This research was supported by VCU startup fund (to Y.G.) and the National Institute Of General Medical Sciences of the National Institutes of Health under Award Number R01GM132329 (to Y.G.) The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. We thank Montserrat Samso and Kevin McRoberts for their generous support for video recording.

1. Protein expression
<ol>
	<li>Inoculate 15 mL of Terrific Broth (TB) media with antibiotics specific to plasmids with BL21(DE3) pLysS cells containing the AcrB expressing plasmids in 50 mL tubes overnight at 37 &#176;C with shaking at 250 rpm.</li>
	<li>Check the optical density of the overnight culture at 600 nm (OD600) and ensure that it is over 2.0.</li>
	<li>Dilute 5 mL of cell culture into 1 L of TB media containing antibiotics specific to plasmids and incubate at 37 &#176;C with shaking until OD600=0.8 and then induce with IPTG that has a final concentration of 1 mM.</li>
	<li>Carry out induction at 20 &#176;C with shaking for 20 h.</li>
	<li>Pellet down the cells using centrifugation for 15 min at 4 &#176;C and 4,000 x g.</li>
</ol>
2. Cell lysis and membrane isolation
<ol>
	<li>Resuspend the cell pellet in Buffer A (Table 1, use DDM Buffer A or NCMNs Buffer A depending on purification scheme that is to follow) using 80 mL for every 20 g of the cell pellet.</li>
	<li>Homogenize the resuspended cell pellets by using a glass Dounce homogenizer at 4 &#176;C or if at room temperature be sure to put on ice immediately after.</li>
	<li>Transfer the sample into a metal beaker on ice and lyse the cells by loading them into a high-pressure homogenizer at 4 &#176;C and 1,500 bar.
	<ol>
		<li>Repeat the process of loading the cells into the homogenizer and allowing them to pass through the homogenizer for 3-4 cycles or until the lysate begins to clarify.</li>
	</ol>
	</li>
	<li>Centrifuge the lysate at 15,000 x g for 30 min at 4 &#176;C.</li>
	<li>Collect the supernatant and load into ultracentrifuge tubes and centrifuge at 215,000 x g for 1 h at 4 &#176;C.</li>
	<li>Discard the supernatant and collect the membrane pellets from the ultracentrifuge tubes. Store any excess membrane pellet at -80 &#176;C.</li>
</ol>
3. Preparation of native cell membrane nanoparticles
<ol>
	<li>Resuspend 1 g of membrane pellet in 10 mL NCMNs Buffer A (Table 1).</li>
	<li>Homogenize the resuspended cell membrane sample with a glass Dounce homogenizer at 20 &#176;C.</li>
	<li>Transfer the suspended membrane sample to a 50 mL polypropylene tube and add membrane active polymers stock solution and additional NCMNs Buffer A to bring the sample to a final concentration of 2.5% (w/v) membrane active polymer (NCMNP1-1 or NCMNP5-2). 
	NOTE: Stock solutions of membrane active polymers should be made in double distilled water and can be kept at varying concentrations, but typically 10% (w/v).</li>
	<li>Shake the sample for 2 h at 20 &#176;C.</li>
	<li>Load the sample into an ultracentrifuge and spin at 150,000 x g for 1 h at 20 &#176;C.</li>
	<li>While the sample is being ultra-centrifuged begin to equilibrate a 5 mL Ni-NTA column with 25 mL of NCMNs Buffer A.</li>
	<li>Collect the supernatant after ultracentrifugation is complete and load it onto 5 mL of Ni-NTA column at room temperature with a flow rate of 0.5 mL/min using a syringe pump.</li>
	<li>Wash fast protein liquid chromatography (FPLC) lines with enough NCMNs Buffer B (Table 1) to completely flush the system and then connect the column to the FPLC machine.</li>
	<li>Wash the column with 30 mL of NCMNs Buffer B with a flow rate of 1 mL/min and collect the flow through.</li>
	<li>Wash the column with 30 mL of NCMNs Buffer C (Table 1) with a flow rate of 1 mL/min and collect the flow through.</li>
	<li>Elute the protein with 20 mL of NCMNs Buffer D (Table 1) at a flow rate of 0.5 mL/min and collect the sample using a fraction collector and the fractions each being set to 1.0 mL.</li>
	<li>Store the protein samples at 4 &#176;C.</li>
	<li>Run an SDS-PAGE gel electrophoresis assay in order to check the samples that correspond to peaks observed on the FPLC elution graph.</li>
</ol>
4. SDS-PAGE gel electrophoresis
<ol>
	<li>Prepare the casting chamber by clamping the glass to the casting apparatus.</li>
	<li>Prepare the 12% separation gel according to the recipe listed in Table 1. 
	NOTE: Once TEMED is added the gel will polymerize quickly so only add this once ready to pour the gel.</li>
	<li>Pour the gel, leaving 2 cm below the bottom of the comb for the stacking gel.</li>
	<li>Remove any bubbles by layering the top of the gel with 100% isopropanol and wait for the separation gel to polymerize.</li>
