Name: optimized negative staining: a high-throughput protocol for examining small and asymmetric protein structure by electron microscopy
Uploaded: 2014-08-15T15:00:00.000+00:00
Duration: 9 min 37 s
Description: Watch this Scientific Journal Video about Optimized Negative Staining: a High-throughput Protocol for Examining Small and Asymmetric Protein Structure by Electron Microscopy at JoVE.com

We thank Dr. Mickey Yang for editing and comments, and Drs. Lei Zhang and Bo Peng for assistance. This work was supported by the Office of Science, Office of Basic Energy Sciences of the United States Department of Energy (contract no. DE-AC02-05CH11231), the National Heart, Lung, And Blood Institute of the National Institutes of Health (no. R01HL115153), and the National Institute of General Medical Sciences of the National Institutes of Health (no. R01GM104427).

1. Preparation of Fresh Negative Staining Solution at 1% (w/v)
<ol>
	<li>Put 1 mg Uranyl Formate (UF) powder into glass bottle containing 100 ml deionized water in a dark room or box.</li>
	<li>Stir the solution O/N at RT in a dark room/box. Wrap the solution bottle with aluminum foil to prevent light from hitting the solution.</li>
	<li>Filter the solution gently through 0.2 &#956;m (pore size) filter mounted on a 5 ml syringe. The filtered solution was collected into several aluminum foil covered Falcon tubes. The filter syringe must also be wrapped with aluminum foil to prevent light exposure.</li>
	<li>Filter the solution again through 0.02 &#956;m (pore size) filter mounted on 1 ml syringe. The filtered solution was collected and aliquot into 2 ml vials. The syringes and vials being covered with aluminum foil prior to use.</li>
	<li>Freeze the aliquot vials using long handled forceps submerging the vials into a liquid nitrogen filled container immediately after the solution was filtered.</li>
	<li>Transfer the frozen vials into a -80 &#176;C freezer for storage and future usage.</li>
</ol>
2. Preparation of Negative Staining Workstation and Incubation Workstation
<ol>
	<li>Thaw a vial of 1% UF solution in a water bath set at RT. Make sure that the aluminum foil remains wrapped around the vial to prevent exposure to light.</li>
	<li>Filter the solution after it&#8217;s completely thawed by using an aluminum foil wrapped 1 ml syringe mounted with a 0.02 &#956;m filter. Collect the filtered solution into a new aluminum foil wrapped vial, and placed it on the ice inside a cap covered ice box waiting for usage.</li>
	<li>Make solution plates by finger pushing a ~8 inch long Parafilm sheet onto the surface of an empty protein crystallization plate. It will generate rows of circular plates with a diameter of ~5 mm. Make 6 plates in each row place the Parafilm plates on a surface of flattened ice inside an ice container lid, as a staining workstation.</li>
	<li>Pipette ~35 &#956;l deionized water onto each of left three plates in each row, and pipette ~35 &#956;l filtered UF solution on to right three plates in each row. Cover the staining workstation with a lid to prevent exposure to light before staining proteins.</li>
	<li>Prepare an EM grid incubation station by filling an empty box halfway with ice, and adhered next to a hanger that can be mounted on a supporting base. Place the sample tweezers in the hanger, in which the tweezers&#8217; tips are nearby the ice surface. Half cover the tweezers&#8217; tips by the box lip height. An ideal ice box to make an incubation station is using an empty pipet tips container.</li>
</ol>
3. OpNS Operation
<ol>
	<li>Glow-discharge the thin carbon film coated 300 mesh copper EM grids for 10 sec.</li>
	<li>Pick up a grid with tweezers and hook the tweezers into the hanger by keeping the grid at a 45&#176; tilt and 1 inch above the ice surface inside the incubation station.</li>
	<li>Dilute the protein sample with Dulbecco&#8217;s Phosphate Buffered Saline (DBPS) at final protein concentration of ~0.01 to ~0.005 mg/ml. Deposit ~4 &#956;l diluted sample immediately after the dilution on the carbon side of EM grid. (DPBS can be changed to another buffer if the protein is sensitive to DPBS)</li>
	<li>Incubate the sample on the EM grids for ~1 min inside the incubation station.</li>
	<li>Remove the excess solution through quickly touching the grid edge with filter paper.</li>
	<li>Touch the grid quickly to the first drop (~35 &#956;l) of pure water surface atop the Parafilm sheet right after the excess solution was removed on the EM grid. And then remove the excess water promptly with filter paper.</li>
	<li>Repeat step 3.6 for another two times by washing the EM grid on the remaining two water droplets on the Parafilm (Step 3.6 and 3.7 can be skipped if the sample is very sensitive to water).</li>
	<li>Float the EM grid immediately on the surface of the first drop of UF solution right after the excess water on the EM grid was removed. And then incubate for 10 sec. Make sure to finish the water washing procedures within 3 sec before floating the grid on the surface of UF solution. The shorter the washing time, the better the quality will be.</li>
	<li>Clean the tweezers by penetrating the tips into a filter paper for ~2 - 3 times.</li>
	<li>Remove the excess solution on the grid through contacting the grid edge with the filter paper, and then float the grid on the second drop of UF.</li>
	<li>Continue above step 3.10 on the second droplet.</li>
	<li>Float the grid on the third droplet of UF solution and cover the staining station for ~1 min.</li>
	<li>Optional step: wash the grid quickly on a top of water as described in 3.6. This additional step will benefit samples containing over loaded free gold particles or stain background.</li>
	<li>Remove the excess solution by touching the filter paper to the entire grid backside (opposite the carbon side). Dry the grid under a gentle stream of nitrogen gas at RT right after observing the solution absorbing to the filter paper.</li>
	<li>Place the grid onto a sheet of filter paper inside a petri dish, and cover the dish partially with a cap to dry for at least 30 min under RT. For some samples, place the grid in a 40 &#176;C incubator baking for an hr.</li>
	<li>Store the grid into an EM grid box for future EM examining.</li>
</ol>
4. EM Examining
<ol>
	<li>Align the TEM properly before EM examination of the small proteins on the grid.
