This work was supported by the National Institutes of Health (NIH) Grant no. R01 GM-130934 and National Science Foundation (NSF) Grant MCB- 2016022. B.N.C, E.J.D.V., and K.S.C. were supported in part by a grant from the National Institute of General Medical Sciences under award T32 GM119999.

NOTE: Forming supported lipid bilayers requires the preparation of monodispersed small unilamellar vesicles (SUVs). Please refer to a previously published protocol24 on SUV formation. Briefly, all SUVs are formed by probe sonication for 12 min in total at 70% amplitude via 4 min sonication periods followed by 2 min rest periods in ice-water. SUV solutions must be well clarified and monodispersed in size. Size distributions of SUVs can be measured, for example, by dynamic light scattering25.
1. Planar lipid bilayers
<ol>
	<li>Plasma cleaning of the slides 
	NOTE: Plasma cleaning removes organic contaminants and increases the hydrophilicity of the glass, ensuring efficient lipid adsorption. The following steps may change or need to be optimized depending on the plasma cleaner and glass coverslips used.
	<ol>
		<li>Purge the plasma cleaner for 5 min with oxygen to remove air from the lines and chamber. This is to ensure that, when the plasma cleaner is run, there is primarily oxygen in the lines and chamber, providing more consistent bilayer preparations.</li>
		<li>While purging, stream an inert gas, such as N2 or Ar, over the micro cover glass slides to remove dust and particulates. 
		OPTIONAL: Cover glass slides can be washed by spraying each side with 100% isopropanol followed by water. Repeat the isopropanol-water wash 3x and then dry with an N2 stream.</li>
		<li>Arrange dry cover glass slides into a ceramic cradle. Place the cradle at the back of the plasma chamber so that the coverslips are parallel with the long edge of the chamber (this keeps them in place during plasma cleaning). Run the plasma cleaner for 15 min with oxygen at maximum power.</li>
	</ol>
	</li>
	<li>Chamber preparation 
	NOTE: The method described here uses homemade chambers from plastic PCR tubes to support the reaction volumes, but other low adhesion materials, such as silicone wells, may also be suitable.
	<ol>
		<li>Using a razor blade or scissors, cut off the cap just below the frosted part of a 0.2 mL PCR tube (Figure 2).</li>
		<li>Paint the rim of the PCR tube with UV-activated adhesive, avoiding the inside of the tube. Gently place the PCR tube glue-down in the center of a plasma-cleaned coverslip. 
		NOTE: To avoid glue inside the chamber, use the minimum amount of glue to get a seal, and promptly treat with UV without bumping or disturbing the chamber.</li>
		<li>Place the chamber under long-wavelength UV light for 5-7 min to cure the adhesive.</li>
	</ol>
	</li>
	<li>Bilayer formation 
	NOTE: Monodispersed SUVs should be used in this step for efficient bilayer formation. Table 1 provides recipes for all buffers used in the bilayer formation, including stock and final buffer concentrations.
	<ol>
		<li>Add 40 &#181;L of supported lipid bilayer buffer (SLBB: 300 mM KCl, 20 mM HEPES, pH 7.4, 1 mM MgCl2; Table 1), 10 &#181;L of 5 mM SUVs, and 1 &#181;L of 100 mM CaCl2 to the well. Gently shake the chambers from side to side to disrupt the SUVs, and then incubate at 37 &#176;C for 20 min. 
		NOTE: To limit evaporation, incubate in a covered Petri dish with a wet wipe.</li>
	</ol>
	</li>
	<li>Washing away excess lipids 
	NOTE: This step removes excess SUVs and acts as a buffer exchange step. It is very important not to scrape the bilayer with the pipette tip while washing; if the glass is exposed, septins and many other proteins will bind irreversibly, reducing the effective protein concentration.
	<ol>
		<li>After incubation, rinse the bilayer 6x with 150 &#181;L of SLBB, pipetting up and down to mix each time while avoiding bubbles. 
		NOTE: Add 150 &#181;L directly to the 50 &#181;L of buffer and SUVs already present for a total of 200 &#181;L in the reaction well.</li>
		<li>When ready to start incubating with the septins or a different protein of choice, wash the bilayer 6x with 150 &#181;L of reaction buffer (RXN: 33.3 mM KCl, 50 mM HEPES, pH 7.4, 1.4 mg/mL BSA, 0.14% methylcellulose, 1 mM BME). 
		NOTE: For best results, make the reaction buffer fresh and be very careful while mixing in the reaction well to avoid bubbles.</li>
		<li>On the last wash step, remove 125 &#181;L of reaction buffer, leaving 75 &#181;L in the well. Add 25 &#181;L of septins diluted in septin storage buffer (SSB: 300 mM KCl, 50 mM HEPES, pH 7.4, 1 mM BME) at the desired concentration and image by TIRF microscopy26. 
		NOTE: When setting up this assay for the first time or incorporating a new lipid composition, it is good practice to ensure lipids are freely diffusing on the surface using fluorescence recovery after photobleaching (FRAP). While the rate of recovery will be different for different lipid compositions and inherently lower than for free-standing systems, the lipids should not be immobile.</li>
	</ol>
	</li>
</ol>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="3">Supported Lipid Bilayer Buffer (SLBB)</td>
		</tr>
		<tr>
			<td>Stock</td>
			<td>Volume</td>
			<td>Final concentration</td>
		</tr>
		<tr>
			<td>2 M KCl</td>
			<td>1.5 mL</td>
			<td>300 mM</td>
		</tr>
		<tr>
			<td>1 M HEPES</td>
			<td>200 &#181;L</td>
			<td>20 mM</td>
		</tr>
		<tr>
			<td>500 mM MgCl2</td>
			<td>20 &#181;L</td>
			<td>1 mM</td>
		</tr>
		<tr>
			<td>Water</td>
			<td>8 mL</td>
			<td></td>
		</tr>
		<tr>
			<td colspan="3">Pre-Reaction Buffer (PRB)</td>
		</tr>
		<tr>
			<td>Stock</td>
			<td>Volume</td>
			<td>Final concentration</td>
		</tr>
		<tr>
			<td>2 M KCl</td>
			<td>166 &#181;L</td>
			<td>33.3 mM</td>
		</tr>
		<tr>
			<td>1 M HEPES</td>
			<td>500 &#181;L</td>
			<td>50 mM</td>
		</tr>
		<tr>
			<td>Water</td>
			<td>9.33 mL</td>
			<td></td>
		</tr>
		<tr>
			<td colspan="3">Reaction Buffer</td>
		</tr>
		<tr>
			<td>Stock</td>
			<td>Volume</td>
			<td>Final concentration</td>
		</tr>
		<tr>