	<li>Remove the isopropanol and wash out any traces of the isopropanol with distilled water.</li>
	<li>Prepare the stacking gel according to the recipe listed in Table 1. 
	NOTE: Once TEMED is added the gel will polymerize quickly so only add this once ready to pour the gel.</li>
	<li>Pour the stacking gel on top of the separation gel.</li>
	<li>Add a comb to the chamber to form the wells and wait for the stacking gel to completely polymerize.</li>
	<li>Place 1 &#181;L of 1 M DTT into a microcentrifuge tube for each fraction sample that needs to be run on the gel.</li>
	<li>Add 7 &#181;L of 4x loading buffer to each tube as well.</li>
	<li>Add 20 &#181;L of sample from the fractions that need to be run on the gel to each microcentrifuge tube.</li>
	<li>Vortex the tubes containing the samples.</li>
	<li>Spin down the sample tubes using a tabletop microcentrifuge for 3 s and make sure all the sample has returned to bottom of the tubes.</li>
	<li>Prepare the gel electrophoresis cell by securing a 12% polyacrylamide gel cassette into place and filling the inner and outer chambers of the cell with Tris-acetate-EDTA (TAE) Buffer (Table 1).</li>
	<li>Load the molecular weight marker into the first lane of the gel with the appropriate volume required by its instructions.</li>
	<li>Load 28 &#181;L of the sample from each tube mixed with DTT and loading buffer.</li>
	<li>Place the lid of the electrophoresis cell on top of the box and plug the lid into the power supply.</li>
	<li>Turn on the power supply and set it to 100 V and allow it to run for 20 min.</li>
	<li>After 20 min, increase the power supply current to 140 V and continue to run it for another 30-40 min or until the band of loading buffer dye reaches the bottom of the gel cassette.</li>
	<li>After completing electrophoresis stain and de-stain the gel to visualize the protein bands contained within the gel.</li>
</ol>
5. Protein purification using DDM
NOTE: The purpose of carrying out this purification process is to serve as a control for the experiments utilizing the membrane active polymers.
<ol>
	<li>Resuspend 6-10 g of membrane pellet, from step 2.6, in DDM Buffer A (Table 1) using 5 mL/g of membrane pellet.</li>
	<li>Homogenize the resuspended sample with a glass Dounce homogenizer at 4 &#176;C or if at room temperature be sure to put on ice immediately after.</li>
	<li>Transfer sample to a 50 mL of polypropylene tube and add DDM and additional DDM Buffer A to bring the sample to a final concentration of 2% DDM.</li>
	<li>Shake the sample for 2 h at 4 &#176;C.</li>
	<li>Load the sample into ultracentrifuge tubes and centrifuge for 1 h at 4 &#176;C and 150,000 x g.</li>
	<li>While sample is being ultra-centrifuged begin to prepare the 5 mL of Ni-NTA column by washing with 25 mL of DDM Buffer A (Table 1).</li>
	<li>After ultracentrifugation is complete, collect the supernatant and load it onto the 5 mL Ni-NTA column at 4 &#176;C with a flow rate of 0.5 mL/min using a syringe pump.</li>
	<li>Wash FPLC lines with enough DDM Buffer B + 0.05% DDM (Table 1) to completely flush the system and then attach the column to the FPLC machine.</li>
	<li>Wash the column with 30 mL of DDM Buffer B + 0.05% DDM with a flow rate of 1 mL/min and collect the flow through.</li>
	<li>Wash the column with 30 mL of DDM Buffer C + 0.05% DDM (Table 1) with a flow rate of 1 mL/min and collect the flow through.</li>
	<li>Elute the protein with 20 mL of DDM Buffer D + 0.05% DDM (Table 1) at a flow rate of 0.5 mL/min and collect the flow through using a fraction collector and the fractions each being set to 1.0 mL.</li>
	<li>Select the fractions containing the elution peak for pooling and concentrating by using a centrifugal concentrator and centrifuging at 4,000 x g and 4 &#176;C until reaching 500 &#181;L.</li>
	<li>Using a 500 &#181;L loop load the sample into the FPLC machine and then onto the 25 mL size exclusion column at 4 &#176;C. Elute using 30 mL of DDM Buffer E + 0.05% DDM (Table 1) at a flow rate of 0.5 mL/min and collect as fractions with the fraction sizes set to being 0.5 mL per a fraction.</li>
	<li>Take the fractions and measure the protein concentration using 280 nm absorbance to confirm UV-Vis curve from the FPLC elution graph.</li>
	<li>Collect 20 &#181;L from each sample fraction that correspond to peaks observed on the FPLC elution graph and were confirmed to be accurate with the absorbance test.</li>
	<li>Freeze the remainder of those sample fractions using liquid nitrogen or dry ice in desired aliquots and store protein samples at -80 &#176;C.</li>
