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>

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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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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-&#945;-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&#160;</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-&#945;-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&#160;</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&#160;</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 &#956;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&#160;</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

Research

JoVE Journal

Biochemistry

2.7K Views.  Nanyang Technological University. 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.

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

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating these studies, reincorporation of detergent-solubilized membrane proteins into liposomes is a stochastic process where protein topology is impossible to enforce. This paper offers an alternative method to these challenging techniques that utilizes a liposome-based scaffold. Protein solubility is enhanced by deletion of the transmembrane domain, and these amino acids are replaced with a tethering moiety, such as a His-tag. This tether interacts with an anchoring group (Ni2+ coordinated by nitrilotriacetic acid (NTA(Ni2+)) for His-tagged proteins), which enforces a uniform protein topology at the surface of the liposome. An example is presented wherein the interaction between Dynamin-related protein 1 (Drp1) with an integral membrane protein, Mitochondrial Fission Factor (Mff), was investigated using this scaffold liposome method. In this work, we have demonstrated the ability of Mff to efficiently recruit soluble Drp1 to the surface of liposomes, which stimulated its GTPase activity. Moreover, Drp1 was able to tubulate the Mff-decorated lipid template in the presence of specific lipids. This example demonstrates the effectiveness of scaffold liposomes using structural and functional assays and highlights the role of Mff in regulating Drp1 activity.

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

This paper describes a method for assessing the interactions and assemblies of integral membrane proteins in vitro with various partner factors in a lipid-proximal environment.

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating ...

Nitrogen Cavitation and Differential Centrifugation Allows for Monitoring the Distribution of Peripheral Membrane Proteins in Cultured Cells

Cultured cells are useful for studying the subcellular distribution of proteins, including peripheral membrane proteins. Genetically encoded fluorescently tagged proteins have revolutionized the study of subcellular protein distribution. However, it is difficult to quantify the distribution with fluorescent microscopy, especially when proteins are partially cytosolic. Moreover, it is often important to study endogenous proteins. Biochemical assays such as immunoblots remain the gold standard for quantification of protein distribution after subcellular fractionation. Although there are commercial kits that aim to isolate cytosolic or certain membrane fractions, most of these kits are based on extraction with detergents, which may be unsuitable for studying peripheral membrane proteins that are easily extracted from membranes. Here we present a detergent-free protocol for cellular homogenization by nitrogen cavitation and subsequent separation of cytosolic and membrane-bound proteins by ultracentrifugation. We confirm the separation of subcellular organelles in soluble and pellet fractions across different cell types, and compare protein extraction among several common non-detergent-based mechanical homogenization methods. Among several advantages of nitrogen cavitation is the superior efficiency of cellular disruption with minimal physical and chemical damage to delicate organelles. Combined with ultracentrifugation, nitrogen cavitation is an excellent method to examine the shift of peripheral membrane proteins between cytosolic and membrane fractions.

Here we present protocols for detergent-free homogenization of cultured mammalian cells based on nitrogen cavitation and subsequent separation of cytosolic and membrane-bound proteins by ultracentrifugation. This method is ideal for monitoring the partitioning of peripheral membrane proteins between soluble and membrane fractions.

Cultured cells are useful for studying the subcellular distribution of proteins, including peripheral membrane proteins. Genetically encoded fluorescently ...

Detergent-free Ultrafast Reconstitution of Membrane Proteins into Lipid Bilayers Using Fusogenic Complementary-charged Proteoliposomes.

Detergents are indispensable for delivery of membrane proteins into 30-100 nm small unilamellar vesicles, while more complex, larger model lipid bilayers are less compatible with detergents.
Here we describe a strategy for bypassing this fundamental limitation using fusogenic oppositely charged liposomes bearing a membrane protein of interest. Fusion between such vesicles occurs within 5 min in a low ionic strength buffer. Positively charged fusogenic liposomes can be used as simple shuttle vectors for detergent-free delivery of membrane proteins into biomimetic target lipid bilayers, which are negatively charged. We also show how to reconstitute membrane proteins into fusogenic proteoliposomes with a fast 30-min protocol.
Combining these two approaches, we demonstrate a fast assembly of an electron transport chain consisting of two membrane proteins from E. coli, a primary proton pump bo3-oxidase and F1Fo ATP synthase, in membranes of vesicles of various sizes, ranging from 0.1 to &#62;10 microns, as well as ATP production by this chain.

Here we present two ultrafast protocols for reconstitution of membrane proteins into fusogenic proteoliposomes, and fusion of such proteoliposomes with target lipid bilayers for detergent-free delivery of these membrane proteins into the postfusion bilayer. The combination of these approaches enables fast and easily controlled assembly of complex multi-component membrane systems.

Detergents are indispensable for delivery of membrane proteins into 30-100 nm small unilamellar vesicles, while more complex, larger model lipid bilayers ...