	<ol>
		<li>Check the resolution of the highest visible Thon-ring that is the power spectrum of an amorphous carbon film to determine whether the TEM has been aligned properly. Since the TEM is shared by many different users, the TEM was often not properly aligned.</li>
		<li>Carefully check the power spectrum on the carbon film area of the specimen to adjust the alignment condition of the machine. The highest visible Thon-rings are better than the targeted resolution. 
		NOTE: As an example of a proper alignment of a LaB6 filament TEM operated under 120 kV high tension, more than 20 Thon-rings can be visualized, in which, the corresponding resolution can be better than 5.0 &#197;. This Thon-ring was achieved by Fourier transfer of an amorphous carbon film imaged under a defocus of ~1.6 &#181;m and a dose of 20.4 e-/&#197;2 (Figure 5). 
		NOTE: Observation of the high resolution Thon-ring is a necessary condition, but not a sufficient condition for proper alignment. To actually achieve the high resolution images, many other parameters are also important, such as sample quality, radiation damage, charging and drifting as well as the illumination dose rate.</li>
	</ol>
	</li>
	<li>Bring the defocus back to near-Scherzer focus for imaging the small proteins on an ideally stained area.</li>
	<li>Search for &#8220;cloudy&#8221; area to image. An ideally stained area is usually located near the edge of a thicker stain area, which looks like a &#8220;cloudy&#8221; area on the grid since the thickness of the stain is normally not evenly distributed on the carbon-coated grid (Figure 6). Typically a low magnification (&#60;100x) was to help to find these cloudy areas first before zooming in for imaging.</li>
</ol>

We have nothing to disclose.

Compared to conventional NS techniques, OpNS can prevent rouleaux artifacts (Figure 1 and 9), providing reliable and reasonable high&#8208;resolution (~1nm) structural details of small proteins (Figure 2). Compared to cryo-EM, OpNS provides a high throughput method and can examine a wide variety of proteins and protein-protein interactions6. However, OpNS still has its disadvantages. Compared to conventional NS, OpNS entails: i) more complicated steps in specimen preparation; ii) using a radioactive substance, UF; iii) keeping the stain fresh due to the shorter potency of the UF stain because of its light sensitivity; iv) to prevent precipitation the UF solution must be stored in -80 &#176;C and requires thawing before use; v) and finally all UF must be handled as hazardous waste. Compared to cryo-EM, OpNS provides relatively lower resolution images and no guarantee for any not yet discovered artifact in future.
NS operation procedural effects on lipoprotein rouleaux formation
NS protocols for preparing proteins for examination16,29-36,55 can be classified into three major groups of methods50: mixing55, drop-by-drop16, and washing procedures29. i) The mixing protocol requires the preliminary mixing of the stain and protein sample at a specific ratio, and then applying the mixed solution onto carbon film coated on a grid, finally removing excess solution with filter paper before air drying for EM examination55. ii) The drop&#8208;by&#8208;drop protocol involves applying (~4 &#956;l) sample drop directly to an EM grid coated in carbon letting sit for ~1 min, subsequently removing excess sample solution before application of a drop (~4 &#956;l) of stain solution on the same side of grid. After ~1 min of incubation, remove the excess stain through contact with clean filter paper, finally submitting for air drying16,56-58. iii) The washing protocol (Figure 7) requiring sample solution application to an EM grid coated in carbon for 1 min, then washing with deionized water three times right after removing the excess solution each time by filter paper3,29,30. OpNS protocol was modified from this washing procedure.
NS stain types and effects on lipoprotein rouleaux formation
In NS, stronger electron scattering due to heavy metal stains provide contrast from the proteins17-20,59. NS samples can additionally suffer greater radiation doses, and enhance protein features by a sharper negative contrast8,9,60. Stains of heavy metals can be classified as: anionic, including phosphotungstic acid (PTA)16, methylamine tungstate14 and silicon tungstate61; and cationic, such as Uranyl acetate (UA)62, UF3,29,30, and Uranyl nitrate (UN)63. One of the anionic NS stains that is most commonly used is PTA33,64-67. PTA is a heteropoly acid, which is typically used around a pH 7.0 - 7.5. PTA can also be used for positive staining3,16,68. The negative charges of PTA may mediate electrostatic interactions with unsaturated lipids positively charged head groups, like POPC, on both lipoprotein and liposome surfaces. PTA may cause rouleaux formation through this interaction 29,30 even under regular buffer salinity29,30 (Figure 1). Uranyl stains, such as UA, UF and UN are alternative choices for cationic heavy metals stains and UA in particular is frequently used for NS of a variety of biological specimens62,69,70. Experiments found UF causes no rouleaux of lipoproteins that contain unsaturated fat acid lipids, such as POPC. However, UF may still induce rouleaux formation on lipoprotein that contain saturated fatty acid lipids, such as the dimyristoyl phosphatidylcholine (DMPC) and 1-hexadecanoyl-2-octadecanoyl-sn-glycero-3-phosphocholine (PSPC). The detailed mechanism of this interaction is unknown. UA/UF/UN can all work at lower pH values ranging from 3.5 to 4.6. Such lower pH values may not suitable for certain biological macromolecules that are sensitive to pH14,71. Interestingly, UA/UF can fix protein structure within a few milliseconds through an unknown mechanism72. Unlike PTA, UA/UF is more similar to a salt than an acid.
