We thank Nanyang NanoFabrication Centre (N2FC) and the Centre for Disruptive Photonic Technologies (CDPT) at Nanyang Technological University (NTU) for supporting nanostructure fabrication and SEM imaging, the Protein Production Platform (PPP) at the School of Biological Sciences NTU for protein purification, and the School of Chemical and Biomedical Engineering NTU for the confocal microscope. This work is funded by the Singapore Ministry of Education (MOE) (W. Zhao, RG112/20, RG95/21, and MOE-T2EP30220-0009), the Institute for Digital Molecular Analytics and Science (IDMxS) supported by MOE funding under the Research Centres of Excellence scheme (W. Zhao), the Human Frontier Science Program Foundation (W. Zhao, RGY0088/2021), the NTU Start-up Grant (W. Zhao), School of Chemical and Biomedical Engineering NTU for the research scholarship (X. Miao), and China Scholarship Council for the research scholarship (J. Wu).

1. Cleaning of nanochip
<ol>
	<li>Place the nanochip in a 10 mL beaker with the patterned side facing up. 
	NOTE: This quartz nanochip has been fabricated via electron beam lithography as described before21. The geometry and arrangement of the nanostructure on the chip can be custom designed. The sizes of the gradient nanobars used here are 2000 nm in length, 600 nm in height, and 100 to 1000 nm in width (100 nm step-set).</li>
	<li>Carefully add 1 mL of 98% sulfuric acid to the beaker, and ensure the acid fully covers the front and backside of the chip. 
	NOTE: 98% sulfuric acid is extremely corrosive and can cause rapid tissue destruction and serious chemical burns upon contact with the skin or eyes. Use it in the fume hood with proper protection.</li>
	<li>Slowly rotate the beaker and add 200 &#181;L of 30% hydrogen peroxide drop by drop until the whole beaker becomes hot. Ensure that sulfuric acid and hydrogen peroxide are well mixed to form piranha solution for the removal of organic molecules from the nanochip17. There are alternative techniques for generating the SLB on clean hydrophilic surfaces, such as UV light and ozone exposure as described earlier23. 
	NOTE: The reaction is extremely exothermic and can cause the solution to boil, so add hydrogen peroxide to the sulfuric acid drop by drop and keep rotating.</li>
	<li>Place the beaker in a secondary glass container and keep the nanochip immersed in the piranha solution overnight to clean the impurities thoroughly. 
	NOTE: Place the beaker in the fume hood without any cover in case the reaction can generate gas.</li>
	<li>Take the beaker out and carefully pipette the piranha solution into an acid waste container.</li>
	<li>Load 5 mL of deionized water into the beaker to dilute the residual acid and discard it into the acid waste. Repeat this step five times and use 5 M NaOH to neutralize the acid waste.</li>
	<li>Grab the chip with tweezers and wash with a continuous stream of deionized water to remove residual acid thoroughly. Blow-dry the chip with 99.9% nitrogen gas for SLB formation22.</li>
</ol>
2. Generation of small unilamellar vesicles (SUVs)
<ol>
	<li>Add 100 &#181;L of chloroform into an amber vial. 
	NOTE: Chloroform is a highly volatile, colorless liquid, and it can be toxic if inhaled or swallowed. Use it in the fume hood with the mask and gloves on.</li>
	<li>Dissolve 500 &#181;g of the lipid mixture in chloroform. The composition of the lipid mixture used here is 89.5 mol% of egg phosphatidylcholines (PC), 10 mol% of brain phosphatidylserine (PS), and 0.5 mol% of Texas Red 1,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine, triethylammonium salt (Texas Red DHPE).</li>
	<li>Blow-dry the lipid mixture in the vial with 99.9% nitrogen gas until the chloroform is volatilized and the lipid mixture is dry. 
	NOTE: This step should be performed in the fume hood. Avoid liquid spatter when blow-drying.</li>
	<li>Place the lipid mixture present in the amber vial without the lid in the aluminum foil-covered vacuum desiccator. Then vacuum dry the sample with a pump for 3 h to completely volatilize the chloroform.</li>
	<li>Add 250 &#181;L of phosphate-buffered saline (PBS) buffer to the vial. Vortex the solution until it is homogeneous. 
