The authors thank the Swedish Research Council, Stockholm County Council, and KI funds for their support. The expression and purification of MSP was performed at the Karolinska Institutet/SciLifeLab Protein Science Core Facility (http://PSF.ki.se). The authors would also like to thank Dr. Pasi Purhonen and Dr. Mathilda Sj&#246;berg for sharing their technical expertise and for their timely assistance.

1. Preparation of Nanodiscs
<ol>
	<li>Expression and purification of the membrane scaffolding protein8,35
	<ol>
		<li>Express the His-tagged MSP1E3D1 in the E. coli BL21 (DE3) T1R pRARE2 strain in flasks. Prepare a 50-mL overnight starter culture with LB medium supplemented with 50 &#181;g/mL Kanamycin at 37 &#176;C. Dilute the overnight starter culture in 2 L of terrific broth medium supplemented with 50 &#181;g/mL kanamycin.</li>
		<li>Grow the cells at 37 &#176;C until the optical density at 600 nm (OD600) reaches approximately 3. Induce the protein expression with 0.5 mM isopropyl &#946;-D-1-thiogalactopyranoside (IPTG) for 3 h at 18 &#176;C.</li>
		<li>Prepare the lysis buffer (100 mM Tris-HCl, pH 8.0; 100 mM NaCl; and 10% glycerol). Add 1 mM TCEP immediately before use.</li>
		<li>After 3 h of induction at 18 &#176;C, harvest the cells by centrifugation for 10 min at 4,500 x g and 4 &#176;C. Discard the supernatant, weigh the harvested cells, and re-suspend them in lysis buffer at a ratio of 2 mL of lysis buffer per g of cells.</li>
		<li>Lyse the cells by pulsed sonication (repetition rate: 4 s ON, 4 s OFF) for 3 min at 80% amplitude.</li>
		<li>Centrifuge the lysates for 20 min at 49,000 x g and 4 &#176;C. Discard the pellet from this centrifugation.</li>
		<li>Inject the supernatant into a 5-mL Ni+ chelating column connected to an automated liquid chromatography system preferably kept at 4 &#176;C.</li>
		<li>Before the elution step (step 1.1.9), wash the column with buffers 1 - 3 below to remove all proteins except the His-tagged MSP. Monitor the protein content at 280 nm to follow the purification process; the UV recording at 280 nm should go back to baseline after each wash.
		<ol>
			<li>Wash with wash buffer 1: 40 mM Tris-HCl, 300 mM NaCl, 10 mM imidazole, and 1% Triton, pH 8.0.</li>
			<li>Wash with wash buffer 2: 40 mM Tris-HCl, 300 mM NaCl, 20 mM imidazole, and 50 mM sodium cholate, pH 8.0.</li>
			<li>Wash with wash buffer 3: 40 mM Tris-HCl, 300 mM NaCl, and 50 mM imidazole, pH 8.0.</li>
		</ol>
		</li>
		<li>Elute the MSP with 40 mM Tris-HCl, 300 mM NaCl, and 500 mM imidazole, pH 8.0.</li>
		<li>Change the buffer to MSP standard buffer (20 mM Tris-HCl, pH 7.5; 100 mM NaCl; and 0.5 mM EDTA) by gel filtration8,35; the yield per batch should be around 7 mg L-1.</li>
	</ol>
	</li>
	<li>Preparation of phospholipid stock solution
	<ol>
		<li>Add 305 &#181;L of 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC) dissolved in chloroform at a concentration of 25 mg/mL to a glass beaker and evaporate the chloroform by purging it with a gentle stream of nitrogen. Dry the lipid overnight in a vacuum desiccator. NOTE: This step is performed to remove the solvent from the lipid, which leaves a thin film of lipid at the bottom of the glass beaker.</li>
		<li>Resuspend the lipid cake in 200 &#181;L of MSP standard buffer containing 100 mM sodium cholate by vortexing the tube until the solution becomes transparent; this will provide a solution with a POPC concentration of 50 mM. 
		NOTE: In general, the molar ratio of lipid to detergent at this point should be 1:2 (lipid:detergent)8. This lipid-detergent mix can be stored at -80 &#176;C for almost 2 months.</li>
	</ol>
	</li>
	<li>Preparation of hydrophobic beads for detergent removal
	<ol>
		<li>Place 5 g of beads (dry-weight) in a 50-mL tube.</li>