			<td>2 M KCl</td>
			<td>166 &#181;L</td>
			<td>33.3 mM</td>
		</tr>
		<tr>
			<td>1 M HEPES</td>
			<td>300 &#181;L</td>
			<td>50 mM</td>
		</tr>
		<tr>
			<td>10 mg/mL BSA</td>
			<td>1.39 mL</td>
			<td>1.39 mg/mL</td>
		</tr>
		<tr>
			<td>1% Methylcellulose</td>
			<td>1.39 mL</td>
			<td>0.0014</td>
		</tr>
		<tr>
			<td>Water</td>
			<td>Up to 10 mL</td>
			<td></td>
		</tr>
		<tr>
			<td>BME</td>
			<td>0.7 &#181;L</td>
			<td>1 mM</td>
		</tr>
		<tr>
			<td colspan="3">Septin Storage Buffer (SSB)</td>
		</tr>
		<tr>
			<td>Stock</td>
			<td>Volume</td>
			<td>Final concentration</td>
		</tr>
		<tr>
			<td>2 M KCl</td>
			<td>1.5 mL</td>
			<td>300 mM</td>
		</tr>
		<tr>
			<td>1 M HEPES</td>
			<td>500 &#181;L</td>
			<td>50 mM</td>
		</tr>
		<tr>
			<td>Water</td>
			<td>Up to 10 mL</td>
			<td></td>
		</tr>
		<tr>
			<td>BME</td>
			<td>0.7 &#181;L</td>
			<td>1 mM</td>
		</tr>
	</tbody>
</table>
Table 1: Buffer components for preparation of supported lipid bilayer and reactions. Volumes of stock solutions that are incorporated into buffers and the final concentrations of each component are shown. SLB and PRB can be stored at room temperature and reused between experiments. Reaction buffer and SSB are made fresh for each experiment.
2. Spherical supported lipid bilayers
NOTE: This protocol uses silica microspheres suspended in ultrapure water at 10% density. For any work on the kinetic parameters of protein assembly, it is important to strictly control the total membrane surface area between experiments and curvatures. Table 2 shows the corrected volumes of beads and buffer to maintain 5 mm2 of total membrane surface area. This protocol expands on a previously published method8,18.
<ol>
	<li>Bilayer formation 
	NOTE: As for planar bilayers, it is important to have high-quality SUVs for robust bilayer formation. Table 2 lists the volume of beads from a 10% density solution and their respective SLBB volumes that are used to obtain a final surface area of 5 mm2 for a range of bead diameters.
	<ol>
		<li>Silica microspheres (also referred to as beads) tend to settle and clump together; to mix the beads and break up any clusters, vortex the bottle for 15 s, then bath-sonicate for 1 min, and vortex again for 15 s.</li>
		<li>Mix the appropriate volume of beads (Table 2) with the corresponding volume of SLBB and 10 &#181;L of 5 mM SUVs in a 0.5 mL low-adhesion microcentrifuge tube.</li>
		<li>Rock the bead-lipid mixture end-over-end for 1 h at room temperature. Ensure that this incubation is done on a rotator to prevent sedimentation.</li>
	</ol>
	</li>
	<li>Prepare chambers on PEGylated coverslips 
	NOTE: As for planar bilayers, homemade chambers from plastic PCR tubes are used to support the reaction volumes, but other low adhesion materials, such as silicone wells, may work just as well.
	<ol>
		<li>While the beads are rocking, thaw a PEGylated coverslip at room temperature. Cut a 0.2 mL PCR tube and glue it to the coverslip as described in step 1.2. For 24 mm x 50 mm slides, up to 10 chambers can feasibly fit on one slide. 
		NOTE: This protocol uses 24 mm x 50 mm PEGylated glass coverslips that are prepared in advance. Briefly, glass coverslips are thoroughly cleaned through sonication in acetone, methanol, and 3 M KOH. Coverslips are then functionalized with n-2-aminoethyl-3-aminopropyltrimethoxysilane and covalently linked to mPEG succinimidyl valerate. Finished PEGylated coverslips are stored vacuum-sealed at &#8722;80 &#176;C. For a similar passivation protocol, please see the work of Gidi et al.27. While PEGylated glass coverslips are suggested here, other means of passivation, such as BSA or poly-L-lysine methods, may be effective.</li>
	</ol>
	</li>
	<li>Washing away excess lipids 
	NOTE: This step functions to remove excess SUVs and perform a buffer exchange. Beads are washed 4x in total with pre-reaction buffer (PRB: 33.3 mM KCl, 50 mM HEPES, pH 7.4). Table 3 lists the spin speeds in RCF for a range of bead diameters.
	<ol>
		<li>Spin beads at the designated RCF for 30 s (Table 3). If using multiple bead sizes, keep beads rocking until it is their time to be washed. 
		NOTE: It is not worth sedimenting larger beads in the same spin as smaller beads with the smaller bead sedimentation velocity to save time. This can cause beads to stick to each other and compromise the integrity of the bilayer.</li>
		<li>After the first spin, remove 50 &#181;L of supernatant and add 200 &#181;L of PRB. After the second and third spins, remove 200 &#181;L and add 200 &#181;L of PRB. After the fourth spin, remove 200 &#181;L and add 220 &#181;L of PRB. Pipette vigorously for each wash. 
		NOTE: The final resuspension volume is different than the washes. Do not vortex the beads after incubation with the lipids as this can leave gaps in the bilayers. To avoid sedimentation while working with the other bead sizes or setting up other parts of the assay, place the bead mixtures back on the end-over-end rotator.</li>
	</ol>
	</li>
	<li>Preparing the reaction 
	NOTE: This reaction is designed to preserve the total membrane surface area of the reaction at 5 mm2 and to produce a final reaction condition of 100 mM KCl, 50 mM HEPES, 1 mg/mL BSA, 0.1% methylcellulose, and 1 mM BME.
	<ol>
		<li>If doing a competition assay with multiple bead sizes, make a 1:1 mixture by mixing equal volumes of each bead size together. Then, mix 29 &#181;L of the desired bead mixture (or of a single bead) with 721 &#181;L of RXN buffer. 
		NOTE: It is important to thoroughly mix the beads at each step with a pipette to avoid dense clusters of beads; this will result in a more even bead distribution and accurate total surface area.</li>
		<li>Mix 75 &#181;L of diluted beads and 25 &#181;L of protein diluted in SSB to the wells. 
		NOTE: The authors typically image yeast septin complexes at 1-50 nM.</li>
		<li>If measuring septins at a steady-state, incubate at room temperature for 1 h, and then image by either near-TIRF or confocal microscopy.</li>
	</ol>
	</li>