UF reportedly can provide better results than UA73, in which, UF and UN stains have smaller grain sizes than UA (UF: 0.3 nm, UN: ~0.5 nm)3,74. An interesting example, secondary structure-like details of a small protein (53 kDa CETP) can be directly visualized from the raw micrographs when UF was used as negative-stain reagent by following the earlier reported cryo-positive-staining (cryo-PS) method (Figure 8)6. In cryo-PS, the stained sample was flash frozen by following cryo-EM sample preparation procedure instead of drying procedure in OpNS protocol, which may cause the secondary structural collapse. The cryo-PS is a method for high-contrast and high-resolution imaging of small proteins75. Since the cryo-PS images have reversed contrast compared to those from the reported cryo-NS protocol 22, but have consistent image contrast to those from conventional cryo-EM images, thus it was named the cryo-PS method. The cryo-PS images show that the staining reagent, Uranyl formate, penetrates the molecular surface, challenging the conventional view that staining can visualize only the outer surface structure. The mechanism of how Uranyl formate penetrates the molecular surface is unknown. One possibility is that the Uranyl cation binds available protein carboxyl groups, and thus the surrounding vitreous ice is of lower density than the staining of the protein and Uranyl groups, thereby acting as a positive stain. The cryo-PS method is similar to multiple isomorphous replacement (MIR) method, which was a common approach to solve the phase problem in X-ray crystallography 76-78. In MIR, crystal samples are soaked with a heavy atom solution, including Uranium, to get an isomorphic form to its native form. The addition of the heavy atom sometimes does not affect the crystal formation or unit cell dimensions but provides additional information of crystal structure determination76-78. By benefiting from the small grain of UF, cryo-PS could provide a high contrast and high-resolution image of a single protein, which is important for protein structure studies, especially when considering that nearly all proteins are naturally dynamic and heterogeneous in solution. For validation of this cryo-PS method, we open for any blind test from the EM field.
Salt concentration effects on the rouleaux formation in lipoproteins
Using traditional PTA negative staining reagent, observed rouleaux formation is more likely related to lipid interactions instead of protein interaction. Imaging the samples of POPC liposome vesicles prepared under considerable salinity (0.5 M NaCl), moderate salinity (0.25 M NaCl), relatively weak salinity (0.1M NaCl), and pure water (Figure 9), the electron micrographs show the rouleaux formation was correlated to salt concentration (Figure 9A - D). Using UF as negative staining reagent, no rouleaux was observed in POPC liposome vesicle sample30 (Figure 2L), suggesting the washing procedure to quickly reduce the salt concentration right before staining is an necessary, but not independently sufficient step for preventing rouleaux in POPC related samples.
TEM imaging small protein requires a near-Scherzer focus condition.
Successful TEM imaging of small proteins at higher resolution requires two critical conditions, a proper beam alignment and a near-Scherzer focus acquisition condition. i) A proper beam alignment can be judged through the number of visible Thon rings (power spectrum) on the Fourier transfer of a carbon film image acquired under a relatively high defocus. The better the alignment condition of the beam, the more Thon rings can be visualized and measurable.. ii) Acquiring image under a near-Scherzer focus is another critical condition. A traditional standard strategy for imaging biological specimens typically utilizes high defocus (1 - 2 &#956;m and even more), which can enhance the biological sample image contrast 79. This strategy is significant different from the typical strategy for imaging atomic resolution structure of hard materials, which often utilizes a near Scherzer focus (~50 nm defocus). Although, a higher defocus could strengthen biological sample contrast, high-resolution signals would be partially eliminated. As a result, the complicity of high-resolution signal would be lower at a higher defocus imaging condition. The reason is that, the TEM image is the convolution of sample structure and the instrument CTF. The CTF is an oscillating curve against the frequency, in which the curve frequently oscillated crossing zero amplitude at high frequency. Every time when CTF across zero, the sample structural signal at this specific frequency will be eliminated (zero times any number will be zero). The eliminated signal at this frequency can never be recovered by any CTF correction algorithm (a zero divided by a zero can be any random number instead of the original structural signal). Under higher defocus imaging, the CTF will oscillate much more aggressively, thus the CTF crosses zero amplitude more frequently, as a result, the structural signals at a greater number of specific frequencies will be eliminated. The more signals that are eliminated, the less complicity the image will have. Notably, the complicity of image cannot be recovered or corrected by any CTF correction. However, a number of incompleted images acquired under different defocuses can be used to fill their gaps to each other to obtained a completed structural signal of the sample structure via an averaging method, in which even an atomic resolution can be achieved 9-12 .