		<li>Wash the beads with 30 mL of 100% methanol and then with 40 mL of ultrapure water.</li>
		<li>Wash the beads with 10 mL of MSP standard buffer. Finally, store the beads with 15 mL of MSP standard buffer at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Nanodisc reconstitution
	<ol>
		<li>Dispense 190 &#181;L of MSP1E3D1 (0.124 mM) into a microfuge tube and add 61.5 &#181;L of phospholipid stock solution (50 mM POPC) to the same tube. Incubate the reconstitution mixture on wet ice for 1 h. NOTE: This equals a molar ratio of 1:130 (MSP1E3D1: POPC).</li>
		<li>Add hydrophobic beads at a concentration of 0.5 g of beads per mL of reconstitution mixture to initiate the self-assembly process. Incubate in a rotary incubator for 16 h at 4 &#176;C.</li>
		<li>After incubation, centrifuge for 10 min at 13,000 x g and 4 &#176;C to remove the precipitates and aggregates. Discard the pellet and save the supernatant.</li>
		<li>Equilibrate the size exclusion chromatography column (mounted on an automated liquid chromatography system) with MSP standard buffer until the absorbance at 280 nm is stable. Inject the supernatant into the column and collect the peak fractions.</li>
		<li>Measure the concentrations of nanodiscs in the peak fractions at 280 nm. Use the molar extinction coefficient of MSP1E3D1 (&#949; = 29,910 cm-1 M-1) for the calculation of the nanodisc concentration. NOTE: The number of lipids per nanodisc can be measured by a combination of radiolabeled lipids and phosphate analysis4 or by phosphate analysis alone.</li>
	</ol>
	</li>
</ol>
2. Preparation of the Monotopic Protein 5-Lipoxygenase35
<ol>
	<li>Prepare an overnight starter culture. Supplement 50 mL of LB medium with 100 &#181;g/mL ampicillin. Inoculate the medium with E. coli BL21 (DE3) containing the plasmid of the 5-lipoxygenase gene (ALOX5). Dilute the overnight starter culture in expression medium containing 42 mM Na2HPO4, 24 mM KH2PO4, 9 mM NaCl, 19 mM NH4Cl, 1 mM MgSO4, 0.1 mM CaCl2, 0.2% D-glucose, 0.1%, 5 &#181;M FeSO4, and 100 &#181;g/mL ampicillin. Grow the cells until the OD600 is ~ 0.5 at 25 &#176;C. Induce protein expression with 0.2 mM IPTG for 16 h at 20 &#176;C35.</li>
	<li>Harvest by centrifugation for 10 min at 7,000 x g and 4 &#176;C and discard the supernatant. Weigh the pellet found at the bottom of the tube containing the harvested cells and resuspend the harvested cells in lysis buffer (100 mM Tris-HCl, pH 8.0; 100 mM NaCl; 10% glycerol; and 1 mM TCEP) containing protease inhibitor and 0.5 mg/mL35 lysozyme at a ratio of 2 mL of lysis buffer per g of cells.</li>
	<li>Lyse the cells by sonication for 5 x 15 s at 80% amplitude. Remove the cell debris by centrifugation for 10 min at 7,000 x g and 4 &#176;C. Perform ammonium sulfate precipitation to 30 - 60% saturation to precipitate the proteins in the solution35. Centrifuge for 15 min at 16,000 x g and 4 &#176;C. NOTE: The 5LO pellet can be rapidly frozen and stored at -80 &#176;C for up to 6 months.</li>
	<li>Resuspend the pellet with 20 mL of lysis buffer and centrifuge for 15 min at 40,000 x g and 4 &#176;C.</li>
	<li>Incubate the supernatant on an ATP agarose column at 4 &#176;C for 30 min. Wash the column once with one column-volume of lysis buffer containing 0.5 M NaCl. Elute the 5LO with 20 mM ATP in lysis buffer containing 10 &#181;M FeSO4 and 20 &#181;g/mL catalase. Perform gel filtration chromatography to remove the ATP35. 
	NOTE: The 5LO is unstable and should be used immediately after purification. Otherwise, it is recommended to halt at the step of ammonium sulphate precipitation (see the note after step 2.1.3).</li>
</ol>
3. Preparation of the Nanodisc-protein Complex
<ol>