	NOTE: PBS buffer consists of 150 mM NaCl, without calcium, magnesium, and phenol red; the pH is adjusted to 7.2 for making the PC and PS lipid bilayer.</li>
	<li>Sonicate the lipid mixture for 30 min at a frequency of 50 kHz in a bath sonicator. Then transfer the lipid mixture into a 1.5 mL centrifuge tube and seal with parafilm.</li>
	<li>Freeze the lipid mixture in liquid nitrogen for 20 s and then thaw at 42 &#176;C for 2 min in a water bath. Repeat the freeze-thaw cycles 30 times. After that, the lipid mixture looks like a clear liquid. 
	CAUTION: Liquid nitrogen has a boiling point of about &#8722;195.8 &#730;C. It can cause frostbite or cryogenic burns. Use it in the unconfined space with heat insulation gloves on.</li>
	<li>Rinse two glass gas-tight syringes and the connector of the mini-extruder with ethanol, chloroform, and methanol respectively. Repeat the rinsing sequence five times to pre-clean the apparatus. Leave them in the fume hood until the organic solvent is completely volatilized.</li>
	<li>Assemble the mini-extruder apparatus and pass 500 &#181;L of PBS buffer through the connector to pre-wet the apparatus, then discard the buffer from another syringe. This step removes impurities and facilitates extrusion.</li>
	<li>Assemble the mini-extruder apparatus with a 100 nm pore-sized polycarbonate filter membrane. 
	NOTE: The pore size of the filter membrane determines the diameter of liposomes.</li>
	<li>Take 500 &#181;L of PBS buffer with one of the syringes and gently push it through the filter to fill the second empty syringe at the other end. Repeat the extrusion back and forth three times to check the apparatus leakage and discard the buffer from the second syringe.</li>
	<li>Pass the lipid mixture through the mini extruder to replace the PBS buffer. Then extrude back and forth 20 times to form SUVs. The multilamellar character of the vesicles may be retained with insufficient number of extrusion times, which will affect SLB formation23.</li>
	<li>Collect the SUVs from the second syringe to reduce the contamination with larger particles and transfer it into a 1.5 mL centrifuge tube. 
	NOTE: Remove the syringe straight out of the connector in case the syringe breaks off.</li>
	<li>Wrap the tube with aluminum foil to prevent bleaching of fluorescent-labeled lipids and store them at 4 &#730;C. Usually, the SUVs can be stored for up to 7 days.</li>
</ol>
3. Formation of the SLB on a nanochip
<ol>
	<li>Take out the cleaned nanochip from deionized water carefully with a pair of tweezer and blow-dry with 99.9% nitrogen gas.</li>
	<li>Perform surface cleaning of the nanochip with air plasma treatment for 1 h.</li>
	<li>Assemble the nanochip in a polydimethylsiloxane (PDMS) chamber, which consists of two pieces of PDMS-a middle PDMS and a top PDMS (Figure 1A). The middle PDMS is thin (~0.5 mm) with a large oval-shaped opening in the center to ensure sufficient exposure to the nanobar pattern. The top PDMS is thick (~4.0 mm) with two small holes within the large oval opening as an inlet and an outlet for liquid handling.
	<ol>
		<li>Place the chip on a clean surface with the pattern facing up.</li>
		<li>Gently cover the middle PDMS with the chip, and make sure that the whole pattern is exposed to the central area of the large oval-shaped opening in the middle PDMS.</li>
		<li>Cover the top PDMS with the middle PDMS and keep its two small holes within the region of the large oval hole of the middle PDMS. Then the PDMS chamber is assembled. 
		NOTE: PDMS is the most widely used silicon-based organic polymer. It is biocompatible, transparent, as well as deformable to be designed as a customized shape for chip assembly and imaging under the microscope. More importantly, it can be covalently stuck to another PDMS layer or glass tightly after plasma treatment, making it suitable as the chamber to prepare the SLB. This step ensures that each layer of PDMS is tightly fitted to avoid gaps and leakage.</li>
	</ol>
	</li>
	<li>Load the SUVs into the PDMS chamber from one of the two small holes in the top PDMS with a pipette and incubate for 15 min at room temperature to form the SLB.</li>