The averaging method is a powerful approach to achieve the structure of some highly symmetric proteins and structurally related rigid proteins. However, it still remains challenging in the structural study of small and asymmetric proteins, especially for structural dynamics and flexible proteins, such as antibodies and HDLs. Averaging is based on the assumption that protein particles are identical in structure and conformation but different in orientations, however many proteins are known to undergo thermodynamic fluctuation in solution. By applying the averaging method without previously knowing the protein thermodynamic fluctuation fluctuations, the averaged structure can often cause missing domains 10,80 or local variation in resolution 81 in cryo-EM reconstructions .
The success in achieving the 3D reconstruction of small protein, such as 53 kDa CETP, and 3D structure of a single protein, such as 160 kDa IgG1 antibody, benefits from using the near-Scherzer focus imaging condition under a proper alignment condition. Although the image contrast seems low in the near-Scherzer focus images, the contrast can be easily enhanced by simply applying a low pass filtering to reduce the high-frequency noise in the background. We believe, the near-Scherzer focus images carry the maximum structural signals, and can increase the accuracy of particle alignment and enhance efficiency in single-particle 3D reconstruction and individual particle electron tomography reconstruction.

Understanding protein function requires knowledge of protein structure. Structural determination is challenging for proteins whose molecular masses are within 40 - 200 kDa. X-ray crystallography is limited by protein crystallization; nuclear magnetic resonance (NMR) spectroscopy is limited to molecular masses less than 40 KDa, while cryo-electron microscopy (cryo-EM) has difficulty in both image acquisition and three-dimensional (3D) reconstructions of small proteins, which molecular masses are less than 200 kDa. Notably, more than 50% proteins have a molecular mass within the range of 40 - 200 kDa1,2, as current methods are challenging in studying proteins of this size, a new method is needed.
Although most transmission electron microscopes (TEMs) are capable of atomic resolution, i.e., better than 3 &#197; resolution, achieving even a near nanometer resolution structure from a biological samples is rather challenging7. Radiation damage, low contrast, structural deviations as well as artifacts such as dehydration all hinder high-resolution TEM imaging3,8.
Among various TEM approaches, cryo-EM is an advanced and cutting edge method to achieve atomic resolution structures of highly symmetric large macromolecules under near physiological conditions9-12. The cryo-EM sample is prepared flash freezing the sample solution, embedding the macromolecules in vitreous ice, which is subsequently imaged at cryogenic temperatures such as liquid nitrogen or helium temperatures13. Cryo-EM is advantageous in that samples present no artifacts and are nearly native in structure8-12. Cryo-EM does have its disadvantages: i) additional devices are required to be installed or purchased to upgrade a standard TEM instrument for a cryo-EM capability. Devices include: anti-contaminator, cryo-holder, low-dose mode software and low-dose sensitive CCD camera, although the prices of these devices are much lower than the price of the TEM instrument itself; ii) cryo-EM operation needs longer time than NS operation. Examining a cryo-EM specimen often requires longer time to prepare specimens and operate the TEM instrument than that of NS because cryo-EM requires addressing additional difficulties, including: liquid nitrogen temperature operation, sample charging, imaging drift, temperature gradients, low-dose model operation, sample radiation sensitivities and dosage limitations. These extra steps will slow down the speed of acquiring useful data compared to NS data acquisition, although a few cryo-EM images can certainly be obtained in 1 hr or less by cryo-EM experts with the instrument prepared with a balanced temperature gradient; iii) users require additional training, such as handling liquid nitrogen, freezing cryo-EM grids, low-dose operation, dose measurement, handling the charging, drifting and knowledge in imaging processing; iv) lack of repeatable imaging for the same cryo-specimen during different TEM sessions. Cryo-EM specimens are easily damaged by ice contamination during specimen loading and unloading to/from the TEM instrument. This damage is especially a concern when the samples are difficult to be isolated/purified14; v) small proteins (&#60;200 kDa molecular mass) are challenging to be imaged because of low contrast; vi) the low contrast and high noise of cryo-EM images reduces the cross-correlation value between images, therefore, decreasing overall accuracy in the determination of protein orientations, conformations and classifications, especially for proteins that are structurally flexible and naturally fluctuate in solution4,5.
Negative staining (NS) is a relatively &#8220;ancient&#8221; and historical method which any laboratory, with any type EM, can utilize to examine protein structure. Brenner and Horne first developed the concept of negative staining a half century ago for examining viruses15. NS is accomplished through coating the specimen with charged heavy metal salts. This concept originally coming from light microscopy and the practice of embedding bacteria into a stain solution providing darkness around the specimens, allowing higher image contrast when viewing the negative image16. Since the heavy metal ions have a greater ability to disperse electrons compared to less dense atoms in the proteins17-20, and coating heavy metal stain permits a higher dosage limitation with improved contrast. NS specimen can provide high contrast images8 for easier particle orientation determination and 3D reconstruction than images from cryo-EM.