	<li>Prepare a total volume of 100 &#181;L of complex. Mix 0.8 &#181;M ND and 0.8 &#181;M 5LO with the 1 mM Ca2+ present in the MSP standard buffer (20 mM Tris-HCl, pH 7.5; 100 mM NaCl; 0.5 mM EDTA; and 1.5 mM CaCl2) and incubate for 10 min on ice. NOTE: The sample can be stored for up to one month at 4 &#176;C35.</li>
</ol>
4. Analysis of the Samples
<ol>
	<li>Gel electrophoresis
	<ol>
		<li>Non-denaturing electrophoresis
		<ol>
			<li>Mix 15 &#181;L of the sample with 5 &#181;L of loading buffer (50 mM BisTris, 6 N HCl, 50 mM NaCl, 10% (w/v) glycerol, and 0.001% Ponceau S, pH 7.2)49 and load it on a 4 - 16% Bis-Tris gel to perform electrophoresis.</li>
			<li>Fill the cathode tank with light cathode buffer (50 mM Bis Tris; 50 mM Tricine, pH 6.8; and 0.002% Coomassie G-250) and the anode tank with running buffer (50 mM Bis Tris and 50 mM Tricine, pH 6.8). Start the separation by using a constant voltage at 150 V.</li>
			<li>Stop the electrophoretic separation when the Coomassie front reaches the end of the gel.</li>
			<li>Stain the gel with a standard Coomassie blue protocol.</li>
		</ol>
		</li>
		<li>Denaturing electrophoresis
		<ol>
			<li>Mix 40 &#181;L of the sample with 10 &#181;L of loading buffer containing SDS (0.05% (w/v) Bromophenol blue; 0.2 M Tris-HCl, pH 6.8; 20% (v/v) glycerol; 10% (w/v) SDS; and 10 mM 2-mercaptoethanol) and load it on a 4 - 20% Tris glycine gel to perform electrophoresis.</li>
			<li>Fill the cathode and anode tanks with running buffer (25 mM Tris-HCl, pH 6.8; 200 mM glycine; and 0.1% (w/v) SDS). Start the separation by using a constant voltage at 150 V.</li>
			<li>Stop the electrophoretic separation when the loading dye front reaches the end of the gel.</li>
			<li>Stain the gel with a standard Coomassie blue protocol.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Preparation of the NaPT solution
	<ol>
		<li>Dissolve 1 g of phosphotungstate sodium salt in 50 mL of water by agitation at room temperature to give a 2% (w/v) acidic solution.</li>
		<li>Adjust the pH to 7.4 with 1 M NaOH. Remove the particles using a 0.22-&#181;m syringe filter. Store the solution at room temperature or at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Preparation of the sample for TEM analysis
	<ol>
		<li>Glow-discharge carbon-coated copper grids (400 mesh) for 20 s at 30 mA to render the grids hydrophilic before the adsorption of the sample. Place 3.5 &#181;L of the sample (0.8 &#181;M with respect to the nanodiscs) on the grid and incubate for 30 s. NOTE: The volume applied may vary between 2.5 and 5 &#181;L, depending on the sample concentration. A suitable concentration range for nanodiscs is 0.5 - 1 &#181;M.</li>
		<li>Blot off surplus solution using filter paper.</li>
		<li>Immediately stain the grid with a drop of 2% NaPT for 30 s. Blot off excess solution and leave the grid to air dry.</li>
		<li>Evaluate the grids by TEM. A microscope with 120-200 keV accelerating voltage suffices to estimate the extent of stacking. NOTE: For the present protocol, a calibrated transmission electron microscope was used, equipped with a 200-keV field emission gun.</li>
		<li>Record the TEM images. For the images showing long stacks, do not process further; images show the complex of nanodiscs, with extrinsic protein that may contain a few short stacks, but most particles are in the complex. Record several images and process these according to standard methods to obtain class-averages and a low-resolution, 3 dimensional model of the complex. 
		NOTE: For the collection of negative-stain data, grids were made on at least three different days. For each day, fresh sample incubations (as in step 3.1) were made before the negative stain was applied.</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