	<li>Run an SDS-PAGE gel electrophoresis assay as previously described to check the samples that correspond to peaks observed on the FPLC elution graph.</li>
</ol>
6. Negative stain grid preparation
<ol>
	<li>Place the grids that are going to be used for the sample preparation on a glass slide wrapped in filter paper with the carbon side facing up.</li>
	<li>Place the glass slide with the electron microscope grids into the chamber of a glow discharger between the two electrodes and replace the glass lid making sure it is centered and well-sealed.</li>
	<li>Run the glow discharging machine and make sure that the purple light generated by the plasma is visible.</li>
	<li>When the machine is done running, wait until the chamber has reached atmospheric pressure to remove the glass lid and then return the slide with the grids to the bench where samples will be loaded onto them.</li>
	<li>Adjust the concentration of the purified protein samples to about 0.1 mg/mL by either diluting the sample with the appropriate buffer or concentrating using a centrifugal concentrator.</li>
	<li>Load 3.5 &#181;L of protein sample onto the 10 nm thick carbon grid and wait for 1 min.</li>
	<li>Remove the liquid from the surface of the EM grid with a filter paper.</li>
	<li>Wash the grid surface 3x by picking up 3 &#181;L droplets of water with the grid and removing the water from the grid with filter paper in between picking up each droplet.</li>
	<li>Wash the grid surface 2x by picking up 3 &#181;L droplets of fresh, filtered 2% uranium acetate and remove the wash solutions on the grid with filter paper in between picking up each droplet.</li>
	<li>Stain the grid with a 3 &#181;L droplet of fresh, filtered 2% uranium acetate for 1 min.</li>
	<li>Remove the uranium acetate solution on the surface of the EM grid with filter paper and air dry the grid for at least 1 min.</li>
	<li>Store the grid in a grid box for the later use.</li>
</ol>
7. EM imaging
<ol>
	<li>Load the prepared grid into the grid holder at a safe workbench.</li>
	<li>Prepare the microscope by placing the microscopes dewar into a polystyrene box and fill the dewar 3/4th full of liquid nitrogen.</li>
	<li>Confirm that the rubber cover of the microscope window is covering it and then load the dewar onto the microscope by placing the copper wires into the dewar until it fits on the platform.</li>
	<li>Fill the remainder of the dewar with liquid nitrogen and place a cap on top of the dewar to cover it.</li>
	<li>Turn on the high tension, condition the microscope, and wait for the microscope to warm up and establish a safe vacuum level for use.</li>
	<li>Once the microscope is ready open the column valve and confirm the beam is present by removing the rubber cover on the microscope window.</li>
	<li>Check the beam stigmation by spreading in and out by adjusting the intensity of the beam. If astigmatism is present the beam will have an elliptical shape as one moves away from cross-over and the beam should be the same oval shape on both sides.</li>
	<li>Adjust the microscope binoculars to be the right height and distance as per the eyes of the observer.</li>
	<li>Check the beam positioning for Search, Focus, and Exposure modes by cycling through all three modes until the beam is center in each.</li>
	<li>Close the column valve and proceed to load the grid holder into the electron microscope.</li>
	<li>Adjust the microscope to eucentric height by using the microscopes wobbler feature and adjusting the movement of the stage using the Z-axis until there is no side-to-side motion of the sample in the center.</li>
	<li>Using the low dose Search mode on the microscope search the grid for a desirable area of reasonable contrast with sample molecules present to focus on the sample.</li>
	<li>Focus on the sample in Focus mode by using a high step size until the sample is seen and then decrease steps to find the correct focus level.</li>
	<li>Zero and blank the beam and then be sure the rubber pad is covering the microscope window.</li>
	<li>Using the Exposure mode take the image of the desired grid area for a 1 s exposure at 62,000x magnification and check the obtained image.</li>
	<li>When done imaging a grid close the column valves and remove the grid holder from the column.</li>
	<li>When finished imaging all grids of interest shutdown the microscope by turning off the filament power and removing the liquid nitrogen dewar from the microscope.</li>
	<li>Place something to absorb moisture on the stand where the dewar sat to catch condensation from the copper coils of the microscope.</li>
	<li>Activate the Cryo Cycle mode and make sure the turbo pump is turned off.</li>
</ol>