Traditional NS, unfortunately, can produce artifacts induced by stain-protein interactions, such as general aggregation, molecular dissociation, flattening and stacking8,21,22. For lipid related proteins, such as lipoproteins16,23-30, a common artifact results in particles that are stacked and packed together into a rouleaux (Figure 1)31-36. Many lipoprotein studies, such as nondenaturing polyacrylamide gradient gel electrophoresis, cryo-EM studies13,29,37-40, mass spectrometry39,41, and small angle X-ray diffraction data42 all show lipoprotein particles are isolated particles instead of naturally stacked together forming a rouleaux 21,29,30,35,42-45. The observation of rouleaux formation by conventional NS is possibly caused by dynamic interactions between lipoproteins composed of apolipoproteins (apo) and phospholipids that are structurally flexible in solution 13,29,30,46-49 and sensitivity to the standard NS protocol. To identify this artifact, apolipoprotein E4 (apoE4) palmitoyl-oleoylphosphatidylcholine (POPC) high-density lipoprotein (HDL) sample were used as a test sample and cryo-EM images for an artifact free standard 29, screening the NS specimens prepared under a series of conditions. By comparing the particle sizes and shapes obtained from NS and cryo-EM, the specific type of staining reagent and salt concentration were found to be two key parameters causing the well known rouleaux phenomena. Thus, an optimized negative staining (OpNS) protocol was reported.
By OpNS, the well known rouleaux phenomenon of apoE4 HDL was eliminated by OpNS (Figure 2A). Statistical analysis demonstrated OpNS yields very similar images (fewer than 5% deviation) in size and shape in comparison to those from cryo-EM, however the contrast was eliminated. The validations of OpNS were performed by examining the elimination of the rouleaux artifact of nearly all classes or subclasses of lipoprotein samples 6,29,30,50,51, including apoA-I 7.8 nm (Figure 2B), 8.4 nm (Figure 2C), 9.6 nm discoidal reconstituted HDL (rHDL) (Figure 2D), 9.3 nm spherical rHDL (Figure 2E), human plasma HDL (Figure 2F), lipid free apoA-I (Figure 2G), plasma HDL (Figure 2H), low density lipoprotein (LDL) (Figure 2I), intermediate&#8208;density lipoprotein (IDL) (Figure 2J), very low density lipoprotein (VLDL) (Figure 2K), and POPC liposome (Figure 2L) 30. Additional validations were performed by imaging small and asymmetric proteins, including the 53 kDa cholesteryl ester transfer protein (CETP) (Figure 3A - C)6,29, and highly flexible 160 kDa IgG antibody (Figure 4A and B)4,5,29,52, and two structurally well known proteins, GroEL and proteasome (Figure 2M and N). For requiring any additional validation from colleagues, we are open to any blind tests on this OpNS method.
OpNS as a high-throughput protocol has also been used to study protein mechanism via examining hundreds of samples of small proteins, such as CETP that was binding to various proteins under a series conditions (including CETP interacting to 4 classes of lipoproteins, recombined HDL, plasma HDL, LDL and VLDL, with/without 2 antibodies, H300 and N13, under 9 incubation times, including 3 min, 20 min, 1 hr, 2 hr, 4 hr, 8 hr, 24 hr, 48 hr and 72 hr, under 4 molar ratios, i.e.,&#160;1:0.5, 1:1, 1:2, 1:4, and 3 dilutions, i.e., 0.1 mg/ml, 0.01 mg/ml and 0.001 mg/ml, plus additional control samples, including CETP alone, LDL alone, and VLDL alone, with triple tests of above experiments by different persons)6,29. OpNS images of CETP provided high contrast images with reasonably fine structural details; allowing us to successfully reconstruct a 3D density map of the 53 kDa small protein CETP (Figure 3D - F) by single particle reconstruction. Moreover, the high contrast OpNS images provide us a sufficient signal from an individual protein (Figure 4A - C), which allowed us to achieve the intermediate resolution (~1.5 nm) of a single (one object, no average) IgG antibody 3D structure via the individual-particle electron tomography (IPET) method (Figure 4E - J)5. The detailed description of IPET reconstruction strategy, methodology, step-by-step processes and structural variation analysis were previously reported4. A movie about IPET antibody reconstruction procedures, including the raw images and intermediate results, 3D density map and structural docking was also available to public by uploaded to YouTube5. Comparison of the 3D reconstructions from different individual antibody particles could reveal the protein dynamics and conformational changes during chemical reactions4,5.
Considering that over 50% of proteins have molecular mass ranging from 40 - 200 kDa1,2, the success in imaging these small proteins evidenced that OpNS method is a useful tool to push the conventional EM boundary toward small and asymmetric structural determinations and mechanism discoveries. Thus, the detailed protocol is provided as below.