	<li>Gently pipette the PBS buffer into the PDMS chamber from one side of the small hole and remove the waste with a cotton bud from the other hole to wash away the unbound SUVs. Then acquire the SLB formed on the nanochip (the quality of the SLB is tested by fluorescence recovery after photobleaching (FRAP) as shown in step 4.4 and discussed in the representative results section). 
	&#8203;NOTE: When pipetting the PBS buffer into the chamber, the pipette tip should be full of the buffer and in contact with the liquid surface to avoid injecting any bubbles.</li>
</ol>
4. Imaging the SLB and the curvature sensing protein binding on the nanochip
NOTE: This section will depend on the microscope system available for the experiment. Here, an overall guideline on how to perform the imaging will be described. The detailed settings can be changed between the different microscope setups.
<ol>
	<li>Set up the laser scanning confocal microscopy using a 100x (N.A.1.4) oil objective. Open the ZEN software to select the excitation laser power that can excite the fluorescence of the lipid and protein. Choose Acquisition mode &#62; Smart Setup &#62; Texas Red DHPE / EGFP.</li>
	<li>Adjust the focus with the focus knob to locate the nanobars on the chip until the nanobar edges are sharp under the lipid channel.</li>
	<li>Set the scanning parameters as below to obtain a control image of the lipid channel before adding protein: Frame Size = 512 px x 512 px (512 pixels x 512 pixels, a pixel size of 124 nm), Bits per Pixel = 16 (16-bit depth), Scan speed = 5, and Averaging = 4 (averaging mode of four times/line).</li>
	<li>Conduct the FRAP assay by bleaching the fluorescent-labeled lipid bilayer on a random single nanobar area.
	<ol>
		<li>Select the Experiment Regions and Bleaching checkboxes. Draw a circular area of 5 &#181;m diameter which can include the whole nanobar at the center (nanobar size is 2 &#181;m) and add to the Experiment Regions. Input time-lapse imaging parameters as the following example: choose Scan speed = 9 and Averaging = 1. Choose Time Series &#62; Duration = 100 cycles and Interval = 2 s. Input bleaching parameters as the following example: select the Start after # images checkbox and choose three images.</li>
		<li>Select the laser checkboxes that perform FRAP and change the power to 100.0%. Click Start Experiment for the FRAP experiment. 
		NOTE: FRAP is a common method to characterize the mobility of cellular molecules. If the SLB has good fluidity, the fluorescence in the bleached area will gradually recover as bleached fluorophores diffuse out and unbleached fluorophores from other areas diffuse in.</li>
	</ol>
	</li>
	<li>Load the protein solution into the PDMS chamber and incubate for 5 min at room temperature to allow the binding of the protein on the SLB. 
	NOTE: The proteins used in Figure 2G,H to demonstrate lipid composition facilitate protein binding on nanobar-curved SLBs are 74 &#181;M GFP and 79 &#181;M GFP-His. These two proteins are green fluorescence proteins without or with His-tag. The proteins used for curvature sensing study in Figure 4 and Figure 5 are 16 &#181;M F-BAR, 16 &#181;M IDRFBP17, and 16 &#181;M FBAR+IDRFBP17. These three proteins are the domains from FBP17, which is a typical BAR protein that is intuitively assumed as both curvature generators and sensors. Each protein volume is 20 &#181;L.</li>
	<li>Re-focus the nanobars and repeat step 4.3 to take images of both lipid and protein channels for protein curvature sensing detection.</li>
	<li>Repeat step 4.4 to conduct the FRAP assay on both lipid and protein channels and perform time-lapse imaging to characterize the mobility of curvature sensing protein.</li>
</ol>
5. Reuse of nanochip
<ol>
	<li>Grab the nanochip with tweezers and carefully detach it from the PDMS chamber in a 50 mL beaker filled with deionized water. 
	NOTE: This is a critical step. Prevent the tweezers from touching the nanostructure, and do not force the chip to separate to avoid cracks.</li>
	<li>Wash the nanochip thoroughly with anhydrous ethanol to remove organic residues attached on the surface. 