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Y.G is listed as inventor of the membrane active polymer NCMNP5-2 and NCMN system.

Protein-protein interactions are important for the integrity of the structure and function of membrane proteins. Many approaches have been developed to investigate protein-protein interactions. When compared with soluble proteins, membrane proteins and their PPIs are more difficult to study due to the unique intrinsic properties of membrane proteins. This difficulty mainly comes from the requirement of membrane proteins to be embedded in a native lipid bilayer environment for structural stability and functionality. This ...

Protein-protein interactions (PPI) play pivotal roles throughout biology, from the maintenance of the structure and function of proteins to the regulation of entire systems. PPIs come in many different forms and can be categorized based on what types of interactions they form. One such categorization is homooligomeric or heterooligomeric, based on whether the interactions are between identical subunits or different proteins acting as subunits. Another categorization is based on the strength of the interaction if the interactions leads to the formation of stable complexes or transient complex states. Structural information about the PPIs between proteins is crucial in understanding the mechanism by which proteins carry out their function. It has been estimated that over 80% of proteins rely on complex formation in order to carry out their functional roles1. Given the percentage of proteins observed to be reliant on PPIs for proper innate functioning their significance in biology is readily apparent, yet being able to properly investigate the proteins that engage in these interactions has remained challenging due to limitations in the techniques available to experimentally observe when proteins are forming PPIs.
There is a high degree of disagreement between the results for many experimentally determined PPIs due to the high amount of noise, false positives, and false negatives that are derived from many of the currently available PPI determining techniques. This is particularly so for the yeast two-hybrid (Y2H) system, tandem affinity purification-mass spectrometry (TAP-MS), and fluorescence resonance energy transfer (FRET), which represent three of the most commonly used methods for PPI determination2,3,4,5,6,7. In comparison, protein structural biology techniques, such as nuclear magnetic resonance (NMR), X-ray crystallography and electron microscopy (EM), can be used to gain high-resolution structural information about protein-protein interactions down to the atomic level and allow for the direct visual confirmation of the interactions occurring for a target protein of interest. All of the currently available non-structure based PPI determining research techniques (e.g., Y2H, TAP-MS, and FRET) lack this ability and additionally suffer from having difficulty in identifying weak and transient interactions between proteins5. These shortcomings are further enhanced when studying membrane proteins due to the added complexity brought on by the additional variable of the lipid environment that influences the formation of proper quaternary structure and heteromeric complex assembly.
Membrane proteins make up a large portion of the proteome and are known to play many crucial roles in proper cell functioning within all living organisms. Despite the fact that membrane proteins are estimated to make up 27% of the human genome and constitute ~60% of all current drug targets there is a major anomaly in the number of solved models for membrane proteins which make up only ~2.2% of all published protein structures8,9,10. The primary cause of the discrepancy in the availability of structural information lies in the intrinsic properties of membrane proteins themselves. Due to their poor solubility, reliance on interactions with a lipid environment for maintaining native structure and function, and various physicochemical properties of the lipid membrane itself, membrane proteins continue to represent a substantial problem when implementing structural biology techniques to study them. Of these intrinsic properties, the most important for obtaining accurate structural information is the requirement of having interactions with their natural lipid environment for them to maintain their native structure and function. The lipid environment is such an integral part of the structure and function of membrane proteins that the concept of memtein (a combination of the words membrane and protein) has been proposed as the fundamental unit of membrane-protein structure and function11. Despite the importance of lipid-protein interactions the currently available structure-based PPI determination techniques often require that the proteins being studied be soluble or be solubilized with detergents. Exposure to harsh detergents can denature the proteins or cause false positives and negatives due to delipidation, which can induce aggregation, denaturation of protein, dissociation of non-covalent interactions, and formation of artificial oligomeric states. Due to the necessity of maintaining a natural lipid environment for accurate determination of the native oligomeric state of membrane proteins we have developed a Native Cell Membrane Nanoparticles system (NCMNs)12 based on the previously reported Styrene Maleic Acid Lipid Particles (SMALP)13 method.
SMALP uses styrene maleic acid copolymer as a membrane active polymer to extract and solubilize membrane proteins. Poly(Styrene-co-Maleic Acid) (SMA) is a unique amphiphilic polymer due to its hydrophobic styrene moiety and diametrically opposing hydrophilic maleic acid moiety. It forms nanoparticles by adsorbing to, destabilizing, and disrupting the cell membrane in a pH-dependent mechanism14. This functional activity of SMA is what allows it to be utilized as a detergent-free system for membrane protein extraction and solubilization. The NCMN system is distinct from the SMALP system in several aspects. The most unique feature of the NCMN system is that it has a membrane active polymer library with a multitude of polymers that have unique properties that make them suitable for the isolation of many different membrane proteins that require different conditions for stability and native functioning. The NCMN system also has different protocols compared to the SMALP method when it comes to preparing the nanoparticles. One such example is that NCMNs use a single-step nickel affinity column purification for high-resolution structure determination; the effect of these different protocols can be observed when comparing the NCMNs protocol, which generated 3.2 and 3.0 &#197; cryo-EM AcrB structures, with a similar study using the SMALP method, which resulted in a cryo-EM AcrB structure at an 8.8 &#197; resolution12,15. These unique features of the NCMN system are the improvements made on the previously established SMALP method and make it an ideal candidate for the study of PPIs.
The multidrug efflux transporter AcrB exists as functional homotrimer in E. coli12. A mutagenic analysis suggested that a single P223G point mutation, located on a loop that is responsible for the stability of the AcrB trimer, destabilizes the trimer state and yields an AcrB monomer when prepared with the detergent DDM, which could be detected with native blue gel electrophoresis16. However, the FRET analysis of the AcrB-P223G mutant suggested that when in the native cell membrane lipid bilayer environment, the majority of mutant AcrB-P223G still existed as a trimer and that the expression levels for both wild type AcrB and AcrB-P223G are similar. Yet despite the FRET analysis results, an activity assay for the mutant transporter showed that activity dramatically decreased when compared to the wild type16. Although FRET technology has been popularly used for the analysis of protein-protein interactions, research has shown that it can frequently give false positives17,18,19,20.
Recently, high-resolution cryo-EM structures of the AcrB trimer were determined which showed the interaction of AcrB with its associated native cell membrane lipid bilayer through the use of the NCMNs12. In principle, the NCMNs can readily be utilized for the analysis of protein-protein interactions found on the native cell membrane. In a test of this system, experiments were carried out to directly observe the oligomeric state of AcrB-P223G with the use of native cell membrane nanoparticles and negative staining electron microscopy. In order to compare with the wild type AcrB nanoparticles, we used the same membrane active polymer (SMA2000 made in our laboratory, which is indexed as NCMNP1-1 in the NCMN library) used for high-resolution cryo-EM structure determination of AcrB and alternative polymers found in the NCMNs library12. Based on the previously reported results, it was expected that the majority of the AcrB-P223G mutant would exist in the form of trimeric native cell membrane nanoparticles21. However, no AcrB trimers were found present in the sample, such as the ones that were observed with the wild type AcrB. This suggests the majority of AcrB-P223G does not form trimers on the native cell membrane as previously suggested.
Here we report a detailed analysis of the mutant E. coli AcrB-P223G in a comparison with the wild type E. coli AcrB using the NCMNs. This case study of AcrB suggests NCMNs is a good system for protein-protein interaction analysis.