</ol>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Bead Diameter&#160;(&#181;m)</td>
			<td>Volume of well-mixed beads (&#181;L)</td>
			<td>Volume of SLB buffer (&#181;L)</td>
			<td>Volume of SUVs (&#181;L)</td>
		</tr>
		<tr>
			<td>6.46</td>
			<td>8.94</td>
			<td>61.1</td>
			<td>10</td>
		</tr>
		<tr>
			<td>5.06</td>
			<td>7</td>
			<td>63</td>
			<td>10</td>
		</tr>
		<tr>
			<td>3.17</td>
			<td>4.39</td>
			<td>65.6</td>
			<td>10</td>
		</tr>
		<tr>
			<td>0.96</td>
			<td>1.33</td>
			<td>68.7</td>
			<td>10</td>
		</tr>
		<tr>
			<td>0.54</td>
			<td>0.75</td>
			<td>69.3</td>
			<td>10</td>
		</tr>
		<tr>
			<td>0.31</td>
			<td>0.43</td>
			<td>69.6</td>
			<td>10</td>
		</tr>
	</tbody>
</table>
Table 2: Normalized volumes of microspheres. In order to maintain an equal surface area of each bead size and to keep the total membrane surface area consistent between experiments, volumes for each bead size and buffer that normalized the total surface area were calculated.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Bead diameter (&#181;m)</td>
			<td>Sedimentation velocity (RCF)</td>
		</tr>
		<tr>
			<td>0.31</td>
			<td>4.5</td>
		</tr>
		<tr>
			<td>0.54</td>
			<td>4.5</td>
		</tr>
		<tr>
			<td>0.96</td>
			<td>2.3</td>
		</tr>
		<tr>
			<td>3.17</td>
			<td>0.8</td>
		</tr>
		<tr>
			<td>5.06</td>
			<td>0.3</td>
		</tr>
	</tbody>
</table>
Table 3: Sedimentation velocities for microspheres of varying diameters. For each bead diameter, the shown minimum sedimentation velocities were used to pellet the beads for washing away unbound liposomes.
3. Rod supported lipid bilayers
NOTE: In contrast to the other assays presented here, the rod assay does not allow for careful control of the total membrane surface area. One can be consistent in amounts and volumes between experiments, but because this results in rods of different lengths and diameters, it is difficult to extrapolate the total membrane surface area in the reaction. Thus, while this is an excellent assay for exploring curvature sensing with multiple curvatures on a single surface and has been useful for exploring septin ultrastructure, it is not recommended for kinetic measurements. This method was previously reported18 and is being expanded upon here.
<ol>
	<li>Obtaining borosilicate rods from glass microfiber filters
	<ol>
		<li>Tear a single 42.5 mm grade GF/C glass microfiber filterinto small pieces and place them into a 100 mL beaker with 60 mL of 100% ethanol. Bath-sonicate until the solution becomes opaque; this can take as little as 20-30 min with a finely shredded filter. 
		NOTE: Sonicate thoroughly to break up big chunks; the more dissociated/opaque, the better.</li>
		<li>Cover the solution with film and store the solution at room temperature overnight.</li>
	</ol>
	</li>
	<li>Forming bilayers on the rods 
	NOTE: This step is performed with excess lipids to achieve complete rod coverage as it is impossible to quantify the total surface area in this method.
	<ol>
		<li>The next day the ethanol solution will be settled (step 3.1.2.), so mix it well before using. Combine 10 &#181;L of rod solution and 70 &#181;L of SLBB.</li>
		<li>Spin at top speed in a mini centrifuge for 30 s and remove 50 &#181;L of the supernatant. Resuspend in another 50 &#181;L of SLBB and repeat the spin for a total of four washes to dilute out the ethanol. 
		NOTE: Rod supports will not form a solid pellet at the bottom of the tube. They are instead smeared along the side of the tube; this makes them difficult to completely preserve when washing. Since the total surface area in this assay is not controlled, it is fine if they are disturbed.</li>
		<li>Combine 10 &#181;L of well-mixed rod solution with 50 &#181;L of 5 mM SUVs and 20 &#181;L of SLBB. Incubate for 1 h at room temperature, rotating end-over-end as when forming bilayers on the microspheres.</li>
	</ol>
	</li>
	<li>Washing away excess lipids
	<ol>
		<li>Similar to step 3.2.2., spin the lipid-rod solution at top speed in a mini centrifuge for 30 s to pellet the membrane-coated rods out of the solution.</li>
		<li>After the first spin, remove 50 &#181;L of the supernatant and add 200 &#181;L of PRB. After the second and third spins, remove 200 &#181;L and add 200 &#181;L of PRB. After the fourth spin, remove 200 &#181;L, add 220 &#181;L of PRB, and pipette vigorously to mix.</li>
	</ol>
	</li>
	<li>Preparing the reaction
	<ol>
		<li>In a 1.5 mL microcentrifuge tube, mix 721 &#181;L of reaction buffer with 29 &#181;L of rod solution. Mix well.</li>
		<li>In a 0.2 mL PCR tube, combine 75 &#181;L of rods in reaction buffer and 25 &#181;L of septins at the desired concentration, diluted in SSB. Allow it to incubate at room temperature for at least 1 h.</li>
	</ol>
	</li>
	<li>Preparing for scanning electron microscopy
	<ol>
		<li>After incubation, add the 100 &#181;L septin-rod mixture onto a round, PEG-coated 12 mm coverslip and incubate at room temperature for 1 h in a closed Petri dish.</li>
		<li>Fix the sample by filling the bottom of the Petri dish (10 mL should be enough) with 2.5% glutaraldehyde in 0.05 M sodium cacodylate (NaCo), pH 7.4, for 30 min. Aspirate the liquid with a glass pipette. Wash the sample by incubating it with 10 mL of 0.05 M NaCo for 5 min and repeat for a total of two washes.</li>
		<li>Postfix in 0.5% OsO4 cacodylate buffer for 30 min, and then wash 3x in 0.05 M NaCo (5 min/wash).</li>
		<li>Incubate the sample with 1% tannic acid for 15 min, and then wash in NaCo 3x. Incubate for 15 min in 0.5% OsO4 and wash 3x with NaCo.</li>
		<li>Dehydrate the samples using increasing ethanol concentrations: 30% EtOH for 5 min, 2x; 50% EtOH for 5 min; 75% EtOH for 5 min; and 100% EtOH for 5 min, 2x. Follow with another 10 min incubation.</li>
		<li>Incubate in transition fluid (hexamethyldisilazane) 3x (5 min, 10 min, then 5 min).</li>
		<li>Allow to air dry, then place the samples in a desiccator until sputter-coating. Sputter-coat the 4 nm layer with a gold/palladium alloy and then image on a scanning electron microscope.</li>
	</ol>
	</li>
</ol>