The method can be separated into three parts: the reconstitution of empty nanodiscs, the preparation of protein-nanodisc complexes, and the negative staining for the TEM of these complexes. Each part will be addressed separately regarding limitations of the technique, critical steps, and useful modifications.
Reconstitution of empty nanodiscs. Critical steps and limitations in the production and use of nanodiscs.
For the preparation of the empty nanodiscs, it is essent...

In medical research, much attention is focused on membrane proteins, either intrinsic or extrinsic, involved in a variety of lipid interactions. Working with lipid-interacting proteins includes either selecting a substitute to the lipids, such as detergents, amphipols1, or small proteins2, or finding a membrane substitute that keeps the protein soluble and active. Lipoic membrane substitutes include liposomes and nanodiscs (ND)3,4.
Nanodiscs are near-native membrane platforms developed by engineering the protein part, ApoA-1, of the high-density lipoprotein (HDL) naturally occurring in blood. ApoA-1 is a 243 residue-long chain of short amphipathic &#945;-helices and has a lipid-free soluble conformation. In vitro when in the presence of lipids, two copies of the protein ApoA-1 spontaneously rearrange to encircle the hydrophobic acyl chain portion of a lipid bilayer patch5. Engineered versions of ApoA-1 are generally called membrane scaffolding proteins (MSP), and an increasing number are commercially available as plasmids or as purified proteins. Repetitions or deletions of the &#945;-helices in ApoA-1 result in longer6 or shorter7 membrane scaffolding proteins. This in turn makes it possible to form discs around 6 nm7 to 17 nm8 in diameter. There are different types of applications for the nanodiscs3,9. The most commonly used application is to provide a near-native membrane environment for the stabilization of an integral membrane protein8, reviewed previously3,9. A less-explored use is to provide a nanoscale membrane surface for the study of peripheral membrane proteins10,11,12,13,14,15,16,17. Section 1 of the protocol below visualizes the procedure for making nanodiscs composed of phospholipids and membrane scaffolding protein.
Sample preparation is a bottleneck in most methods. Method-specific samples may add particular information, but they also make comparisons of results difficult. Therefore, it is simpler when samples are multimodal and can be used directly in several different methods. One advantage with the use of nanodiscs is the small size of the nanodisc in comparison to liposomes (e.g., the samples can be directly used for both TEM and non-denaturing gel electrophoresis, as in the present protocol).
Vesicles and liposomes have long been used to understand the function of membrane-interacting proteins. For structural studies and visualization, an example of the structural determination of a transmembrane protein in liposomes is available18. However, no high-resolution 3D structure of a monotopic membrane protein embedded on a liposome membrane has been published yet, as far as we know. Gold nanoparticles or antibodies can be used to visualize proteins binding to liposomes or vesicles using TEM19. Even though these probes are very specific, they might interfere with membrane-binding proteins by veiling the membrane binding site or by masking areas of interest with the flexible parts. Gold-labeled or antibody-complexed proteins could probably be analyzed on a gel, but this would increase the cost of the experiment.
Though liposomes are an excellent platform, one cannot be certain that the population has a particular ratio of protein per liposome, a feature that can be explored by the use of nanodiscs20. In a liposome, cofactors and substrates can be trapped in the soluble interior. Substances that are membrane-soluble will share the same fate for both types of membrane mimetics. Nevertheless, as the bilayer area is smaller in nanodiscs, a smaller amount of substance is required to saturate the nanodisc membranes.
Understanding protein function through the determination of the atomic structure has been essential for many fields of research. Methods for protein structure determination include X-ray21; nuclear magnetic resonance (NMR)22,23; and transmission electron microscopy (TEM)24 at cryogenic temperatures, cryoEM. The resolution by cryoEM has lately been greatly improved, mainly due to the use of direct electron detectors25,26. The macromolecules are imaged in thin, vitreous ice27 in a near-native state. However, due to the low contrast of biological molecules, they become hard to detect in the size range of 100 - 200 kDa. For suitably sized samples, data collection can be made and the method of single particle reconstruction can be applied to obtain a structure28.
However, the determination of protein structure by TEM is a multistep process. It usually starts with the evaluation of sample monodispersity by negative-stain TEM29 using salts of heavy metals like phosphotungsten (PT)30 or uranium31. Reconstruction of a low-resolution model of the negatively stained macromolecule is usually made and may yield important information on the molecular structure29. In parallel, data collection using cryoEM may start. Care should be taken when evaluating negative-stain TEM data to avoid the misinterpretation of artefact formation. One particular artefact is the effect of the PT stain on phospholipids and liposomes32, resulting in the formation of long rods resembling stacks of coins viewed from the side33. Such &#34;rouleau&#34; or &#34;stacks&#34; (hereafter denoted as &#34;stacks&#34;) were observed early on for HDL34, and later also for nanodiscs35.
The stacking and reshaping of membranes may occur for many reasons. For example, it can be induced by co-factors like copper, shown by TEM imaging in an ammonium molybdate stain36. A fraction of the membrane lipids in liposomes contained an iminodiacetic acid head group mimicking metal complexation by EDTA, thus stacking liposomes after the addition of copper ions36. Stacking could also be due to a protein-protein interaction by a protein in or on the lipid bilayers (the stain used is not mentioned)37. The stack formation of phospholipids by PT was observed early on; however, later work has focused on removing or abolishing this artifact formation38.
Here, we propose a method to take advantage of the NaPT-induced nanodisc stacking for the study of membrane-binding proteins by TEM. In short, protein binding on the nanodiscs would prevent the nanodiscs from stacking. Though the reasons for the stacking are not clear, it was proposed39 that there is an electrostatic interaction between the phospholipids and the phosphoryl group of PT, causing the discs to stick to each other (Figure 1A). The hypothesis behind our protocol is that when a protein binds to a nanodisc, most of the phospholipid surface is not available for the interaction with the PT due to steric hindrance by the protein. This would prevent stack formation (Figure 1B). Two conclusions can be drawn. First, the prevention of stacking means that the protein of interest has bound to the membrane. Secondly, the protein-ND complex can be treated with standard single-particle processing methods24,40 to get a rough morphology of the complex. Furthermore, analyses by methods like non-denaturing gel electrophoresis or dynamic light scattering can be performed.
To demonstrate this hypothesis, we used the membrane-binding protein 5-lipoxygenase (5LO), which is involved in many inflammatory diseases41,42. This 78-kDa protein requires calcium ions to bind to its membrane43. Though this membrane association has been studied extensively using liposomes44,45,46 and membrane fractions47, these cannot be used for TEM analysis and structure determination.
The preparation of nanodiscs starts by mixing MSP with lipid resuspended in the detergent sodium cholate. After incubation on ice for 1 h, the detergent is slowly removed from the reconstitution mixture using an adsorbent resin. This kind of material is frequently made of polystyrene shaped into small beads. They are relatively hydrophobic and have a strong preference for binding detergent compared to lipids48. After removing the hydrophobic beads and performing clarification using centrifugation, the nanodiscs are purified by size exclusion chromatography (SEC). The purified nanodiscs are mixed with a monotopic membrane protein (and possible cofactors) in an equimolar ratio (or several ratios for a titration) and are left to react (15 min). Analysis by TEM is carried out by applying &#181;L-amounts of sample onto glow-discharged, carbon-coated grids and then by performing negative staining with NaPT. The same sample from when the aliquots were applied to the TEM grids can be used for analysis by non-denaturing or SDS PAGE gel-electrophoresis, as well as by various kinds of activity measurements, with no major changes.