	NOTE: Use clean tissue paper to remove water on the surface before washing with anhydrous ethanol. 
	CAUTION: Anhydrous ethanol is a highly volatile and slightly hazardous chemical. Avoid contact with the skin and eyes. Use it in the fume hood with the mask and gloves on.</li>
	<li>Wash the chip thoroughly with plenty of deionized water to remove residual ethanol. Then blow-dry the chip with 99.9% nitrogen gas. 
	NOTE: It is important to remove all traces of ethanol on the chip before proceeding to the next step because the piranha acid can react violently with organic solvent and may cause explosions.</li>
	<li>Place the chip in a clean 10 mL beaker with the pattern side facing up and clean the chip with piranha solution for reuse which follows the same steps as &#39;cleaning of nanochip&#39;.</li>
</ol>
6. Image quantification
<ol>
	<li>Rotate the images acquired by a microscope so that the nanobar array is in a vertical position for subsequent analysis in Fiji software.</li>
	<li>Use a custom-written MATLAB code &#39;c_pillarmask_averaging&#39; to locate the individual nanobar by a square mask (51 pixels x 51 pixels) both in the lipid channel and protein channel. 
	NOTE: This MATLAB code is specially designed for the nanochip. It can be obtained on GitHub&#160;via&#160;the link:&#160;https://github.com/wtzhaolab/GNB_SLB_MX.</li>
	<li>Subtract the background signals of each image with the three ROIs (5 pixels x 5 pixels) at the corners of the image by the meshgrid function in MATLAB code &#39;BarGra_avg&#39;. This operation aims to correct potential uneven background noises caused by the microscope setup or chip leveling on the microscope stage.</li>
	<li>Generate the average images of the same-sized nanobars using MATLAB code &#39;BarGra_avg&#39;, where each point is the mean of the values at that point in all the individual nanobar square masks. Generate 3D surface plots of the average images in Fiji software to show the average signal distribution around the nanobars intuitively. Choose Analyze &#62; Surface Plot &#62; Input 100 for Polygon Multiplier, and select Shade, Draw Axis, and Smooth checkboxes.</li>
	<li>Segment each nanobar into three areas, which include two nanobar-end areas and one nanobar-center area to extract their fluorescence intensities respectively by MATLAB code &#39;avg_nanobar_quantification&#39;. The sizes of the nanobar ROIs are adjusted according to the dimension of the nanobars using the &#39;position&#39; file.</li>
	<li>Divide the protein intensities by lipid intensities extracted from the MATLAB code &#39;avg_nanobar_quantification&#39; at the bar-end area to get the nanobar-end density which excludes the surface area effect.</li>
	<li>Plot the nanobar-end density with different concentrations in GraphPad Prism to get the binding curve. Open the Analyze menu. Choose Nonlinear regression (curve fit) &#62; Binding - Saturation &#62; Specific binding with Hill slope function to fit the curve with the Hill equation and then calculate the KD and Hill coefficient value.</li>
	<li>The lipid and protein intensities are normalized to their corresponding intensities at the 600 nm nanobars center acquired in the same image using the MATLAB code &#39;avg_nanobar_quantification&#39;.</li>
	<li>Use the brightest nine pixels of normalized protein intensities at the nanobar-end area divided by the bar-center area to quantify the protein curvature sensing with same-sized nanobars, the value is named as &#34;end-to-center ratio&#34;. Use the ratio of normalized protein intensity to normalize lipid intensity to quantify the protein curvature sensing range with different-sized nanobars, the value is named as &#34;Normalized Nanobar-End Density&#34;. These values are calculated by the MATLAB code &#39;avg_nanobar_quantification&#39;.</li>
	<li>Load the FRAP stack image in Fiji software. Choose the FRAP area and generate an ROI. Choose the More &#62; Multi measure function to measure the intensity of the FRAP area for each time. Plot the intensity in GraphPad Prism to generate the FRAP recovery curve.</li>
</ol>