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A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stochasticity+and+positive+feedback+enable+enzyme+kinetics+at+the+membrane+to+sense+reaction+size.">Stochasticity and positive feedback enable enzyme kinetics at the membrane to sense reaction size.</a> Proceedings of the National Academy of Sciences. 118 (47), 2103626118 (2021).</li><li>Ferhan, A. R., 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=Nanoplasmonic+sensing+architectures+for+decoding+membrane+curvature-dependent+biomacromolecular+interactions.">Nanoplasmonic sensing architectures for decoding membrane curvature-dependent biomacromolecular interactions.</a> Analytical Chemistry. 90 (12), 7458-7466 (2018).</li><li>Beber, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Membrane+reshaping+by+micrometric+curvature+sensitive+septin+filaments.">Membrane reshaping by micrometric curvature sensitive septin filaments.</a> Nature Communications. 10 (1), 420 (2019).</li><li>Lou, H. -. Y., 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=Membrane+curvature+underlies+actin+reorganization+in+response+to+nanoscale+surface+topography.">Membrane curvature underlies actin reorganization in response to nanoscale surface topography.</a> Proceedings of the National Academy of Sciences. 116 (46), 23143-23151 (2019).</li><li>Bridges, A. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Septin+assemblies+form+by+diffusion-driven+annealing+on+membranes.">Septin assemblies form by diffusion-driven annealing on membranes.</a> Proceedings of the National Academy of Sciences. 111 (6), 2146-2151 (2014).</li><li>Cannon, K. S., Woods, B. L., Crutchley, J. M., Gladfelter, A. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+amphipathic+helix+enables+septins+to+sense+micrometer-scale+membrane+curvature.">An amphipathic helix enables septins to sense micrometer-scale membrane curvature.</a> Journal of Cell Biology. 218 (4), 1128-1137 (2019).</li><li>Johnson, J. M., Ha, T., Chu, S., Boxer, S. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Early+steps+of+supported+bilayer+formation+probed+by+single+vesicle+fluorescence+assays.">Early steps of supported bilayer formation probed by single vesicle fluorescence assays.</a> Biophysical Journal. 83 (6), 3371-3379 (2002).</li><li>Lobato-M&#225;rquez, D., 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=Mechanistic+insight+into+bacterial+entrapment+by+septin+cage+reconstitution.">Mechanistic insight into bacterial entrapment by septin cage reconstitution.</a> Nature Communications. 12 (1), 4511 (2021).</li><li>Gladfelter, A. S., Pringle, J. R., Lew, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+septin+cortex+at+the+yeast+mother-bud+neck.">The septin cortex at the yeast mother-bud neck.</a> Current Opinion in Microbiology. 4 (6), 681-689 (2001).</li><li>Ong, K., Wloka, C., Okada, S., Svitkina, T., Bi, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Architecture+and+dynamic+remodelling+of+the+septin+cytoskeleton+during+the+cell+cycle.">Architecture and dynamic remodelling of the septin cytoskeleton during the cell cycle.</a> Nature Communications. 5, 1-10 (2014).</li><li>Bertin, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phosphatidylinositol-4,5-bisphosphate+promotes+budding+yeast+septin+filament+assembly+and+organization.">Phosphatidylinositol-4,5-bisphosphate promotes budding yeast septin filament assembly and organization.</a> Journal of Molecular Biology. 404 (4), 711-731 (2010).</li><li>Bridges, A. A., Gladfelter, A. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+reconstitution+of+septin+assemblies+on+supported+lipid+bilayers.">In vitro reconstitution of septin assemblies on supported lipid bilayers.</a> Methods in Cell Biology. 136, 57-71 (2016).</li><li>Hupfeld, S., Hols&#230;ter, A. M., Skar, M., Frantzen, C. B., Brandl, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Liposome+size+analysis+by+dynamic/static+light+scattering+upon+size+exclusion-/field+flow-fractionation.">Liposome size analysis by dynamic/static light scattering upon size exclusion-/field flow-fractionation.</a> Journal of Nanoscience and Nanotechnology. 6 (9), 3025-3031 (2006).</li><li>Johnson, D. S., Jaiswal, J. K., Simon, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Total+internal+reflection+fluorescence+(TIRF)+microscopy+illuminator+for+improved+imaging+of+cell+surface+events.">Total internal reflection fluorescence (TIRF) microscopy illuminator for improved imaging of cell surface events.</a> Current Protocols in Cytometry. (1), Chapter 12, Unit 12.29 (2012).</li><li>Gidi, Y., Bayram, S., Ablenas, C. J., Blum, A. S., Cosa, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efficient+one-step+PEG-silane+passivation+of+glass+surfaces+for+single-molecule+fluorescence+studies.">Efficient one-step PEG-silane passivation of glass surfaces for single-molecule fluorescence studies.</a> ACS Applied Materials &amp; Interfaces. 10 (46), 39505-39511 (2018).</li><li>Woods, B. L., 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=Biophysical+properties+governing+septin+assembly.">Biophysical properties governing septin assembly.</a> bioRxiv. , (2021).</li><li>Cannon, K. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+gene+duplication+of+a+septin+provides+a+developmentally-regulated+filament+length+control+mechanism.">A gene duplication of a septin provides a developmentally-regulated filament length control mechanism.</a> bioRxiv. , (2021).</li><li>Pincet, F., 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=FRAP+to+characterize+molecular+diffusion+and+interaction+in+various+membrane+environments.">FRAP to characterize molecular diffusion and interaction in various membrane environments.</a> PLOS One. 11 (7), 0158457 (2016).</li><li>Reimhult, E., H&#246;&#246;k, F., Kasemo, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intact+vesicle+adsorption+and+supported+biomembrane+formation+from+vesicles+in+solution:+Influence+of+surface+chemistry,+vesicle+size,+temperature,+and+osmotic+pressure.">Intact vesicle adsorption and supported biomembrane formation from vesicles in solution: Influence of surface chemistry, vesicle size, temperature, and osmotic pressure.</a> Langmuir. 19 (5), 1681-1691 (2003).</li><li>Cha, T., Guo, A., Zhu, X. -. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Formation+of+supported+phospholipid+bilayers+on+molecular+surfaces:+Role+of+surface+charge+density+and+electrostatic+interaction.">Formation of supported phospholipid bilayers on molecular surfaces: Role of surface charge density and electrostatic interaction.</a> Biophysical Journal. 90 (4), 1270-1274 (2006).</li><li>Andrews, J. T., 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=Formation+of+supported+lipid+bilayers+(SLBs)+from+buffers+containing+low+concentrations+of+group+I+chloride+salts.">Formation of supported lipid bilayers (SLBs) from buffers containing low concentrations of group I chloride salts.</a> Langmuir. 37 (44), 12819-12833 (2021).</li><li>Gurtovenko, A. A., Vattulainen, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effect+of+NaCl+and+KCl+on+phosphatidylcholine+and+phosphatidylethanolamine+lipid+membranes:+Insight+from+atomic-scale+simulations+for+understanding+salt-induced+effects+in+the+plasma+membrane.">Effect of NaCl and KCl on phosphatidylcholine and phosphatidylethanolamine lipid membranes: Insight from atomic-scale simulations for understanding salt-induced effects in the plasma membrane.</a> The Journal of Physical Chemistry B. 112 (7), 1953-1962 (2008).</li><li>Danuser, G., Waterman-Storer, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantitative+fluorescent+speckle+microscopy+of+cytoskeleton+dynamics.">Quantitative fluorescent speckle microscopy of cytoskeleton dynamics.</a> Annual Review of Biophysics and Biomolecular Structure. 35 (1), 361-387 (2006).</li><li>Chan, Y. -. H. M., Boxer, S. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Model+membrane+systems+and+their+applications.">Model membrane systems and their applications.</a> Current Opinion in Chemical Biology. 11 (6), 581-587 (2007).</li></ol>