<table><tbody><tr><td>Stain: Uranyl Formate</td><td>The SPI Supplies Division of Structure Probe, Inc.</td><td>02545-AA</td><td>http://www.2spi.com/catalog/chem/Uranyl_Formate.shtml</td></tr><tr><td>Syringe: 1 ml NORM-JECT</td><td>VWR</td><td>89174-491</td><td>https://us.vwr.com/store/catalog/product.jsp?catalog_number=89174-491</td></tr><tr><td>Syringe filter: 0.2 &amp;mu;m&amp;nbsp;</td><td>VWR</td><td>28145-487</td><td>https://us.vwr.com/store/catalog/product.jsp?catalog_number=28145-487</td></tr><tr><td>Syringe filter: 0.02 &amp;mu;m filter (Anotop 10)</td><td>Whatman</td><td>6809-1002</td><td>http://www.whatman.com/AnotopSyringeFilters.aspx</td></tr><tr><td>Dulbecco&amp;rsquo;s Phosphate Buffered Saline</td><td>SIGMA-Aldrich</td><td>D8537</td><td>http://www.sigmaaldrich.com/catalog/product/sigma/d8537?lang=en&amp;amp;region=US</td></tr><tr><td>Parafilm: 4 in. x 125 ft.</td><td>VWR</td><td>470152-246</td><td>https://us.vwr.com/store/catalog/product.jsp?catalog_number=WLS65710-A</td></tr><tr><td>Carbon film coated TEM grid: 300 mesh</td><td>Pacific Grid-Tech</td><td>Cu-300CN</td><td>http://www.grid-tech.com/catalog.htm#1-thincarbon</td></tr><tr><td>Carbon film coated TEM grid: 200 mesh</td><td>EMS (Electro Microscopy Sciences)</td><td>CF200-Cu</td><td>http://www.emsdiasum.com/microscopy/products/grids/support.aspx</td></tr><tr><td>Tweezer: Dumont Tweezer L5</td><td>EMS (Electro Microscopy Sciences)</td><td>72882-D</td><td>http://www.emsdiasum.com/microscopy/products/tweezers/dumont_clamping.aspx#02-L5</td></tr><tr><td>Milli-Q Advantage A10 Ultrapure Water Purification System</td><td>EMD Millipore</td><td>Milli-Q Advantage A10</td><td>http://www.millipore.com/catalogue/module.do?id=C10117</td></tr><tr><td>Filter Paper: Grade 1 circles</td><td>Whatman</td><td>1001-090</td><td>http://www.whatman.com/QualitativeFilterPapersStandardGrades.aspx</td></tr><tr><td>Aluminum Foil</td><td></td><td></td><td>Any type</td></tr><tr><td>Filter Tips: Low-Retention Aerosol Filter Tips for 10&amp;micro;L Pipettors</td><td>VWR</td><td>89174-520</td><td>https://us.vwr.com/store/catalog/product.jsp?catalog_number=89174-520</td></tr><tr><td colspan="4">[header]</td></tr><tr><td>Glow Discharge Unit</td><td>Ted Pella</td><td>91000</td><td>http://www.tedpella.com/easiglow_html/Glow-Discharge-Cleaning-System.htm</td></tr><tr><td>Filter Tips: Low-Retention Aerosol Filter Tips for 40&amp;micro;L Pipettors</td><td>VWR</td><td>89174-524</td><td>https://us.vwr.com/store/catalog/product.jsp?catalog_number=89174-524</td></tr><tr><td>Filter Tips: VWR Signature 200 &amp;micro;L Pipet Tips</td><td>VWR</td><td>37001-528</td><td>https://us.vwr.com/store/catalog/product.jsp?catalog_number=37001-528</td></tr><tr><td>Pipet</td><td>Pipetman</td><td>F161201 and F167300</td><td>http://www.pipetman.com/Pipettes/Single-Channel/PIPETMAN-Classic.aspx</td></tr></tbody></table>

optimized negative staining: a high-throughput protocol for examining small and asymmetric protein structure by electron microscopy

Structural determination of proteins is rather challenging for proteins with molecular masses between 40 - 200 kDa. Considering that more than half of natural proteins have a molecular mass between 40 - 200 kDa1,2, a robust and high-throughput method with a nanometer resolution capability is needed. Negative staining (NS) electron microscopy (EM) is an easy, rapid, and qualitative approach which has frequently been used in research laboratories to examine protein structure and protein-protein interactions. Unfortunately, conventional NS protocols often generate structural artifacts on proteins, especially with lipoproteins that usually form presenting rouleaux artifacts. By using images of lipoproteins from cryo-electron microscopy (cryo-EM) as a standard, the key parameters in NS specimen preparation conditions were recently screened and reported as the optimized NS protocol (OpNS), a modified conventional NS protocol 3 . Artifacts like rouleaux can be greatly limited by OpNS, additionally providing high contrast along with reasonably high&#8208;resolution (near 1 nm) images of small and asymmetric proteins. These high-resolution and high contrast images are even favorable for an individual protein (a single object, no average) 3D reconstruction, such as a 160 kDa antibody, through the method of electron tomography4,5. Moreover, OpNS can be a high&#8208;throughput tool to examine hundreds of samples of small proteins. For example, the previously published mechanism of 53 kDa cholesteryl ester transfer protein (CETP) involved the screening and imaging of hundreds of samples 6. Considering cryo-EM rarely successfully images proteins less than 200 kDa has yet to publish any study involving screening over one hundred sample conditions, it is fair to call OpNS a high-throughput method for studying small proteins. Hopefully the OpNS protocol presented here can be a useful tool to push the boundaries of EM and accelerate EM studies into small protein structure, dynamics and mechanisms.

The implementations of OpNS include the structural and morphological examinations of various lipoprotein species such as: nascent recombinant HDL (rHDL), spherical recombinant HDL, plasma HDL, LDL, IDL and VLDL (Figure 2)30; as well as small proteins such as 53 kDa CETP (among the smallest proteins successfully imaged through EM) (Figure 3)6 and 160kDa IgG antibodies (one of the most dynamic and heterogeneous proteins) (Figure 4)4,5,29,52, even 28 kDa lipid free apolipoprotein A-1 (Figure 2L), and GroEL and Proteasomes (Figure 2M and N).