<table><tbody><tr><td>&lt;strong&gt;Chemicals&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>30% Acrylamide/BIS SOL (37.5:1)</td><td>Bio-Rad</td><td>161-0158</td><td></td></tr><tr><td>4x Laemmli Sample Buffer (Loading Buffer)</td><td>Bio-Rad</td><td>1610747</td><td></td></tr><tr><td>Acetic Acid Glacial</td><td>ThermoFisher Scientific</td><td>A38S-212</td><td></td></tr><tr><td>Ammonium Persulfate (APS)</td><td>Bio-Rad</td><td>161-0700</td><td></td></tr><tr><td>Chloramphenicol</td><td>Goldbio</td><td>C-105-5</td><td></td></tr><tr><td>Coomassie Brilliant Blue R-250 protein stain powder</td><td>Bio-Rad</td><td>161-0400</td><td></td></tr><tr><td>DTT (Dithiothreitol) (&gt; 99% pure) Protease free</td><td>Goldbio</td><td>DTT10</td><td></td></tr><tr><td>Glycerol</td><td>ThermoFisher Scientific</td><td>G33-4</td><td></td></tr><tr><td>HEPES</td><td>ThermoFisher Scientific</td><td>BP310-1</td><td></td></tr><tr><td>Imidazole</td><td>Affymetrix</td><td>17525 1 KG</td><td></td></tr><tr><td>IPTG</td><td>Goldbio</td><td>I2481C100</td><td></td></tr><tr><td>Kanamycin</td><td>Goldbio</td><td>K-120-25</td><td></td></tr><tr><td>Magnesium chloride hexahydrate</td><td>ThermoFisher Scientific</td><td>AA3622636</td><td></td></tr><tr><td>Methanol</td><td>ThermoFisher Scientific</td><td>A412-4</td><td></td></tr><tr><td>N,N-dimethylethylenediamine (EDTA)</td><td>Merck</td><td>8.03779.0100</td><td></td></tr><tr><td>NCMNS-P5-2</td><td>Not commercially available yet</td><td>Submit request for obtaining to corresponding author</td><td></td></tr><tr><td>Precision Plus Protein Dual Color Standard</td><td>Bio-Rad</td><td>161-0374</td><td></td></tr><tr><td>SDS (Sodium Dodecyl Sulfate)</td><td>Bio-Rad</td><td>161-0301</td><td></td></tr><tr><td>SMA2000</td><td>Cray Valley</td><td>Submit request for obtaining to corresponding author</td><td></td></tr><tr><td>Sodium Chloride</td><td>ThermoFisher Scientific</td><td>S271-10</td><td></td></tr><tr><td>TCEP-HCl</td><td>Goldbio</td><td>TCEP25</td><td></td></tr><tr><td>TEMED</td><td>Bio-Rad</td><td>161-0800</td><td></td></tr><tr><td>Terrific Broth Media</td><td>Affymetrix</td><td>75856 1 KG</td><td></td></tr><tr><td>Tris Base</td><td>Bio-Rad</td><td>161-0719</td><td></td></tr><tr><td>Uranyl Acetate</td><td>Ambinter</td><td>Amb22348393</td><td></td></tr><tr><td>&lt;strong&gt;Equipment&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Avanti J-26S XPI</td><td>Beckman Coulter</td><td>B14538</td><td></td></tr><tr><td>Avanti JXN-30</td><td>Beckman Coulter</td><td>B34193</td><td></td></tr><tr><td>Carbon Electron Microscope Grids (10 nm)</td><td>Electron Microscopy Sciences</td><td>CF300-Cu-TH</td><td></td></tr><tr><td>Con-Torque Tissue Homogenizer</td><td>Eberbach</td><td>E7265</td><td></td></tr><tr><td>Corning LSE Mini Microcentrifuge</td><td>ThermoFisher Scientific</td><td>07-203-954</td><td></td></tr><tr><td>EmulsiFlex-C3</td><td>Avestin</td><td></td><td></td></tr><tr><td>Fraction Collector F9-R</td><td>GE Healthcare Life Sciences</td><td>29003875</td><td></td></tr><tr><td>Mini-PROTEAN Tetra Vertical Electrophoresis Cell</td><td>Bio-Rad</td><td>165-8004</td><td></td></tr><tr><td>NanoDrop 2000 Spectrophotometer</td><td>ThermoFisher Scientific</td><td>ND-2000</td><td></td></tr><tr><td>Optima L-90K Ultracentrifuge</td><td>Beckman Coulter</td><td>PN LL-IM-12AB</td><td></td></tr><tr><td>PELCO easiGlow Glow Discharge Cleaning System</td><td>Ted Pella</td><td>91000S-230</td><td></td></tr><tr><td>Potter-Elvehjem Safe Grind Tissue Grinder</td><td>Wheaton</td><td>358013</td><td></td></tr><tr><td>PowerPac Basic Power Supply</td><td>Bio-Rad</td><td>164-5050</td><td></td></tr><tr><td>Razel R99-E Variable Speed Syringe Pump</td><td>Razel Scientific Instruments</td><td></td><td></td></tr><tr><td>Superdex 200 Increase 10/300 GL</td><td>GE Healthcare Life Sciences</td><td>28990944</td><td></td></tr><tr><td>Tecnai F20 200kV</td><td>FEI</td><td></td><td></td></tr><tr><td>Type 70 Ti Fixed-Angle Rotor</td><td>Beckman Coulter</td><td></td><td></td></tr><tr><td>&lt;strong&gt;General Materials&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>1.5 ml Microcentrifuge Tubes</td><td>ThermoFisher Scientific</td><td>05-408-129</td><td></td></tr><tr><td>4 ml Amicon Ultra-4 30 kDa</td><td>Millipore Sigma</td><td>UFC803024</td><td></td></tr><tr><td>AKTA pure 25 L1 FPLC</td><td>GE Healthcare Life Sciences</td><td>29018225</td><td></td></tr><tr><td>BL21(DE3)pLysS Cells</td><td>ThermoFisher Scientific</td><td>C606003</td><td></td></tr><tr><td>Falcon 50 ml Conical Centrifuge Tube</td><td>ThermoFisher Scientific</td><td>14-959-49A</td><td></td></tr><tr><td>HisTrap HP 5 ml Column</td><td>GE Healthcare Life Sciences</td><td>17524802</td><td></td></tr><tr><td>pET-24a</td><td>EMD Biosciences</td><td>69749-3</td><td></td></tr></tbody></table>