The authors have no conflicts of interest.

Cell membranes take on many different shapes, curvatures, and physicochemical properties. In order to study the nanometer-scale machinery through which cells build micrometer-scale assemblies, it is necessary to design minimal reconstitution systems of membrane mimetics. This protocol presents techniques that precisely control both membrane curvature and composition while allowing the user to easily take quantitative fluorescence measurements using widely available microscopy techniques.
The m...

The shapes of cells and many of their internal compartments are dependent on the lipid membranes that surround them. Membranes are viscoelastic structures that can be deformed through interactions with proteins, lipid sorting, and acting internal and external forces to generate a variety of shapes1,2,3,4. These shapes are often described in terms of membrane curvature. Cells use a diverse suite of proteins capable of preferentially assembling onto, or &#34;sensing&#34;, particular membrane curvatures to ensure defined spatio-temporal control over processes including cell trafficking, cytokinesis, and migration5,6. The dynamics of cell machinery at the membrane are notably difficult to observe due to the difficulty of balancing time and spatial resolution with cell health. While super-resolution techniques can offer a detailed view of such structures, they require lengthy acquisitions that are not amenable to the timescales of assembly/disassembly for most machinery. Additionally, the molecular complexity of these assemblies in their native environment and the multitude of roles a single component can play make minimal reconstitution systems a valuable tool for studying the functional capacity of molecules.
Minimal membrane mimetics have been developed to study membrane properties and protein-membrane interactions outside of the cell. Membrane mimetics vary from free-standing lipid bilayers, such as liposomes or giant unilamellar vesicles, to supported lipid bilayers (SLBs)7,8,9,10. SLBs are biomimetic membranes anchored to underlying support, typically composed of glass, mica, or silica11,12.&#160;A variety of geometries can be used, including planar surfaces, spheres, rods, and even undulating or micropatterned substrates to probe protein-membrane interactions on both concave and convex curvatures simultaneously13,14,15,16,17,18. Bilayer formation begins with vesicle adsorption onto a hydrophilic surface, followed by fusion and rupture to form a continuous bilayer (Figure 1)19. Supported bilayers are particularly amenable to light and electron microscopy, providing both better time and spatial resolution than is often achievable in cells. Curved SLBs especially provide an attractive means to probe protein curvature sensitivity in the absence of significant membrane deformation, allowing one to distinguish between curvature sensing and curvature induction, which are often impossible to separate in free-standing systems.
Septins are a class of filament-forming cytoskeletal proteins well known for their ability to assemble on positively curved membranes6,18,20. Over the course of the cell cycle in yeast, septins assemble into a ring and must rearrange to form the hourglass and double ring structures associated with bud emergence and cytokinesis, respectively21. While beautiful work has been done using platinum replica electron microscopy to observe septin architecture at varying cell cycle stages22, watching septin assembly over time using light microscopy in yeast has met with limited spatial resolution. Previous work on septins using lipid monolayers visualized by transmission electron microscopy (TEM) was able to reconstitute several interesting septin structures such as rings, bundles, and gauzes23. However, EM techniques are likewise limited in their temporal resolution, unlike fluorescence microscopy. In order to better resolve the kinetic parameters of the multi-scale process of septin assembly, we turned to supported membrane mimetics, where one can carefully control membrane geometry, sample conditions, and imaging modality.
The protocols described here use planar or curved SLBs, purified protein, and a combination of microscopy techniques. Quantitative fluorescence confocal microscopy and total internal reflection fluorescence microscopy (TIRFM) were used to measure both bulk protein binding onto various membrane curvatures, as well as to measure the binding kinetics of single molecules. Furthermore, this protocol has been adapted to be used with scanning electron microscopy (SEM) to examine protein ultrastructure on different membrane curvatures. While the focus of these protocols is on the septin cytoskeleton, the protocols can be easily modified to investigate the curvature sensitivity of any protein the reader finds interesting. Additionally, those working in fields such as endocytosis or vesicular trafficking may find these techniques useful for probing the curvature-dependent assemblies of multi-protein complexes.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.2 mL PCR Tubes with flat cap, Natural</td>
			<td>Watson</td>
			<td>137-211C(EX)</td>
			<td></td>
		</tr>
		<tr>
			<td>0.5 mL low adhesion tubes</td>
			<td>USA Scientific</td>
			<td>1405-2600</td>
			<td></td>
		</tr>
		<tr>
			<td>Beta mercaptoethanol (BME)</td>
			<td>Sigma-Aldrich</td>
			<td>M6250-100ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin (BSA)</td>
			<td>Sigma-Aldrich</td>
			<td>A4612-25G</td>
			<td></td>
		</tr>
		<tr>
			<td>Coverglass for making PEGylated coverslips</td>
			<td>Thermo Scientific</td>
			<td>152450</td>
			<td>Richard-Allan Scientific SLIP-RITE Cover Glass 24x50 #1.5</td>
		</tr>
		<tr>
			<td>DOPC</td>
			<td>Avanti Polar Lipids</td>
			<td>850375</td>
			<td></td>
		</tr>
		<tr>
			<td>Egg Liss Rhodamine PE</td>
			<td>Avanti Polar Lipids</td>
			<td>810146</td>
			<td></td>
		</tr>
		<tr>
			<td>EMS Glutaraldehyde Aqueous 25%, EM Grade</td>
			<td>VWR</td>
			<td>16220</td>
			<td></td>
		</tr>
		<tr>
			<td>EMS Sodium Cacodylate Buffer</td>
			<td>VWR</td>
			<td>11652</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol, 200 proof</td>
			<td>Fisher Scientific</td>
			<td>04-355-223EA</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma Aldrich</td>
			<td>H3375-1KG</td>
			<td></td>
		</tr>
		<tr>
			<td>Hexamethyldisilazane</td>
			<td>Sigma-Aldrich</td>
			<td>440191</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium chloride</td>
			<td>VWR</td>
			<td>7791-18-6</td>
			<td></td>
		</tr>
		<tr>
			<td>Methyl cellulose 4000cp</td>
			<td>Sigma-Aldrich</td>
			<td>M052-100G</td>
			<td></td>
		</tr>
		<tr>
			<td>Microglass coverslips for planar bilayers</td>
			<td>Matsunami</td>
			<td>Discontinued</td>
			<td>22x22</td>
		</tr>
		<tr>
			<td>Mini centrifuge</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Non-Functionalized Silica Microspheres</td>
			<td>Bangs Laboratories, Inc.</td>
			<td>Depends on size: SS0200*-SS0500*</td>
			<td>Silica in aqueous suspension</td>
		</tr>
		<tr>
			<td>Optical Adhesive</td>
			<td>Norland Thorlabs</td>
			<td>NOA 68</td>
			<td>Flexible adhesive for glass or plastics</td>
		</tr>
		<tr>
			<td>Osmium tetroxide</td>
			<td>Millipore Sigma</td>
			<td>20816-12-0</td>
			<td></td>
		</tr>
		<tr>
			<td>Parafilm</td>
			<td>VWR</td>
			<td>52858-000</td>
			<td></td>
		</tr>
		<tr>
			<td>Plasma Cleaner</td>
			<td>Plasma Etch</td>
			<td>PE-25</td>
			<td>Voltage: 120V, 60Hz. Current: 15 AMPS</td>
		</tr>
		<tr>
			<td>Potassium chloride</td>
			<td>VWR</td>
			<td>0395-1kg</td>
			<td></td>
		</tr>
		<tr>
			<td>Round coverglass, #1.5 12mm&#160;&#160;</td>
			<td>VWR</td>
			<td>64-0712</td>
			<td></td>
		</tr>
		<tr>
			<td>Sonicator bath</td>
			<td>Branson</td>
			<td>1510R-MT</td>
			<td>Bransonic Ultrasonic cleaner. 50-60 Hz. Output: 70W</td>
		</tr>
		<tr>
			<td>Soy PI</td>
			<td>Avanti Polar Lipids</td>
			<td>840044</td>
			<td></td>
		</tr>
		<tr>
			<td>Tabletop centrifuge</td>
			<td>Eppendorf</td>
			<td>22331</td>
			<td></td>
		</tr>
		<tr>
			<td>UV Lamp</td>
			<td>Spectroline</td>
			<td>ENF-260C</td>
			<td>115 Volts, 60 Hz, 0.20 AMPS</td>
		</tr>
		<tr>
			<td>WhatmanGlass Microfiber Filter Paper</td>
			<td>VWR</td>
			<td>28455-030</td>
			<td>42.5 mm diameter, Grade GF/C</td>
		</tr>
	</tbody>
</table>