Other examples of OpNS as a high-throughput method are examining protein‐protein interactions for uncovering the protein mechanisms, such as 53 kDa CETP. As described in the introduction, how CETP interacts with different subclasses of lipoproteins were investigated via examining more than 400 EM specimens6 As far as we know, there was no such a case of cryo-EM study that involves screening nearly a hundred different conditions .
OpNS can expand EM boundaries to study small and asymmetric protein 3D structure and even expand to achieve 3D structure form a single and individual protein (without averaging). For example, IgG antibody is 160 kDa molecule with a highly flexible structure, in which the 3D reconstruction at intermediate resolution is challenging to be achieved. the OpNS method was used to image an individual IgG antibody from a series of tilting angles (Figure 4E and H). The reasonably high-resolution and high contrast protein images from OpNS allowed for the successful reconstruction of a single protein by individual particle electron tomography (IPET) (Figure 4E - J)4,5. In IPET 3D reconstruction4, the 2D tilt series of the targeted antibody were gradually aligned to their calculated global center via subsequently iterative reconstruction to achieve a 3D density at a resolution of ~14.1 Å (Figure 4F and G)4. By the same approach, a single antibody‐peptide complex was reconstructed, and the peptide induced the internal domain conformational change of a single antibody particle was discovered (Figure 4I and J, resolution ~16.6 Å)4,5. Comparing the 3D reconstructions from different individual particles, the structural analysis may allow us to reveal protein thermodynamic fluctuations, and even “snap-shot” the intermediate stages of a chemical reaction4,5,29,53,54.
In summary, the proteins with molecular mass between 40 kDa - 200 kDa are challenging for standard EM structural examinations and reconstructions. Considering that over 50% of proteins have molecular mass ranging from 40 - 200 kDa1,2, we report an OpNS method as a robust and high-throughput tool to push the conventional EM boundary toward small and asymmetric structural determinations and even mechanistic studies.
<img alt="High-resolution microscopy images of ApoE4 rHDL, ApoA-I rHDL, ApoB LDL, and POPC Liposome structures." fo:content-width="5in" src="/files/ftp_upload/51087/51087fig1highres.jpg" width="500" data-altupdatedat="1747911167000" data-alt="Figure 1"> 
Figure 1. Rouleau artifact of lipoproteins by conventional negative staining (NS) EM. Electron micrographs (above panels) and selected particles (below panels) show various lipoproteins with rouleaux after NS with phosphotungstic acid (PTA) under standard mix procedure. (A) Reconstituted HDL (rHDL) with ApoE4, (B) apoA-I-containing 9.6 nm discoidal rHDL, (C) apoB containing plasma LDL, (D) non-apolipoprotein-containing POPC liposomes. Bars: 50 nm. Window size: A and C, 20 nm; B, 30 nm. The Journal of Lipid Research initially published this work29,30. <a href="https://www.jove.com/files/ftp_upload/51087/51087fig1highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Cryo-EM micrographs, ApoE4 rHDL, LDL, VLDL particles; structural analysis, lipid-protein complexes." fo:content-width="5in" src="/files/ftp_upload/51087/51087fig2highres.jpg" width="500" data-altupdatedat="1747911167000" data-alt="Figure 2"> 
Figure 2. Protein structure by optimized negative staining (OpNS) EM. Electron micrographs show various lipoproteins and liposomes without rouleaux artifact by OpNS. (A) apoE4-containing rHDL; (B) 7.8 nm rHDL; (C) 8.4 nm rHDL; (D) 9.6 nm rHDL; (E) 9.3 nm spherical rHDL; (F) Human plasma α-HDL; (G) Plasma HDL; (H) Human plasma LDL; (I) Human plasma IDL; (J) Human plasma VLDL; (K) POPC liposome; (L) lipid free apoA-I. Additional verification was done using (M) GroEL and (N) Proteasome samples. Micrographs (above panels) and selected particles (below panels). Bars: 50 nm. Window size: A - D, 20 nm; E and F, 25 nm; G, 20 nm; H, 25 nm; I, 50 nm; J and K, 100 nm; L, 80 nm., Except lipid free apoA-I, GroEL and Proteasome, This research was originally published in the Journal of Lipid Research29,30. <a href="https://www.jove.com/files/ftp_upload/51087/51087fig2highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Cryo-EM protein structure analysis, particle images and 3D reconstructions, experimental result." fo:content-width="5in" src="/files/ftp_upload/51087/51087fig3highres.jpg" width="500" data-altupdatedat="1747911167000" data-alt="Figure 3"> 
Figure 3. OpNS electron micrographs of CETP. (A) Overview micrograph with CETP (dashed circles) banana shaped particles. (B) Windowed select raw particles. (C) Three single raw CETP particle images clearly showing a bigger, heavier and larger globular tip including the other smaller, lighter and narrower tip (left panel), top view showing similar terminal regions with less overall curvature (middle panel), and end view down the long axis of CETP (right panel). (D) Superimposing the crystal structure (PDB: 2OBD) onto these raw CETP particle images, matches well in both structural shape and domain size showing the orientation can be nearly directly defined even without assistance by computer (corresponding panels to (C)). (E) Reference free 2D class averages (an ab initio process to compute the similarities (cross-correlation calculation) of these randomly oriented particles, the particles having a high similarity to each other were grouped, aligned and then averaged to reduce the background noise and enhanced the particle image contrast. The reference free 2D class averages are calculated by refine2D.py software in the EMAN package, in which, no any human made initial model was involved in this class average. The reference class averages can be