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.

Using the procedures presented here, samples of E. coli AcrB wild type and E. coli mutant AcrB-P223G were purified. The samples were then adsorbed to carbon negative stain electron microscopy grids and stained using uranyl acetate with the side blotting method22. Negative stain images were collected using transmission electron microscopy. The negative stain image for the AcrB wild type sample purified with DDM reveals a homogenous solution of monodispersed particles with the prot...

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

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 ...

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Research

JoVE Journal

Biochemistry

5.9K Views.  Virginia Commonwealth University. Hier wordt een protocol gepresenteerd voor de bepaling van oligomere toestand van membraaneiwitten dat gebruik maakt van een inheems celmembraan nanodeeltjessysteem in combinatie met elektronenmicroscopie.

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

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 ...

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 ...

Monitoring Protein Adsorption with Solid-state Nanopores

Solid-state nanopores have been used to perform measurements at the single-molecule level to examine the local structure and flexibility of nucleic acids 1-6, the unfolding of proteins 7, and binding affinity of different ligands 8. By coupling these nanopores to the resistive-pulse technique 9-12, such measurements can be done under a wide variety of conditions and without the need for labeling 3. In the resistive-pulse technique, an ionic salt solution is introduced on both sides of the nanopore. Therefore, ions are driven from one side of the chamber to the other by an applied transmembrane potential, resulting in a steady current. The partitioning of an analyte into the nanopore causes a well-defined deflection in this current, which can be analyzed to extract single-molecule information. Using this technique, the adsorption of single proteins to the nanopore walls can be monitored under a wide range of conditions 13. Protein adsorption is growing in importance, because as microfluidic devices shrink in size, the interaction of these systems with single proteins becomes a concern. This protocol describes a rapid assay for protein binding to nitride films, which can readily be extended to other thin films amenable to nanopore drilling, or to functionalized nitride surfaces. A variety of proteins may be explored under a wide range of solutions and denaturing conditions. Additionally, this protocol may be used to explore more basic problems using nanopore spectroscopy.

A method of using solid-state nanopores to monitor the non-specific adsorption of proteins onto an inorganic surface is described. The method employs the resistive-pulse principle, allowing for the adsorption to be probed in real-time and at the single-molecule level. Because the process of single protein adsorption is far from equilibrium, we propose the employment of parallel arrays of synthetic nanopores, enabling for the quantitative determination of the apparent first-order reaction rate constant of protein adsorption as well as and the Langmuir adsorption constant.

Solid-state nanopores have been used to perform measurements at the single-molecule level to examine the local structure and flexibility of nucleic acids ...

Bioengineering

Biotinylated Cell-penetrating Peptides to Study Intracellular Protein-protein Interactions

Here we present a protocol to study intracellular protein-protein interactions that is based on the widely used biotin-avidin pull-down system. The modification presented includes the combination of this technique with cell-penetrating sequences. We propose to design cell-penetrating baits that can be incubated with living cells instead of cell lysates and therefore the interactions found will reflect those that occur within the intracellular context. Connexin43 (Cx43), a protein that forms gap junction channels and hemichannels is down-regulated in high-grade gliomas. The Cx43 region comprising amino acids 266-283 is responsible for the inhibition of the oncogenic activity of c-Src in glioma cells. Here we use TAT as the cell-penetrating sequence, biotin as the pull-down tag and the region of Cx43 comprised between amino acids 266-283 as the target to find intracellular interactions in the hard-to-transfect human glioma stem cells. One of the limitations of the proposed method is that the molecule used as bait could fail to fold properly and, consequently, the interactions found could not be associated with the effect. However, this method can be especially interesting for the interactions involved in signal transduction pathways because they are usually carried out by intrinsically disordered regions and, therefore, they do not require an ordered folding. In addition, one of the advantages of the proposed method is that the relevance of each residue on the interaction can be easily studied. This is a modular system; therefore, other cell-penetrating sequences, other tags, and other intracellular targets can be employed. Finally, the scope of this protocol is far beyond protein-protein interaction because this system can be applied to other bioactive cargoes such as RNA sequences, nanoparticles, viruses or any molecule that can be transduced with cell-penetrating sequences and fused to pull-down tags to study their intracellular mechanism of action.