Dissipative Microgravimetry to Study the Binding Dynamics of the Phospholipid Binding Protein Annexin A2 to Solid-supported Lipid Bilayers Using a Quartz Resonator

The dissipative quartz crystal microbalance technique is a simple and label-free approach to measure simultaneously the mass uptake and viscoelastic properties of the absorbed/immobilized mass on sensor surfaces, allowing the measurements of the interaction of proteins with solid-supported surfaces, such as lipid bilayers, in real-time and with a high sensitivity. Annexins are a highly conserved group of phospholipid-binding proteins that interact reversibly with the negatively charged headgroups via the coordination of calcium ions. Here, we describe a protocol that was employed to quantitatively analyze the binding of annexin A2 (AnxA2) to planar lipid bilayers prepared on the surface of a quartz sensor. This protocol is optimized to obtain robust and reproducible data and includes a detailed step-by-step description. The method can be applied to other membrane-binding proteins and bilayer compositions.

Here, we present an experimental protocol that can be employed to determine the binding affinities and mode of interaction of label-free phospholipid-binding protein annexin A2 with immobilized solid-supported bilayers (SLB) by simultaneously measuring the mass uptake and the viscoelastic properties of the protein annexin A2.

The dissipative quartz crystal microbalance technique is a simple and label-free approach to measure simultaneously the mass uptake and viscoelastic ...

A Fluorescence Fluctuation Spectroscopy Assay of Protein-Protein Interactions at Cell-Cell Contacts

A variety of biological processes involves cell-cell interactions, typically mediated by proteins that interact at the interface between neighboring cells. Of interest, only few assays are capable of specifically probing such interactions directly in living cells. Here, we present an assay to measure the binding of proteins expressed at the surfaces of neighboring cells, at cell-cell contacts. This assay consists of two steps: mixing of cells expressing the proteins of interest fused to different fluorescent proteins, followed by fluorescence fluctuation spectroscopy measurements at cell-cell contacts using a confocal laser scanning microscope. We demonstrate the feasibility of this assay in a biologically relevant context by measuring the interactions of the amyloid precursor-like protein 1 (APLP1) across cell-cell junctions. We provide detailed protocols on the data acquisition using fluorescence-based techniques (scanning fluorescence cross-correlation spectroscopy, cross-correlation number and brightness analysis) and the required instrument calibrations. Further, we discuss critical steps in the data analysis and how to identify and correct external, spurious signal variations, such as those due to photobleaching or cell movement.
In general, the presented assay is applicable to any homo- or heterotypic protein-protein interaction at cell-cell contacts, between cells of the same or different types and can be implemented on a commercial confocal laser scanning microscope. An important requirement is the stability of the system, which needs to be sufficient to probe diffusive dynamics of the proteins of interest over several minutes.

This protocol describes a fluorescence fluctuation spectroscopy-based approach to investigate interactions among proteins mediating cell-cell interactions, i.e. proteins localized in cell junctions, directly in living cells. We provide detailed guidelines on instrument calibration, data acquisition and analysis, including corrections to possible artefact sources.

A variety of biological processes involves cell-cell interactions, typically mediated by proteins that interact at the interface between neighboring ...

Single Liposome Measurements for the Study of Proton-Pumping Membrane Enzymes Using Electrochemistry and Fluorescent Microscopy

Proton-pumping enzymes of electron transfer chains couple redox reactions to proton translocation across the membrane, creating a proton-motive force used for ATP production. The amphiphilic nature of membrane proteins requires particular attention to their handling, and reconstitution into the natural lipid environment is indispensable when studying membrane transport processes like proton translocation. Here, we detail a method that has been used for the investigation of the proton-pumping mechanism of membrane redox enzymes, taking cytochrome bo3 from Escherichia coli as an example. A combination of electrochemistry and fluorescence microscopy is used to control the redox state of the quinone pool and monitor pH changes in the lumen. Due to the spatial resolution of fluorescent microscopy, hundreds of liposomes can be measured simultaneously while the enzyme content can be scaled down to a single enzyme or transporter per liposome. The respective single enzyme analysis can reveal patterns in the enzyme functional dynamics that might be otherwise hidden by the behavior of the whole population. We include a description of a script for automated image analysis.

Here, we present a protocol to study the molecular mechanism of proton translocation across lipid membranes of single liposomes, using cytochrome bo3 as an example. Combining electrochemistry and fluorescence microscopy, pH changes in the lumen of single vesicles, containing single or multiple enzyme, can be detected and analyzed individually.

Proton-pumping enzymes of electron transfer chains couple redox reactions to proton translocation across the membrane, creating a proton-motive force used ...

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 ...