The authors declare no competing financial interest in this work.

The nanobar-SLB system described here offers a unique combination of the advantages in several existing in vitro assays. It efficiently reveals the preferential binding of proteins to highly curved membranes as the liposome floatation or sedimentation assay but requires much fewer samples and offers more accurately defined curvature on individual nanobars8,29. It also offers a wide range of precisely controlled curvature for simultaneous comparison as th...

Membrane curvature is a critical physical parameter of a cell that occurs in a variety of cellular processes such as morphogenesis, cell division, and cell migration1. It is widely recognized now that membrane curvature is beyond a simple result of cellular events; instead, it has emerged as an effective regulator of protein interactions and intracellular signaling. For example, several proteins involved in clathrin-mediated endocytosis were found to preferentially bind to the curved membrane, resulting in the formation of a hotspot for endocytosis2. There are many different causes of membrane deformation such as membrane pulling by the cytoskeletal forces, the presence of lipid asymmetry with different sized head groups, the existence of transmembrane proteins with conical shape, the accumulation of membrane-shaping proteins like BAR-domain proteins (named after Bin, amphiphysin, and Rvs proteins), and the insertion of amphipathic helices domain into the membrane1. Interestingly, these proteins and lipids not only deform the membrane but can also sense the membrane curvature and exhibit preferential accumulation1. Therefore, it is crucial to study whether and how membranes with different curvatures alter the distribution and dynamics of proteins and lipids attached to them and the potential impacts on the related intracellular processes.
Many techniques have been developed to analyze the interaction between curved membrane and proteins in both live cell and in vitro systems. The live cell system provides a real cell environment with rich lipid diversity and dynamic protein signaling regulation2,3,4,5,6,7. However, such a sophisticated system is difficult to study due to the uncertainties and fluctuations in the intracellular environment. Hence, the in vitro assays using an artificial membrane composed of known lipid species and purified proteins have become powerful reconstitution systems to characterize the relationship between proteins and curved membranes. Traditionally, liposomes of different diameters are generated by extrusion to detect curvature-sensitive proteins via either a co-sedimentation assay using centrifugal force or a co-flotation assay with a density gradient to avoid protein aggregation8,9. However, the curvature of the extruded liposomes is limited by the available pore size of the membrane filter used in the extruder10. Single liposome curvature (SLiC) assay has been proven to overcome this limitation, in which liposomes with different diameters are fluorescence-labeled and immobilized onto the surface so that the curvature can be marked by the fluorescent intensity11. However, strong variability in lipid composition has been observed in small vesicles, which affects the accuracy of the curvature measurement12. Tether-pulling experiments provide a more accurate measurement of the curvature on the transient tether pulled from giant unilamellar vesicles (GUVs) using an optical tweezer, where the curvature can be well controlled by the membrane tension generated13,14. This method is suitable to study either positive- or negative-curvature sensing proteins, but is constrained by the throughput of tube generation10. Supported membrane tubes (SMrT) assay affords simultaneous generation of multiple membrane tubes that are extruded from the same lipid reservoir by microfluidic flows. Nevertheless, the membrane curvature varies intrinsically along the nanotube, which compromises the accuracy of fluorescence-intensity-based curvature measurement15,16. In comparison, using small unilamellar vesicles (SUVs, diameter &#60;100 nm17) to form a supported lipid bilayer (SLB) on surfaces containing designed topographies generated a single bilayer membrane with curvatures predetermined by nanofabrication or nanomaterials in high accuracy18,19,20.
Here, we present a protocol for the formation of the SLB on fabricated nanochip surfaces with nanobar arrays and how it can be used to probe the curvature sensitivity of proteins in vitro. As shown in Figure 1, there are six essential components of the assay: A) Cleaning and assembly of the chip with a microfluidic chamber; B) Preparation of SUVs&#160;with defined lipid composition; C) Formation of the SLB on a nanochip and binding with curvature sensitive proteins; D) Imaging and characterization of the SLB and curvature sensitive proteins under fluorescence microscopy; E) Cleaning the chip for reuse; F) Image processing for quantitative analysis of protein curvature sensing ability. The detailed protocol is described step-by-step below.