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.

Following the preparation of each SLB, septins or the protein of interest may be incubated with the desired support and imaged via TIRFM, confocal microscopy, or SEM. The results shown here use septins recombinantly expressed and purified from E. coli17. Using TIRFM on planar SLBs, it is possible to determine the length of filaments and their flexibility, measure the diffusion coefficients and observe assembly over time28,29. In o...

Watch this Scientific Journal Video about Reconstitution of Septin Assembly at Membranes to Study Biophysical Properties and Functions at JoVE.com

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

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 ...

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Biochemistry

2.1K Views.  University of North Carolina at Chapel Hill. 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.

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

Isolation of Intermediate Filament Proteins from Multiple Mouse Tissues to Study Aging-associated Post-translational Modifications

Intermediate filaments (IFs), together with actin filaments and microtubules, form the cytoskeleton &#8211; a critical structural element of every cell. Normal functioning IFs provide cells with mechanical and stress resilience, while a dysfunctional IF cytoskeleton compromises cellular health and has been associated with many human diseases. Post-translational modifications (PTMs) critically regulate IF dynamics in response to physiological changes and under stress conditions. Therefore, the ability to monitor changes in the PTM signature of IFs can contribute to a better functional understanding, and ultimately conditioning, of the IF system as a stress responder during cellular injury. However, the large number of IF proteins, which are encoded by over 70 individual genes and expressed in a tissue-dependent manner, is a major challenge in sorting out the relative importance of different PTMs. To that end, methods that enable monitoring of PTMs on IF proteins on an organism-wide level, rather than for isolated members of the family, can accelerate research progress in this area. Here, we present biochemical methods for the isolation of the total, detergent-soluble, and detergent-resistant fraction of IF proteins from 9 different mouse tissues (brain, heart, lung, liver, small intestine, large intestine, pancreas, kidney, and spleen). We further demonstrate an optimized protocol for rapid isolation of IF proteins by using lysing matrix and automated homogenization of different mouse tissues. The automated protocol is useful for profiling IFs in experiments with high sample volume (such as in disease models involving multiple animals and experimental groups). The resulting samples can be utilized for various downstream analyses, including mass spectrometry-based PTM profiling. Utilizing these methods, we provide new data to show that IF proteins in different mouse tissues (brain and liver) undergo parallel changes with respect to their expression levels and PTMs during aging.

In this method, we present biochemical procedures for rapid and efficient isolation of intermediate filament (IF) proteins from multiple mouse tissues. Isolated IFs can be used to study changes in post-translational modifications by mass spectrometry and other biochemical assays.

Intermediate filaments (IFs), together with actin filaments and microtubules, form the cytoskeleton &#8211; a critical structural element of every cell. ...

Atomic Scale Structural Studies of Macromolecular Assemblies by Solid-state Nuclear Magnetic Resonance Spectroscopy

Supramolecular protein assemblies play fundamental roles in biological processes ranging from host-pathogen interaction, viral infection to the propagation of neurodegenerative disorders. Such assemblies consist in multiple protein subunits organized in a non-covalent way to form large macromolecular objects that can execute a variety of cellular functions or cause detrimental consequences. Atomic insights into the assembly mechanisms and the functioning of those macromolecular assemblies remain often scarce since their inherent insolubility and non-crystallinity often drastically reduces the quality of the data obtained from most techniques used in structural biology, such as X-ray crystallography and solution Nuclear Magnetic Resonance (NMR). We here present magic-angle spinning solid-state NMR spectroscopy (SSNMR) as a powerful method to investigate structures of macromolecular assemblies at atomic resolution. SSNMR can reveal atomic details on the assembled complex without size and solubility limitations. The protocol presented here describes the essential steps from the production of 13C/15N isotope-labeled macromolecular protein assemblies to the acquisition of standard SSNMR spectra and their analysis and interpretation. As an example, we show the pipeline of a SSNMR structural analysis of a filamentous protein assembly.