This is a protocol to study intracellular protein-protein interactions based on the biotin-avidin pull-down system with the novelty of combining cell-penetrating sequences. The main advantage is that the target sequence is incubated with living cells instead of cell lysates and therefore the interactions will occur within the cellular context.

Here we present a protocol to study intracellular protein-protein interactions that is based on the widely used biotin-avidin pull-down system. The ...

Membrane-SPINE: A Biochemical Tool to Identify Protein-protein Interactions of Membrane Proteins In Vivo

Membrane proteins are essential for cell viability and are therefore important therapeutic targets1-3. Since they function in complexes4, methods to identify and characterize their interactions are necessary5. To this end, we developed the Membrane Strep-protein interaction experiment, called Membrane-SPINE6. This technique combines in vivo cross-linking using the reversible cross-linker formaldehyde with affinity purification of a Strep-tagged membrane bait protein. During the procedure, cross-linked prey proteins are co-purified with the membrane bait protein and subsequently separated by boiling. Hence, two major tasks can be executed when analyzing protein-protein interactions (PPIs) of membrane proteins using Membrane-SPINE: first, the confirmation of a proposed interaction partner by immunoblotting, and second, the identification of new interaction partners by mass spectrometry analysis. Moreover, even low affinity, transient PPIs are detectable by this technique. Finally, Membrane-SPINE is adaptable to almost any cell type, making it applicable as a powerful screening tool to identify PPIs of membrane proteins.

Membrane-SPINE: A Biochemical Tool to Identify Protein-protein Interactions of Membrane Proteins In Vivo

A biochemical approach is described to identify in vivo protein-protein interactions (PPI) of membrane proteins. The method combines protein cross-linking, affinity purification and mass spectrometry, and is adaptable to almost any cell type or organism. With this approach, even the identification of transient PPIs becomes possible.

Membrane proteins are essential for cell viability and are therefore important therapeutic targets1-3. Since they function in complexes4, methods to ...

Evaluation of Protein&#8211;Protein Interactions using an On-Membrane Digestion Technique

Numerous intracellular proteins physically interact in accordance with their intracellular and extracellular circumstances. Indeed, cellular functions largely depend on intracellular protein&#8211;protein interactions. Therefore, research regarding these interactions is indispensable to facilitating the understanding of physiologic processes. Co-precipitation of associated proteins, followed by mass spectrometry (MS) analysis, enables the identification of novel protein interactions. In this study, we have provided details of the novel technique of immunoprecipitation-liquid chromatography (LC)-MS/MS analysis combined with on-membrane digestion for the analysis of protein&#8211;protein interactions. This technique is suitable for crude immunoprecipitants and can improve the throughput of proteomic analyses. Tagged recombinant proteins were precipitated using specific antibodies; next, immunoprecipitants blotted onto polyvinylidene difluoride membrane pieces were subjected to reductive alkylation. Following trypsinization, the digested protein residues were analyzed using LC-MS/MS. Using this technique, we were able to identify several candidate associated proteins. Thus, this method is convenient and useful for the characterization of novel protein&#8211;protein interactions.

Here, we present a protocol for an on-membrane digestion technique for the preparation of samples for mass spectrometry. This technique facilitates the convenient analysis of protein&#8211;protein interactions.

Numerous intracellular proteins physically interact in accordance with their intracellular and extracellular circumstances. Indeed, cellular functions ...

Cell-Free Production of Proteoliposomes for Functional Analysis and Antibody Development Targeting Membrane Proteins

Membrane proteins play essential roles in a variety of cellular processes and perform vital functions. Membrane proteins are medically important in drug discovery because they are the targets of more than half of all drugs. An obstacle to conducting biochemical, biophysical, and structural studies of membrane proteins as well as antibody development has been the difficulty in producing large amounts of high-quality membrane protein with correct conformation and activity. Here we describe a &#8220;bilayer-dialysis method&#8221; using a wheat germ cell-free system, liposomes, and dialysis cups to efficiently synthesize membrane proteins and prepare purified proteoliposomes in a short time with a high success rate. Membrane proteins can be produced as much as in several milligrams, such as GPCRs, ion channels, transporters, and tetraspanins. This cell-free method contributes to reducing the time, cost and effort for preparing high-quality proteoliposomes, and provides suitable means for functional analysis of membrane proteins, drug targets screening, and antibody development.

This protocol describes an efficient cell-free method for production of high-quality proteoliposome by bilayer-dialysis method using wheat cell-free system and liposomes. This method provides suitable means for functional analysis of membrane proteins, drug targets screening, and antibody development.