<table><tbody><tr><td>Anhydrous Ethanol</td><td>Sigma-Aldrich</td><td>100983</td><td></td></tr><tr><td>Aluminum foil</td><td>Diamond</td><td>RN0879999FU</td><td></td></tr><tr><td>Amber Vial</td><td>Sigma-Aldrich</td><td>27115-U</td><td></td></tr><tr><td>Brain PS: L-&amp;alpha;-phosphatidylserine (Brain, Porcine) (sodium salt)</td><td>Avanti Polar Lipids, Inc.</td><td>840032</td><td></td></tr><tr><td>10 mL Beaker</td><td>Schott-Duran</td><td>SCOT211060804</td><td></td></tr><tr><td>50 mL Beaker</td><td>Schott-Duran</td><td>SCOT211061706</td><td></td></tr><tr><td>1000 mL Beaker</td><td>Schott-Duran</td><td>SCOT211065408</td><td>The second container&amp;nbsp;</td></tr><tr><td>Chloroform</td><td>Sigma-Aldrich</td><td>V800117</td><td></td></tr><tr><td>Cotton buds</td><td>Watsons</td><td></td><td></td></tr><tr><td>18:1 DGS-NTA(Ni): 1,2-dioleoyl-sn-glycero-3-[(N-(5-amino-1-carboxypentyl)iminodiacetic acid)succinyl] (nickel salt)</td><td>Avanti Polar Lipids, Inc.</td><td>790404</td><td></td></tr><tr><td>Egg PC: L-&amp;alpha;-phosphatidylcholine (Egg, Chicken)</td><td>Avanti Polar Lipids, Inc.</td><td>840051</td><td></td></tr><tr><td>F-BAR</td><td>Protein Production Plaftorm, School of Biological Sciences, Nanyang Technological University, Singapore</td><td></td><td>Proteins and peptide</td></tr><tr><td>F-BAR+IDR</td><td>Protein Production Plaftorm, School of Biological Sciences, Nanyang Technological University, Singapore</td><td></td><td>Proteins and peptide</td></tr><tr><td>GFP</td><td>Protein Production Plaftorm, School of Biological Sciences, Nanyang Technological University, Singapore</td><td></td><td>Proteins and peptide</td></tr><tr><td>GFP-His</td><td>Protein Production Plaftorm, School of Biological Sciences, Nanyang Technological University, Singapore</td><td></td><td>Proteins and peptide</td></tr><tr><td>GraphPad Prism</td><td>GraphPad</td><td>V9.0.0</td><td></td></tr><tr><td>Hydrogen Peroxide, 30% (Certified ACS)</td><td>Thermo Scientific</td><td>H325-500</td><td></td></tr><tr><td>IDR from human FBP17</td><td>Sangon Biotech (Shanghai) Co., Ltd.</td><td></td><td></td></tr><tr><td>ImageJ</td><td>National Institutes of Health</td><td>1.50d</td><td></td></tr><tr><td>Laser Scanning Confocal Microscopy</td><td>Zeiss&amp;nbsp;</td><td>LSM 800 with Airyscan</td><td>100x (N.A.1.4) oil objective.</td></tr><tr><td>Methanol</td><td>Fisher scientific</td><td>10010240</td><td></td></tr><tr><td>Mini-extuder&amp;nbsp;</td><td>Avanti Polar Lipids, Inc.</td><td>610000-1EA</td><td></td></tr><tr><td>1.5 mL Microtubes</td><td>Greiner</td><td>616201</td><td></td></tr><tr><td>MATLAB</td><td>Mathworks</td><td>R2018b</td><td></td></tr><tr><td>Nuclepore Hydrophilic Membrane,0.1 &amp;mu;m</td><td>Whatman</td><td>800309</td><td></td></tr><tr><td>Phosphate Bufferen Saline (PBS)</td><td>Life Technologies Holdings Pte Ltd.</td><td>70013</td><td></td></tr><tr><td>Polydimethylsiloxane (PDMS) Base</td><td>Dow Corning Corporation</td><td>SYLGARD 184</td><td></td></tr><tr><td>Polydimethylsiloxane (PDMS) Crosslinker</td><td>Dow Corning Corporation</td><td>SYLGARD 184</td><td></td></tr><tr><td>Plasma Cleaner</td><td>HARRICK PLASMA</td><td>PDC-002-HP</td><td></td></tr><tr><td>Quartz Nanochip</td><td>Donghai County Alfa Quartz Products CO., LTD</td><td></td><td></td></tr><tr><td>Sodium Hydroxide&amp;nbsp;</td><td>Sigma-Aldrich</td><td>795429</td><td></td></tr><tr><td>Sulfuric acid</td><td>Sigma-Aldrich</td><td>258105</td><td></td></tr><tr><td>Texas Red DHPE: Texas Red 1,2-Dihexadecanoyl-sn-Glycero-3-Phosphoethanolamine, Triethylammonium Salt</td><td>Life Technologies Holdings Pte Ltd.</td><td>T1395MP</td><td></td></tr><tr><td>Tweezer</td><td>Gooi</td><td>PDC-002-HP</td><td></td></tr><tr><td>Ultrasonic Cleaners</td><td>Elma</td><td>D-78224</td><td></td></tr><tr><td>Voterx</td><td>Scientific Industries</td><td>G560E</td><td></td></tr><tr><td>Vacuum Desiccator</td><td>NUCERITE</td><td>5312</td><td></td></tr><tr><td>Water Bath</td><td>Julabo</td><td>TW8</td><td></td></tr></tbody></table>

a nanobar-supported lipid bilayer system for the study of membrane curvature sensing proteins in vitro

Membrane curvature plays important roles&#160;in various essential processes of cells, such as cell migration, cell division, and vesicle trafficking. It is not only passively generated by cellular activities, but also actively regulates protein interactions and is involved in many intracellular signaling. Thus, it is of great value to examine the role of membrane curvature in regulating the distribution and dynamics of proteins and lipids. Recently, many techniques have been developed to study the relationship between the curved membrane and protein in vitro. Compared to traditional techniques, the newly developed nanobar-supported lipid bilayer (SLB) offers both high-throughput and better accuracy in membrane curvature generation by forming a continuous lipid bilayer on patterned arrays of nanobars with a pre-defined membrane curvature and local flat control. Both the lipid fluidity and protein sensitivity to curved membranes can be quantitatively characterized using fluorescence microscopy imaging. Here, a detailed procedure on how to form a SLB on fabricated glass surfaces containing nanobar arrays&#160;and the characterization of curvature-sensitive proteins on such SLB are introduced. In addition, protocols for nanochip reusing and image processing are covered. Beyond the nanobar-SLB, this protocol is readily applicable to all types of nanostructured glass chips for curvature sensing studies.