Structures of supramolecular protein assemblies at atomic resolution are of high relevance because of their crucial roles in a variety of biological phenomena. Herein, we present a protocol to perform high-resolution structural studies on insoluble and non-crystalline macromolecular protein assemblies by magic-angle spinning solid-state nuclear magnetic resonance spectroscopy (MAS SSNMR).

Supramolecular protein assemblies play fundamental roles in biological processes ranging from host-pathogen interaction, viral infection to the ...

Characterization of Glycoproteins with the Immunoglobulin Fold by X-Ray Crystallography and Biophysical Techniques

Glycoproteins on the surface of cells play critical roles in cellular function, including signalling, adhesion and transport. On leukocytes, several of these glycoproteins possess immunoglobulin (Ig) folds and are central to immune recognition and regulation. Here, we present a platform for the design, expression and biophysical characterization of the extracellular domain of human B cell receptor CD22. We propose that these approaches are broadly applicable to the characterization of mammalian glycoprotein ectodomains containing Ig domains. Two suspension human embryonic kidney (HEK) cell lines, HEK293F and HEK293S, are used to express glycoproteins harbouring complex and high-mannose glycans, respectively. These recombinant glycoproteins with different glycoforms allow investigating the effect of glycan size and composition on ligand binding. We discuss protocols for studying the kinetics and thermodynamics of glycoprotein binding to biologically relevant ligands and therapeutic antibody candidates. Recombinant glycoproteins produced in HEK293S cells are amenable to crystallization due to glycan homogeneity, reduced flexibility and susceptibility to endoglycosidase H treatment. We present methods for soaking glycoprotein crystals with heavy atoms and small molecules for phase determination and analysis of ligand binding, respectively. The experimental protocols discussed here hold promise for the characterization of mammalian glycoproteins to give insight into their function and investigate the mechanism of action of therapeutics.

We present approaches for the biophysical and structural characterization of glycoproteins with the immunoglobulin fold by biolayer interferometry, isothermal titration calorimetry, and X-ray crystallography.

Glycoproteins on the surface of cells play critical roles in cellular function, including signalling, adhesion and transport. On leukocytes, several of ...

Purification and Reconstitution of TRPV1 for Spectroscopic Analysis

Polymodal ion channels transduce multiple stimuli of different natures into allosteric changes; these dynamic conformations are challenging to determine and remain largely unknown. With recent advances in single-particle cryo-electron microscopy (cryo-EM) shedding light on the structural features of agonist binding sites and the activation mechanism of several ion channels, the stage is set for an in-depth dynamic analysis of their gating mechanisms using spectroscopic approaches. Spectroscopic techniques such as electron paramagnetic resonance (EPR) and double electron-electron resonance (DEER) have been mainly restricted to the study of prokaryotic ion channels that can be purified in large quantities. The requirement for large amounts of functional and stable membrane proteins has hampered the study of mammalian ion channels using these approaches. EPR and DEER offer many advantages, including determination of the structure and dynamic changes of mobile protein regions, albeit at low resolution, that might be difficult to obtain by X-ray crystallography or cryo-EM, and monitoring reversible gating transition (i.e., closed, open, sensitized, and desensitized). Here, we provide protocols for obtaining milligrams of functional detergent-solubilized transient receptor potential cation channel subfamily V member 1 (TRPV1) that can be labeled for EPR and DEER spectroscopy.

This article describes specific methods to obtain biochemical quantities of detergent-solubilized TRPV1 for spectroscopic analysis. The combined protocols provide biochemical and biophysical tools that can be adapted to facilitate structural and functional studies for mammalian ion channels in a membrane-controlled environment.

Polymodal ion channels transduce multiple stimuli of different natures into allosteric changes; these dynamic conformations are challenging to determine ...

Using In Vitro Fluorescence Resonance Energy Transfer to Study the Dynamics Of Protein Complexes at a Millisecond Time Scale

Proteins are the primary operators of biological systems, and they usually interact with other macro- or small molecules to carry out their biological functions. Such interactions can be highly dynamic, meaning the interacting subunits are constantly associated and dissociated at certain rates. While measuring the binding affinity using techniques such as quantitative pull-down reveals the strength of the interaction, studying the binding kinetics provides insights on how fast the interaction occurs and how long each complex can exist. Furthermore, measuring the kinetics of an interaction in the presence of an additional factor, such as a protein exchange factor or a drug, helps reveal the mechanism by which the interaction is regulated by the other factor, providing important knowledge for the advancement of biological and medical research. Here, we describe a protocol for measuring the binding kinetics of a protein complex that has a high intrinsic association rate and can be dissociated quickly by another protein. The method uses fluorescence resonance energy transfer to report the formation of the protein complex in vitro, and it enables monitoring the fast association and dissociation of the complex in real time on a stopped-flow fluorimeter. Using this assay, the association and dissociation rate constants of the protein complex are quantified.

Protein-protein interactions are critical for biological systems, and studies of the binding kinetics provide insights into the dynamics and function of protein complexes. We describe a method that quantifies the kinetic parameters of a protein complex using fluorescence resonance energy transfer and the stopped-flow technique. &#160;

Proteins are the primary operators of biological systems, and they usually interact with other macro- or small molecules to carry out their biological ...

A Planarian Motility Assay to Gauge the Biomodulating Properties of Natural Products

A straightforward, controllable means of using the non-parasitic planarian, Dugesia tigrina, a free-living aquatic flatworm, to study the stimulant and withdrawal properties of natural products is described. Experimental assays benefitting from unique aspects of planarian physiology have been applied to studies on wound healing, regeneration, and tumorigenesis. In addition, because planarians exhibit sensitivity to a variety of environmental stimuli and are capable of learning and developing conditioned responses, they can be used in behavioral studies examining learning and memory. Planarians possess a basic bilateral symmetry and a central nervous system that uses neurotransmitter systems amenable to studies examining the effects of neuromuscular biomodulators. Consequently, experimental systems monitoring planarian movement and motility have been developed to examine substance addiction and withdrawal. Because planarian motility offers the potential for a sensitive, easily standardized motility assay system to monitor the effect of stimuli, the planarian locomotor velocity (pLmV) test was adapted to monitor both stimulation and withdrawal behaviors by planarians through the determination of the number of grid lines crossed by the animals with time. Here, the technique and its application are demonstrated and explained.