Isolation of Lipoprotein Particles from Chicken Egg Yolk for the Study of Bacterial Pathogen Fatty Acid Incorporation into Membrane Phospholipids

Staphylococcus aureus and other Gram-positive pathogens incorporate fatty acids from the environment into membrane phospholipids. During infection, the majority of exogenous fatty acids are present within host lipoprotein particles. Uncertainty remains as to the reservoirs of host fatty acids and the mechanisms by which bacteria extract fatty acids from the lipoprotein particles. In this work, we describe protocols for enrichment of low-density lipoprotein (LDL) particles from chicken egg yolk and determining whether LDLs serve as fatty acid reservoirs for S. aureus. This method exploits unbiased lipidomic analysis and chicken LDLs, an effective and economical model for the exploration of interactions between LDLs and bacteria. The analysis of S. aureus integration of exogenous fatty acids from LDLs is performed using high-resolution/accurate mass spectrometry and tandem mass spectrometry, enabling the characterization of the fatty acid composition of the bacterial membrane and unbiased identification of novel combinations of fatty acids that arise in bacterial membrane lipids upon exposure to LDLs. These advanced mass spectrometry techniques offer an unparalleled perspective of fatty acid incorporation by revealing the specific exogenous fatty acids incorporated into the phospholipids. The methods outlined here are adaptable to the study of other bacterial pathogens and alternative sources of complex fatty acids.

This method provides a framework for studying incorporation of exogenous fatty acids from complex host sources into bacterial membranes, particularly Staphylococcus aureus. To achieve this, protocols for the enrichment of lipoprotein particles from chicken egg yolk and subsequent fatty acid profiling of bacterial phospholipids utilizing mass spectrometry are described.

Staphylococcus aureus and other Gram-positive pathogens incorporate fatty acids from the environment into membrane phospholipids. During infection, the ...

PeptiQuick, a One-Step Incorporation of Membrane Proteins into Biotinylated Peptidiscs for Streamlined Protein Binding Assays

Membrane proteins, including transporters, channels, and receptors, constitute nearly one-fourth of the cellular proteome and over half of current drug targets. Yet, a major barrier to their characterization and exploitation in academic or industrial settings is that most biochemical, biophysical, and drug screening strategies require these proteins to be in a water-soluble state. Our laboratory recently developed the peptidisc, a membrane mimetic offering a &#8220;one-size-fits-all&#8221; approach to the problem of membrane protein solubility. We present here a streamlined protocol that combines protein purification and peptidisc reconstitution in a single chromatographic step. This workflow, termed PeptiQuick, allows for bypassing dialysis and incubation with polystyrene beads, thereby greatly reducing exposure to detergent, protein denaturation, and sample loss. When PeptiQuick is performed with biotinylated scaffolds, the preparation can be directly attached to streptavidin-coated surfaces. There is no need to biotinylate or modify the membrane protein target. PeptiQuick is showcased here with the membrane receptor FhuA and antimicrobial ligand colicin M, using biolayer interferometry to determine the precise kinetics of their interaction. It is concluded that PeptiQuick is a convenient way to prepare and analyze membrane protein-ligand interactions within one&#160;day in a detergent-free environment.

We present a method that combines membrane protein purification and reconstitution into peptidiscs in a single chromatographic step. Biotinylated scaffolds are used for direct surface attachment and measurement of protein-ligand interactions via biolayer interferometry.

Membrane proteins, including transporters, channels, and receptors, constitute nearly one-fourth of the cellular proteome and over half of current drug ...

Native Cell Membrane Nanoparticles System for Membrane Protein-Protein Interaction Analysis

Protein-protein interactions in cell membrane systems play crucial roles in a wide range of biological processes- from cell-to-cell interactions to signal transduction; from sensing environmental signals to biological response; from metabolic regulation to developmental control. Accurate structural information of protein-protein interactions is crucial for understanding the molecular mechanisms of membrane protein complexes and for the design of highly specific molecules to modulate these proteins. Many in vivo and in vitro approaches have been developed for the detection and analysis of protein-protein interactions. Among them the structural biology approach is unique in that it can provide direct structural information of protein-protein interactions at the atomic level. However, current membrane protein structural biology is still largely limited to detergent-based methods. The major drawback of detergent-based methods is that they often dissociate or denature membrane protein complexes once their native lipid bilayer environment is removed by detergent molecules. We have been developing a&#160;native cell membrane nanoparticle system for membrane protein structural biology. Here, we demonstrate the use of this system in the analysis of protein-protein interactions on the cell membrane with a case study of the oligomeric state of AcrB.

Presented here is a protocol for the determination of oligomeric state of membrane proteins that utilizes a native cell membrane nanoparticle system in conjunction with electron microscopy.

Protein-protein interactions in cell membrane systems play crucial roles in a wide range of biological processes- from cell-to-cell interactions to signal ...