<table><tbody><tr><td>Transmission electron microscope: JEOL2100F</td><td>JEOL</td><td></td><td></td></tr><tr><td>CCD camera</td><td>Tiez Video and Imaging Processing System GmbH, Germany</td><td></td><td></td></tr><tr><td>Glow discharger</td><td>Baltec</td><td></td><td></td></tr><tr><td>TEM grid: 400 mesh</td><td>TAAB</td><td>GM016/C</td><td></td></tr><tr><td>Size exclusion chromatography: Agilent SEC-5</td><td>Agilent Technologies</td><td>5190-2526</td><td></td></tr><tr><td>Superdex 200 HR 10/300</td><td>GE Healthcare Life Sciences</td><td>17-5172-01</td><td></td></tr><tr><td>Plasmid: MSP1E3D1</td><td>Addgene</td><td>20066</td><td></td></tr><tr><td>Bacteria: BL21DE3</td><td>NEB</td><td>C2527H</td><td></td></tr><tr><td>Bacteria: BL21 (DE3) T1R pRARE2</td><td>Protein Science Facility, KI, Solna</td><td></td><td></td></tr><tr><td>Purification Matrix: ATP agarose</td><td>Sigma Aldrich</td><td>A2767</td><td></td></tr><tr><td>Purification Matrix: HisTrap HP-5 mL</td><td>GE Healthcare Life Sciences</td><td>17-5247-01</td><td></td></tr><tr><td>Lipid: POPC</td><td>Avanti polar lipids</td><td>850457C</td><td>25 mg/mL in chloroform</td></tr><tr><td>Hydrophobic beads: Bio-Beads, SM-2 Resin</td><td>Bio-Rad</td><td>1523920</td><td></td></tr><tr><td>13 mm syringe filter: 0.2 &amp;mu;m</td><td>Pall life sciences</td><td>PN 4554T</td><td></td></tr><tr><td>Stain: Sodium phosphotungstate tribasic hydrate</td><td>Sigma Aldrich</td><td>31648</td><td></td></tr><tr><td>2-mercaptoethanol</td><td>Sigma Aldrich</td><td>M3148-250ML</td><td></td></tr><tr><td>Sodium Dodecyl Sulfate (SDS)</td><td>Bio-Rad</td><td>161-0301</td><td></td></tr><tr><td>Protease inhibitor cocktail</td><td>Sigma Aldrich</td><td>4693132001</td><td></td></tr><tr><td>TCEP</td><td>Sigma Aldrich</td><td>646547</td><td></td></tr><tr><td>Detergent: Sodium cholate hydrate</td><td>Sigma Aldrich</td><td>C6445-10G</td><td></td></tr><tr><td>Sodium Cholate</td><td></td><td></td><td>500 mM Sodium cholate. Resuspend in miliQ water and store at -20 &amp;deg;C.</td></tr><tr><td>Lipid Stock</td><td></td><td></td><td>50 mM POPC, 100 mM sodium cholate, 20 mM Tris-HCl pH 7.5, 100 mM NaCl. &lt;bt /&gt;Store at 4 &amp;deg;C for a week; or&lt;br/&gt; Store -80 &amp;deg;C for a month, after purging the solution with nitrogen.</td></tr><tr><td>MSP standard buffer</td><td></td><td></td><td>20 mM Tris-HCl, pH 7.5, 100 mM NaCl, 0.5 mM EDTA. &lt;br /&gt;Store at 4 &amp;deg;C.</td></tr><tr><td>Non-Denaturaing Electrophoresis Anode Buffer</td><td>Thermo Fisher Scientific</td><td>BN2001</td><td>50 mM Bis-Tris, 50 mM Tricine, pH 6.8</td></tr><tr><td>Non-Denaturaing Electrophoresis Cathode Buffer</td><td>Thermo Fisher Scientific</td><td>BN2002</td><td>50 mM Bis-Tris, 50 mM Tricine, pH 6.8, 0.002% Coomassie G-250</td></tr><tr><td>Non-Denaturaing Electrophoresis 4x Sample loading Buffer</td><td>Thermo Fisher Scientific</td><td>BN2003</td><td>50 mM Bis-Tris, pH 7.2, 6 N HCl, 50 mM NaCl, 10% (w/v) glycerol, 0.001% Ponceau S</td></tr><tr><td>Denaturaing Electrophoresis Running Buffer</td><td></td><td></td><td>In-house recipe: 25 mM Tris-HCl, pH 6.8, 200 mM Glycine, 0.1% (w/v) SDS</td></tr><tr><td>Denaturaing Electrophoresis 5x Sample loading Buffer</td><td></td><td></td><td>In-house recipe: 0.05% (w/v) Bromophenolblue, 0.2 M Tris-HCl, pH 6.8, 20% (v/v) glycerol, 10% (w/v) SDS, 10 mM 2-mercaptoethanol</td></tr><tr><td>Terrific broth</td><td></td><td></td><td>Tryptone - 12.0 g, Yeast Extract - 24.0 g, 100 mL 0.17 M KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt; and 0.72 M K&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;, Glycerol - 4 mL.&lt;br /&gt;Tryptone, yeast extract and glycerol were prepared to 900 mL and autoclaved seperately. KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt; and K&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt; were prepared and autoclaved separately. Both were mixed before using the medium.</td></tr></tbody></table>