Membrane proteins play essential roles in a variety of cellular processes and perform vital functions. Membrane proteins are medically important in drug ...

Ligand Nano-cluster Arrays in a Supported Lipid Bilayer

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

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

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

Protein Membrane Overlay Assay: A Protocol to Test Interaction Between Soluble and Insoluble Proteins in vitro

Validating interactions between different proteins is vital for investigation of their biological functions on the molecular level. There are several methods, both in vitro and in vivo, to evaluate protein binding, and at least two methods that complement the shortcomings of each other should be conducted to obtain reliable insights. 
For an in vivo assay, the bimolecular fluorescence complementation (BiFC) assay represents the most popular and least invasive approach that enables to detect protein-protein interaction within living cells, as well as identify the intracellular localization of the interacting proteins 1,2. In this assay, non-fluorescent N- and C-terminal halves of GFP or its variants are fused to tested proteins, and when the two fusion proteins are brought together due to the tested proteins&#x2019; interactions, the fluorescent signal is reconstituted3-6. Because its signal is readily detectable by epifluorescence or confocal microscopy, BiFC has emerged as a powerful tool of choice among cell biologists for studying about protein-protein interactions in living cells 3. This assay, however, can sometimes produce false positive results. For example, the fluorescent signal can be reconstituted by two GFP fragments arranged as far as 7 nm from each other due to close packing in a small subcellular compartment, rather that due to specific interactions7.
Due to these limitations, the results obtained from live cell imaging technologies should be confirmed by an independent approach based on a different principle for detecting protein interactions. Co-immunoprecipitation (Co-IP) or glutathione transferase (GST) pull-down assays represent such alternative methods that are commonly used to analyze protein-protein interactions in vitro. However, iIn these assays, however, the tested proteins must be readily soluble in the buffer that supportsused for the binding reaction. Therefore, specific interactions involving an insoluble protein cannot be assessed by these techniques.
 Here, we illustrate the protocol for the protein membrane overlay binding assay, which circumvents this difficulty. In this technique, interaction between soluble and insoluble proteins can be reliably tested because one of the proteins is immobilized on a membrane matrix. This method, in combination with in vivo experiments, such as BiFC, provides a reliable approach to investigate and characterize interactions faithfully between soluble and insoluble proteins. In this article, binding between Tobacco mosaic virus (TMV) movement protein (MP), which exerts multiple functions during viral cell-to-cell transport8-14, and a recently identified plant cellular interactor, tobacco ankyrin repeat-containing protein (ANK) 15, is demonstrated using this technique.

Protein Membrane Overlay Assay: A Protocol to Test Interaction Between Soluble and Insoluble Proteins in vitro

Testing protein-protein interaction is indispensable for dissection of protein functionality. Here, we introduce an in vitro protein-protein binding assay to probe a membrane-immobilized protein with a soluble protein. This assay provides a reliable method to test interaction between an insoluble protein and a protein in solution.

Validating interactions between different proteins is vital for investigation of their biological functions on the molecular level.  There are several ...

Biology

Membrane Transport Processes Analyzed by a Highly Parallel Nanopore Chip System at Single Protein Resolution

Membrane protein transport on the single protein level still evades detailed analysis, if the substrate translocated is non-electrogenic. Considerable efforts have been made in this field, but techniques enabling automated high-throughput transport analysis in combination with solvent-free lipid bilayer techniques required for the analysis of membrane transporters are rare. This class of transporters however is crucial in cell homeostasis and therefore a key target in drug development and methodologies to gain new insights desperately needed.
The here presented manuscript describes the establishment and handling of a novel biochip for the analysis of membrane protein mediated transport processes at single transporter resolution. The biochip is composed of microcavities enclosed by nanopores that is highly parallel in its design and can be produced in industrial grade and quantity. Protein-harboring liposomes can directly be applied to the chip surface forming self-assembled pore-spanning lipid bilayers using SSM-techniques (solid supported lipid membranes). Pore-spanning parts of the membrane are freestanding, providing the interface for substrate translocation into or out of the cavity space, which can be followed by multi-spectral fluorescent readout in real-time. The establishment of standard operating procedures (SOPs) allows the straightforward establishment of protein-harboring lipid bilayers on the chip surface of virtually every membrane protein that can be reconstituted functionally. The sole prerequisite is the establishment of a fluorescent read-out system for non-electrogenic transport substrates.
High-content screening applications are accomplishable by the use of automated inverted fluorescent microscopes recording multiple chips in parallel. Large data sets can be analyzed using the freely available custom-designed analysis software. Three-color multi spectral fluorescent read-out furthermore allows for unbiased data discrimination into different event classes, eliminating false positive results.
The chip technology is currently based on SiO2 surfaces, but further functionalization using gold-coated chip surfaces is also possible.

The presented protocol describes the analysis of membrane protein mediated transport on the single transporter level using pore-spanning solvent-free lipid bilayers. This is achieved by the creation of bulk produced nanopore array chips, combined with highly parallel data acquisition and analysis, enabling the future establishment of membrane protein effector screenings.

Membrane protein transport on the single protein level still evades detailed analysis, if the substrate translocated is non-electrogenic. Considerable ...

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

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