodeled by the ...

Neuroscience

Neutron Spin Echo Spectroscopy as a Unique Probe for Lipid Membrane Dynamics and Membrane-Protein Interactions

Lipid bilayers form the main matrix of cell membranes and are the primary platform for nutrient exchange, protein-membrane&#160;interactions, and viral budding, among other vital cellular processes. For efficient biological activity, cell membranes should be rigid enough to maintain the integrity of the cell and its compartments yet fluid enough to allow membrane components, such as proteins and functional domains, to diffuse and interact. This delicate balance of elastic and fluid membrane properties, and their impact on biological function, necessitate a better understanding of collective membrane dynamics over mesoscopic length and time scales of key biological processes, e.g., membrane deformations and protein binding events. Among the techniques that can effectively probe this dynamic range is neutron spin echo (NSE) spectroscopy. Combined with deuterium labeling, NSE can be used to directly access bending and thickness fluctuations as well as mesoscopic dynamics of select membrane features. This paper provides a brief description of the NSE technique and outlines the procedures for performing NSE experiments on liposomal membranes, including details of sample preparation and deuteration schemes, along with instructions for data collection and reduction. The paper also introduces data analysis methods used to extract key membrane parameters, such as the bending rigidity modulus, area compressibility modulus, and in-plane viscosity. To illustrate the biological importance of NSE studies, select examples of membrane phenomena probed by NSE are discussed, namely, the effect of additives on membrane bending rigidity, the impact of domain formation on membrane fluctuations, and the dynamic signature of membrane-protein interactions.

This paper describes the protocols for sample preparation, data reduction, and data analysis in neutron spin echo (NSE) studies of lipid membranes. Judicious deuterium labeling of lipids enables access to different membrane dynamics on mesoscopic length and time scales, over which vital biological processes occur.

Lipid bilayers form the main matrix of cell membranes and are the primary platform for nutrient exchange, protein-membrane&#160;interactions, and viral ...

Dynamic Light-Induced Protein Patterns at Model Membranes

Summary

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Chapters in this video

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

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

Single Molecule Fluorescence Microscopy on Planar Supported Bilayers

Ligand Nano-cluster Arrays in a Supported Lipid Bilayer

Mass-Sensitive Particle Tracking to Characterize Membrane-Associated Macromolecule Dynamics

Single-Molecule Diffusion and Assembly on Polymer-Crowded Lipid Membranes

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

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Light-Induced Molecular Adsorption of Proteins Using the PRIMO System for Micro-Patterning to Study Cell Responses to Extracellular Matrix Proteins

Neutron Spin Echo Spectroscopy as a Unique Probe for Lipid Membrane Dynamics and Membrane-Protein Interactions