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.

The method we propose depends upon the preparation of nanodiscs to provide the membrane surface for monotopic membrane-protein binding. As there is no transmembrane protein embedded into the nanodisc lipid bilayer, the nanodiscs are here denoted as &#34;empty nanodiscs&#34; (Figure 2A). These have a calculated molecular weight of 256 kDa for a composition of two MSP1E3D1 scaffolding proteins and around 260 molecules of POPC8. Using this protein:lip...

Watch this Scientific Journal Video about Method to Visualize and Analyze Membrane Interacting Proteins by Transmission Electron Microscopy at JoVE.com

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

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

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 ...

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13.1K Views.  Karolinska Institutet. 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.

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

Visualizing Proteins and Macromolecular Complexes by Negative Stain EM: from Grid Preparation to Image Acquisition

Single particle electron microscopy (EM), of both negative stained or frozen hydrated biological samples, has become a versatile tool in structural biology 1. In recent years, this method has achieved great success in studying structures of proteins and macromolecular complexes 2, 3. Compared with electron cryomicroscopy (cryoEM), in which frozen hydrated protein samples are embedded in a thin layer of vitreous ice 4, negative staining is a simpler sample preparation method in which protein samples are embedded in a thin layer of dried heavy metal salt to increase specimen contrast 5. The enhanced contrast of negative stain EM allows examination of relatively small biological samples. In addition to determining three-dimensional (3D) structure of purified proteins or protein complexes 6, this method can be used for much broader purposes. For example, negative stain EM can be easily used to visualize purified protein samples, obtaining information such as homogeneity/heterogeneity of the sample, formation of protein complexes or large assemblies, or simply to evaluate the quality of a protein preparation.
In this video article, we present a complete protocol for using an EM to observe negatively stained protein sample, from preparing carbon coated grids for negative stain EM to acquiring images of negatively stained sample in an electron microscope operated at 120kV accelerating voltage. These protocols have been used in our laboratory routinely and can be easily followed by novice users.

Visualizing protein samples by negative stain electron microscopy (EM) has become a popular structural analysis method. It is useful for quantitative structural analysis, such as calculating a 3D reconstruction of the molecules being studied, and also for qualitative examination of the quality of protein preparations. In this article we present detailed protocols for preparing the EM grids, staining the sample and visualizing the sample in an electron microscope. Novice users can follow these protocols easily and to utilize negative stain EM as a routine assay, in addition to other biochemical assays, for evaluating their protein samples.

Single particle electron microscopy (EM), of both negative stained or frozen hydrated biological samples, has become a versatile tool in structural ...

Bioengineering

Analyzing Dynamic Protein Complexes Assembled On and Released From Biolayer Interferometry Biosensor Using Mass Spectrometry and Electron Microscopy

In vivo, proteins are often part of large macromolecular complexes where binding specificity and dynamics ultimately dictate functional outputs. In this work, the pre-endosomal anthrax toxin is assembled and transitioned into the endosomal complex. First, the N-terminal domain of a cysteine mutant lethal factor (LFN) is attached to a biolayer interferometry (BLI) biosensor through disulfide coupling in an optimal orientation, allowing protective antigen (PA) prepore to bind (Kd 1 nM). The optimally oriented LFN-PAprepore complex then binds to soluble capillary morphogenic gene-2 (CMG2) cell surface receptor (Kd 170 pM), resulting in a representative anthrax pre-endosomal complex, stable at pH 7.5. This assembled complex is then subjected to acidification (pH 5.0) representative of the late endosome environment to transition the PAprepore into the membrane inserted pore state. This PApore state results in a weakened binding between the CMG2 receptor and the LFN-PApore and a substantial dissociation of CMG2 from the transition pore. The thio-attachment of LFN to the biosensor surface is easily reversed by dithiothreitol. Reduction on the BLI biosensor surface releases the LFN-PAprepore-CMG2 ternary complex or the acid transitioned LFN-PApore complexes into microliter volumes. Released complexes are then visualized and identified using electron microscopy and mass spectrometry. These experiments demonstrate how to monitor the kinetic assembly/disassembly of specific protein complexes using label-free BLI methodologies and evaluate the structure and identity of these BLI assembled complexes by electron microscopy and mass spectrometry, respectively, using easy-to-replicate sequential procedures.