Nanobar design is recommended for probing positive curvature sensing proteins, which contains a half circle at each end with curvature defined by the nanobar width and one flat/zero curvature control locally at the center (Figure 2A,B). Successful formation of the SLB on nanobars results in evenly distributed lipid marker signals across the entire nanobar surface as shown in Figure 2C. Signals from multiple nanobars can be combined by averaging ...

Watch this Scientific Journal Video about A Nanobar-Supported Lipid Bilayer System for the Study of Membrane Curvature Sensing Proteins in vitro at JoVE.com

A Nanobar-Supported Lipid Bilayer System for the Study of Membrane Curvature Sensing Proteins in vitro

Here, a nanobar-supported lipid bilayer system is developed to provide a synthetic membrane with a defined curvature that enables the characterization of proteins with curvature sensing ability in vitro.

A Nanobar-Supported Lipid Bilayer System for the Study of Membrane Curvature Sensing Proteins in vitro

Membrane curvature plays important roles&#160;in various essential processes of cells, such as cell migration, cell division, and vesicle trafficking. It ...

a-nanobar-supported-lipid-bilayer-system-for-study-membrane-curvature

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Biochemistry

2.8K Views.  Nanyang Technological University. Dit nanobar-ondersteunde lipide bilayer-systeem kan worden gebruikt om de rol van membraankromming te bestuderen bij het reguleren van de dynamiek en distributie van eiwitten en lipiden tijdens celactiviteiten. Deze techniek biedt zowel hoge subwortel als slechte in de membraankrommingsgeneratie door een continue lipide bilayer te vormen op patent nanobarfouten met membraankromming vooraf gedefinieerd door nanofabricage met hoge resolutie. Plaats om te beginnen de nanochip in een bekerglas va...

Video: A Nanobar-Supported Lipid Bilayer System for the Study of Membrane Curvature Sensing Proteins in vitro

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

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

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

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

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

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

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

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

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

Single Molecule Methods for Monitoring Changes in Bilayer Elastic Properties

Membrane protein function is regulated by the cell membrane lipid composition.  This regulation is due to a combination of specific lipid-protein interactions and more general lipid bilayer-protein interactions. These interactions are particularly important in pharmacological research, as many current pharmaceuticals on the market can alter the lipid bilayer material properties, which can lead to altered membrane protein function.   The formation of gramicidin channels are dependent on conformational changes in gramicidin subunits which are in turn dependent on the properties of the lipid.  Hence the gramicidin channel  current is a reporter of altered properties of the bilayer due to certain compounds.  

Membrane protein function is regulated by the cell membrane lipid composition. This video-article details how to form a patch using bilayer patch electrodes, as well as how to use gramicidin channels as reporters of altered membrane properties.

Membrane protein function is regulated by the cell membrane lipid composition.  This regulation is due to a combination of specific lipid-protein ...

Biology

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 ...

Bioengineering

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

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

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

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

A Quantitative Fluorescence Microscopy-based Single Liposome Assay for Detecting the Compositional Inhomogeneity Between Individual Liposomes

Most research employing liposomes as membrane model systems or drug delivery carriers relies on bulk read-out techniques and thus intrinsically assumes all liposomes of the ensemble to be identical. However, new experimental platforms able to observe liposomes at the single-particle level have made it possible to perform highly sophisticated and quantitative studies on protein-membrane interactions or drug carrier properties on individual liposomes, thus avoiding errors from ensemble averaging. Here we present a protocol for preparing, detecting, and analyzing single liposomes using a fluorescence-based microscopy assay, facilitating such single-particle measurements. The setup allows for imaging individual liposomes in a massive parallel manner and is employed to reveal intra-sample size and compositional inhomogeneities. Additionally, the protocol describes the advantages of studying liposomes at the single liposome level, the limitations of the assay, and the important features to be considered when modifying it to study other research questions.

This protocol describes the fabrication of liposomes and how these can be immobilized on a surface and imaged individually in a massive parallel manner using fluorescence microscopy. This allows for the quantification of the size and compositional inhomogeneity between single liposomes of the population.