Planarian motility is used to gauge the stimulant and withdrawal properties of natural products when compared to the movement of the animals in spring water alone.

A straightforward, controllable means of using the non-parasitic planarian, Dugesia tigrina, a free-living aquatic flatworm, to study the stimulant and ...

Co-Translational Insertion of Membrane Proteins into Preformed Nanodiscs

Cell-free expression systems allow the tailored design of reaction environments to support the functional folding of even complex proteins such as membrane proteins. The experimental procedures for the co-translational insertion and folding of membrane proteins into preformed and defined membranes supplied as nanodiscs are demonstrated. The protocol is completely detergent-free and can generate milligrams of purified samples within one day. The resulting membrane protein/nanodisc samples can be used for a variety of functional studies and structural applications such as crystallization, nuclear magnetic resonance, or electron microscopy. The preparation of basic key components such as cell-free lysates, nanodiscs with designed membranes, critical stock solutions as well as the assembly of two-compartment cell-free expression reactions is described. Since folding requirements of membrane proteins can be highly diverse, a major focus of this protocol is the modulation of parameters and reaction steps important for sample quality such as critical basic reaction compounds, membrane composition of nanodiscs, redox and chaperone environment, or DNA template design. The whole process is demonstrated with the synthesis of proteorhodopsin and a G-protein coupled receptor.

Co-translational insertion into pre-formed nanodiscs makes it possible to study cell-free synthesized membrane proteins in defined lipid environments without contact with detergents. This protocol describes the preparation of essential system components and the critical parameters for improving expression efficiency and sample quality.

Cell-free expression systems allow the tailored design of reaction environments to support the functional folding of even complex proteins such as ...

High-Resolution Neutron Spectroscopy to Study Picosecond-Nanosecond Dynamics of Proteins and Hydration Water

Neutron scattering offers the possibility to probe the dynamics within samples for a wide range of energies in a nondestructive manner and without labeling other than deuterium. In particular, neutron backscattering spectroscopy records the scattering signals at multiple scattering angles simultaneously and is well suited to study the dynamics of biological systems on the ps-ns timescale. By employing D2O-and possibly deuterated buffer components-the method allows monitoring of both center-of-mass diffusion and backbone and side-chain motions (internal dynamics) of proteins in liquid state.
Additionally, hydration water dynamics can be studied by employing powders of perdeuterated proteins hydrated with H2O. This paper presents the workflow employed on the instrument IN16B at the Institut Laue-Langevin (ILL) to investigate protein and hydration water dynamics. The preparation of solution samples and hydrated protein powder samples using vapor exchange is explained. The data analysis procedure for both protein and hydration water dynamics is described for different types of datasets (quasielastic spectra or fixed-window scans) that can be obtained on a neutron backscattering spectrometer.
The method is illustrated with two studies involving amyloid proteins. The aggregation of lysozyme into &#181;m sized spherical aggregates-denoted particulates-is shown to occur in a one-step process on the space and time range probed on IN16B, while the internal dynamics remains unchanged. Further, the dynamics of hydration water of tau was studied on hydrated powders of perdeuterated protein. It is shown that translational motions of water are activated upon the formation of amyloid fibers. Finally, critical steps in the protocol are discussed as to how neutron scattering is positioned regarding the study of dynamics with respect to other experimental biophysical methods.

Neutron backscattering spectroscopy offers a nondestructive and label-free access to the ps-ns dynamics of proteins and their hydration water. The workflow is presented with two studies on amyloid proteins: on the time-resolved dynamics of lysozyme during aggregation and on the hydration water dynamics of tau upon fiber formation.

Neutron scattering offers the possibility to probe the dynamics within samples for a wide range of energies in a nondestructive manner and without ...

Purification and Quality Control of Recombinant Septin Complexes for Cell-Free Reconstitution

Septins are a family of conserved eukaryotic GTP-binding proteins that can form cytoskeletal filaments and higher-order structures from hetero-oligomeric complexes. They interact with other cytoskeletal components and the cell membrane to participate in important cellular functions such as migration and cell division. Due to the complexity of septins' many interactions, the large number of septin genes (13 in humans), and the ability of septins to form hetero-oligomeric complexes with different subunit compositions, cell-free reconstitution is a vital strategy to understand the basics of septin biology. The present paper first describes a method to purify recombinant septins in their hetero-oligomeric form using a two-step affinity chromatography approach. Then, the process of quality control used to check for the purity and integrity of the septin complexes is detailed. This process combines native and denaturing gel electrophoresis, negative stain electron microscopy, and interferometric scattering microscopy. Finally, a description of the process to check for the polymerization ability of septin complexes using negative stain electron microscopy and fluorescent microscopy is given. This demonstrates that it is possible to produce high-quality human septin hexamers and octamers containing different isoforms of septin_9, as well as Drosophila septin hexamers.

In vitro reconstitution of cytoskeletal proteins is a vital tool to understand the basic functional properties of these proteins. The present paper describes a protocol to purify and assess the quality of recombinant septin complexes, which play a central role in cell division and migration.

Septins are a family of conserved eukaryotic GTP-binding proteins that can form cytoskeletal filaments and higher-order structures from hetero-oligomeric ...

Using Microfluidics and Fluorescence Microscopy to Study the Assembly Dynamics of Single Actin Filaments and Bundles

In order to decipher the complex molecular mechanisms that regulate the assembly and disassembly of actin filaments, it is a great asset to monitor individual reactions live in well-controlled conditions. To do so, live single-filament experiments have emerged over the past 20 years, mostly using total internal reflection fluorescence (TIRF) microscopy, and have provided a trove of key results. In 2011, in order to further expand the possibilities of these experiments and to avoid recurring problematic artifacts, we introduced simple microfluidics in these assays. This study details our basic protocol, where individual actin filaments are anchored by one end to the passivated coverslip surface, align with the flow, and can be successively exposed to different protein solutions. We also present the protocols for specific applications and explain how controlled mechanical forces can be applied, thanks to the viscous drag of the flowing solution. We highlight the technical caveats of these experiments and briefly present possible developments based on this technique. These protocols and explanations, along with today&#39;s availability of easy-to-use microfluidics equipment, should allow non-specialists to implement this assay in their labs.

We present protocols for simple actin filament microfluidic assays, in combination with fluorescence microscopy, that allow one to accurately monitor individual actin filaments in real-time while sequentially exposing them to different protein solutions.

In order to decipher the complex molecular mechanisms that regulate the assembly and disassembly of actin filaments, it is a great asset to monitor ...