Here we present a protocol to monitor the assembly and disassembly of the anthrax toxin using biolayer interferometry (BLI). Following assembly/disassembly on the biosensor surface, the large protein complexes are released from the surface for visualization and identification of components of the complexes using electron microscopy and mass spectrometry, respectively.

In vivo, proteins are often part of large macromolecular complexes where binding specificity and dynamics ultimately dictate functional outputs. In this ...

Mitochondria and Endoplasmic Reticulum Imaging by Correlative Light and Volume Electron Microscopy

Cellular organelles, such as mitochondria and endoplasmic reticulum (ER), create a network to perform a variety of functions. These highly curved structures are folded into various shapes to form a dynamic network depending on the cellular conditions. Visualization of this network between mitochondria and ER has been attempted using super-resolution fluorescence imaging and light microscopy; however, the limited resolution is insufficient to observe the membranes between the mitochondria and ER in detail. Transmission electron microscopy provides good membrane contrast and nanometer-scale resolution for the observation of cellular organelles; however, it is exceptionally time-consuming when assessing the three-dimensional (3D) structure of highly curved organelles. Therefore, we observed the morphology of mitochondria and ER via correlative light-electron microscopy (CLEM) and volume electron microscopy techniques using enhanced ascorbate peroxidase 2 and horseradish peroxidase staining. An en bloc staining method, ultrathin serial sectioning (array tomography), and volume electron microscopy were applied to observe the 3D structure. In this protocol, we suggest a combination of CLEM and 3D electron microscopy to perform detailed structural studies of mitochondria and ER.

We present a protocol to study the distribution of mitochondria and endoplasmic reticulum in whole cells after genetic modification using correlative light and volume electron microscopy including ascorbate peroxidase 2 and horseradish peroxidase staining, serial sectioning of cells with and without the target gene in the same section, and serial imaging via electron microscopy.

Cellular organelles, such as mitochondria and endoplasmic reticulum (ER), create a network to perform a variety of functions. These highly curved ...

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 ...

Freeze-Fracture Electron Microscopy for Extracellular Vesicle Analysis

Extracellular vesicles (EVs) are membrane-limited structures released from the cells into the extracellular space and are implicated in intercellular communication. EVs consist of three populations of vesicles, namely microvesicles (MVs), exosomes, and apoptotic bodies. The limiting membrane of EVs is crucially involved in the interactions with the recipient cells, which could lead to the transfer of biologically active molecules to the recipient cells and, consequently, affect their behavior. The freeze-fracture electron microscopy technique is used to study the internal organization of biological membranes. Here, we present a protocol for MV isolation from cultured cancerous urothelial cells and the freeze-fracture of MVs in the steps of rapid freezing, fracturing, making and cleaning the replicas, and analyzing them with transmission electron microscopy. The results show that the protocol for isolation yields a homogenous population of EVs, which correspond to the shape and size of MVs. Intramembrane particles are found mainly in the protoplasmic face of the limiting membrane. Hence, freeze-fracture is the technique of choice to characterize the MVs' diameter, shape, and distribution of membrane proteins. The presented protocol is applicable to other populations of EVs.

We present a protocol for the isolation and freeze-fracturing of extracellular vesicles (EVs) originating from cancerous urothelial cells. The freeze-fracture technique revealed the EVs' diameter and shape and-as a unique feature-the internal organization of the EV membranes. These are of immense importance in understanding how EVs interact with the recipient membranes.

Extracellular vesicles (EVs) are membrane-limited structures released from the cells into the extracellular space and are implicated in intercellular ...

Cancer Research

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

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

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

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

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

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

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

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

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

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

Visualization of ATP Synthase Dimers in Mitochondria by Electron Cryo-tomography

Electron cryo-tomography is a powerful tool in structural biology, capable of visualizing the three-dimensional structure of biological samples, such as cells, organelles, membrane vesicles, or viruses at molecular detail. To achieve this, the aqueous sample is rapidly vitrified in liquid ethane, which preserves it in a close-to-native, frozen-hydrated state. In the electron microscope, tilt series are recorded at liquid nitrogen temperature, from which 3D tomograms are reconstructed. The signal-to-noise ratio of the tomographic volume is inherently low. Recognizable, recurring features are enhanced by subtomogram averaging, by which individual subvolumes are cut out, aligned and averaged to reduce noise. In this way, 3D maps with a resolution of 2 nm or better can be obtained. A fit of available high-resolution structures to the 3D volume then produces atomic models of protein complexes in their native environment. Here we show how we use electron cryo-tomography to study the in situ organization of large membrane protein complexes in mitochondria. We find that ATP synthases are organized in rows of dimers along highly curved apices of the inner membrane cristae, whereas complex I is randomly distributed in the membrane regions on either side of the rows. By subtomogram averaging we obtained a structure of the mitochondrial ATP synthase dimer within the cristae membrane.