Most research employing liposomes as membrane model systems or drug delivery carriers relies on bulk read-out techniques and thus intrinsically assumes ...

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

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

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

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

Pulling Membrane Nanotubes from Giant Unilamellar Vesicles

The reshaping of the cell membrane is an integral part of many cellular phenomena, such as endocytosis, trafficking, the formation of filopodia, etc. Many different proteins associate with curved membranes because of their ability to sense or induce membrane curvature. Typically, these processes involve a multitude of proteins making them too complex to study quantitatively in the cell. We describe a protocol to reconstitute a curved membrane in vitro, mimicking a curved cellular structure, such as the endocytic neck. A giant unilamellar vesicle (GUV) is used as a model of a cell membrane, whose internal pressure and surface tension are controlled with micropipette aspiration. Applying a point pulling force on the GUV using optical tweezers creates a nanotube of high curvature connected to a flat membrane. This method has traditionally been used to measure the fundamental mechanical properties of lipid membranes, such as bending rigidity. In recent years, it has been expanded to study how proteins interact with membrane curvature and the way they affect the shape and the mechanics of membranes. A system combining micromanipulation, microinjection, optical tweezers, and confocal microscopy allows measurement of membrane curvature, membrane tension, and the surface density of proteins, concurrently. From these measurements, many important mechanical and morphological properties of the protein-membrane system can be inferred. In addition, we lay out a protocol of creating GUVs in the presence of physiological salt concentration, and a method of quantifying the surface density of proteins on the membrane from fluorescence intensities of labeled proteins and lipids.

Many proteins in the cell sense and induce membrane curvature. We describe a method to pull membrane nanotubes from lipid vesicles to study the interaction of proteins or any curvature-active molecule with curved membranes in vitro.

The reshaping of the cell membrane is an integral part of many cellular phenomena, such as endocytosis, trafficking, the formation of filopodia, etc. Many ...

Bottom-Up In Vitro Methods to Assay the Ultrastructural Organization, Membrane Reshaping, and Curvature Sensitivity Behavior of Septins

Membrane remodeling occurs constantly at the plasma membrane and within cellular organelles. To fully dissect the role of the environment (ionic conditions, protein and lipid compositions, membrane curvature) and the different partners associated with specific membrane reshaping processes, we undertake in vitro bottom-up approaches. In recent years, there has been keen interest in revealing the role of septin proteins associated with major diseases. Septins are essential and ubiquitous cytoskeletal proteins that interact with the plasma membrane. They are implicated in cell division, cell motility, neuro-morphogenesis, and spermiogenesis, among other functions. It is, therefore, important to understand how septins interact and organize at membranes to subsequently induce membrane deformations and how they can be sensitive to specific membrane curvatures. This article aims to decipher the interplay between the ultra-structure of septins at a molecular level and the membrane remodeling occurring at a micron scale. To this end, budding yeast, and mammalian septin complexes were recombinantly expressed and purified. A combination of in vitro assays was then used to analyze the self-assembly of septins at the membrane. Supported lipid bilayers (SLBs), giant unilamellar vesicles (GUVs), large unilamellar vesicles (LUVs), and wavy substrates were used to study the interplay between septin self-assembly, membrane reshaping, and membrane curvature.

Bottom-Up In Vitro Methods to Assay the Ultrastructural Organization, Membrane Reshaping, and Curvature Sensitivity Behavior of Septins

Septins are cytoskeletal proteins. They interact with lipid membranes and can sense but also generate membrane curvature at the micron scale. We describe in this protocol bottom-up in vitro methodologies for analyzing membrane deformations, curvature-sensitive septin binding, and septin filament ultrastructure.

Membrane remodeling occurs constantly at the plasma membrane and within cellular organelles. To fully dissect the role of the environment (ionic ...

Spherical Supported Lipid Bilayer Assay: A Technique to Observe the Septin Assembly on Microspheres

Source: Curtis, B. N. et al., <a href="https://www.jove.com/v/64090">Reconstitution of Septin Assembly at Membranes to Study Biophysical Properties and Functions.</a> J. Vis. Exp. (2022).
This video describes the technique of studying the septin protein assembly on microspheres coated with lipid bilayers. This technique helps in understanding the curvature-dependent assemblies of protein on curved surfaces.

Source: Curtis, B. N. et al., Reconstitution of Septin Assembly at Membranes to Study Biophysical Properties and Functions. J. Vis. Exp. (2022).
This ...

A Nanobar-Supported Lipid Bilayer System for the Study of Membrane Curvature Sensing Proteins in vitro

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