We present a protocol of how to collect and process electron cryo-tomograms of whole mitochondria. The technique provides detailed insights into the structure, function, and organization of large membrane protein complexes in native biological membranes.

Electron cryo-tomography is a powerful tool in structural biology, capable of visualizing the three-dimensional structure of biological samples, such as ...

Determining Membrane Protein Topology Using Fluorescence Protease Protection (FPP)

The correct topology and orientation of integral membrane proteins are essential for their proper function, yet such information has not been established for many membrane proteins. A simple technique called fluorescence protease protection (FPP) is presented, which permits the determination of membrane protein topology in living cells. This technique has numerous advantages over other methods for determining protein topology, in that it does not require the availability of multiple antibodies against various domains of the membrane protein, does not require large amounts of protein, and can be performed on living cells. The FPP method employs the spatially confined actions of proteases on the degradation of green fluorescent protein (GFP) tagged membrane proteins to determine their membrane topology and orientation. This simple approach is applicable to a wide variety of cell types, and can be used to determine membrane protein orientation in various subcellular organelles such as the mitochondria, Golgi, endoplasmic reticulum and components of the endosomal/recycling system. Membrane proteins, tagged on either the N-termini or C-termini with a GFP fusion, are expressed in a cell of interest, which is subject to selective permeabilization using the detergent digitonin. Digitonin has the ability to permeabilize the plasma membrane, while leaving intracellular organelles intact. GFP moieties exposed to the cytosol can be selectively degraded through the application of protease, whereas GFP moieties present in the lumen of organelles are protected from the protease and remain intact. The FPP assay is straightforward, and results can be obtained rapidly.

Determining Membrane Protein Topology Using Fluorescence Protease Protection FPP

Here, we present a protocol to determine the orientation and topology of integral membrane proteins in living cells. This simple protocol relies on selective protease sensitivity of chimeras between the protein of interest and GFP.

The correct topology and orientation of integral membrane proteins are essential for their proper function, yet such information has not been established ...

Miniaturized Sample Preparation for Transmission Electron Microscopy

Due to recent technological progress, cryo-electron microscopy (cryo-EM) is rapidly becoming a standard method for the structural analysis of protein complexes to atomic resolution. However, protein isolation techniques and sample preparation methods for EM remain a bottleneck. A relatively small number (100,000 to a few million) of individual protein particles need to be imaged for the high-resolution analysis of proteins by the single particle EM approach, making miniaturized sample handling techniques and microfluidic principles feasible.
A miniaturized, paper-blotting-free EM grid preparation method for sample pre-conditioning, EM grid priming and post processing that only consumes nanoliter-volumes of sample is presented. The method uses a dispensing system with sub-nanoliter precision to control liquid uptake and EM grid priming, a platform to control the grid temperature thereby determining the relative humidity above the EM grid, and a pick-and-plunge-mechanism for sample vitrification. For cryo-EM, an EM grid is placed on the temperature-controlled stage and the sample is aspirated into a capillary. The capillary tip is positioned in proximity to the grid surface, the grid is loaded with the sample and excess is re-aspirated into the microcapillary. Subsequently, the sample film is stabilized and slightly thinned by controlled water evaporation regulated by the offset of the platform temperature relative to the dew-point. At a given point the pick-and-plunge mechanism is triggered, rapidly transferring the primed EM grid into liquid ethane for sample vitrification. Alternatively, sample-conditioning methods are available to prepare nanoliter-sized sample volumes for negative stain (NS) EM.
The methodologies greatly reduce sample consumption and avoid approaches potentially harmful to proteins, such as the filter paper blotting used in conventional methods. Furthermore, the minuscule amount of sample required allows novel experimental strategies, such as fast sample conditioning, combination with single-cell lysis for &#34;visual proteomics,&#34; or &#34;lossless&#34; total sample preparation for quantitative analysis of complex samples.

An instrument and methods for the preparation of nanoliter-sized sample volumes for transmission electron microscopy is presented. No paper-blotting steps are required, thus avoiding the detrimental consequences this can have for proteins, significantly reducing sample loss and enabling the analysis of single cell lysate for visual proteomics.

Due to recent technological progress, cryo-electron microscopy (cryo-EM) is rapidly becoming a standard method for the structural analysis of protein ...

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

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Chapters in this video

Visualizing Proteins and Macromolecular Complexes by Negative Stain EM: from Grid Preparation to Image Acquisition

Analyzing Dynamic Protein Complexes Assembled On and Released From Biolayer Interferometry Biosensor Using Mass Spectrometry and Electron Microscopy

Mitochondria and Endoplasmic Reticulum Imaging by Correlative Light and Volume Electron Microscopy

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

Freeze-Fracture Electron Microscopy for Extracellular Vesicle Analysis

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

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

Visualization of ATP Synthase Dimers in Mitochondria by Electron Cryo-tomography

Determining Membrane Protein Topology Using Fluorescence Protease Protection (FPP)

Miniaturized Sample Preparation for Transmission Electron Microscopy