used as an independent method to validate the 3D reconstruction from single-particle reconstruction). (F) Selected reference free 2D class averages show a larger distal end compared to the other. (G) Superimposed crystal structure (PDB: 2OBD) comparison to the reference free averages, the overlay images show a near perfect matches between crystal-structure and reference-free class average in both structure shape and domain size (low panel). (F) 3D density map of CETP at a resolution of 13 Å was reconstructed from 8,879 particles imaged by OpNS and single-particle reconstruction method (top panel) and the ridged-body docked into this single-particle reconstruction by the crystal structure (low panel). The detailed information of the single particle reconstruction, including image preprocessing, initial model, refinement procedures, angle distribution and Fourier Shell Correlation (FSC) were published in the method section and supporting information of the original paper (H) Final comparison of single particle reconstruction (above panel) and additional PDB fit comparison (bottom panel) 6. Bars: A, 50 nm; B - D, 10 nm; F, 3 nm. Some of these figures were previously published in the Nature Chemical Biology6. <a href="https://www.jove.com/files/ftp_upload/51087/51087fig3highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Cryo-EM protein structure analysis; projection data, 3D reconstruction, color-coded density maps." fo:content-width="5in" src="/files/ftp_upload/51087/51087fig4highres.jpg" width="500" data-altupdatedat="1747911167000" data-alt="Figure 4"> 
Figure 4. OpNS images and reconstructions of Human IgG1 antibody particles. (A) Overview of human IgG1 antibody OpNS images. Particles shown in white circles clearly display the particles with 3 subdomains. (B) Selected high-resolution raw images of five individual unconjugated antibody particles and (C) their corresponding noise-reduced images by manually reducing noise surrounding raw particle edge (without touch any density inside the particles) to visualize easily. (D) The crystal structure (PDB entry 1IGT) of IgG1 antibody that was oriented to a similarly viewing to the EM images. Comparison of the corresponding images shows many common features between the EM image and the crystal structure, including domain positions and shapes. (E) The step-to-step procedure of an ab initio 3D reconstruction from a single instance IgG1 antibody (no average, one individual object) using individual‐particle electron tomography (IPET) method. (F) The final 3D reconstruction of a single antibody particle at resolution of 14.1 Å showed three donut shaped domains forming in a “Y” shape. (G) Docking the domains crystal structures into each of corresponding domain density maps respectively, and manually repeating the loops among the domains. (H) The step-to-step procedure of 3D reconstruction of a single IgG-peptide complex (no average, one individual object) by IPET. (I) The final 3D reconstruction of an IgG peptide complex was displayed at resolution of 16.6 Å showed three rod shaped domains forming in a “Y” shape. (J) Respectively, docking the crystal structure of each IgG antibody domains (PDB entry: 1IGT) into each rod shape densities showed a poorly matching with domain crystal structures, suggesting an inner domain conformational change after peptide conjugation. The detailed procedure, resolution analysis and statistics were described in the original paper 5 Bars: A, 50 nm; B - D, 10 nm. This work initially published in PLoS ONE4 and Scientific Reports5. <a href="https://www.jove.com/files/ftp_upload/51087/51087fig4highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Electron diffraction pattern with a contrast transfer function (CTF) intensity graph for microscopy analysis." fo:content-width="5in" src="/files/ftp_upload/51087/51087fig5highres.jpg" width="500" data-altupdatedat="1747911167000" data-alt="Figure 5"> 
Figure 5. Power spectrum of amorphous carbon film imaged under 1.6 µm defocus by a Zeiss Libra 120 LaB6 TEM. High-resolution imaging requires sufficient evidence to show that EM has the proper alignment and high-resolution capability. The analysis of the power spectrum of nearby carbon area on the grid is an efficient method to check the beam coherence condition prior to data acquisition. Fourier transfer of the image of an amorphous carbon area show the instrument related contrast transfer function (CTF), so called Thon-rings. A proper alignment and coherence can be reflected by the number of visible Thon-rings and the highest resolution of visible Thon rings under a relatively high defocus condition. As an example of a near proper alignment condition of a LaB6 filament equipped TEM, more than ~20 Thon rings (~5 Å) can be generated from a carbon film are imaged at 1.6 µm defocus using dose 20.4 e-/Å2. The Thon rings achieved from a low-end TEM have a similar to that from high-end TEM, such as a field-emission gun (FEG) enhanced TEM operated under a proper alignment condition. This work initially published in Scientific Reports5. <a href="https://www.jove.com/files/ftp_upload/51087/51087fig5highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Microscopy images showing crystal structure at 80x, 4Kx, 80Kx magnifications; material analysis." fo:content-width="5in" src="/files/ftp_upload/51087/51087fig6highres.jpg" width="500" data-altupdatedat="1747911167000" data-alt="Figure 6"> 
Figure 6. Example micrographs of “ideal” areas for OpNS imaging. Three protein samples, (A) GroEL (B) Proteasome (C) Spherical HDL, were used as examples to demonstrate the ideal OpNS for imaging. Since stains are unevenly distributed in the specimen, the low-magnification of sample usually appears “cloudy” (leftmost panel). The ideal areas for imaging are normally located in the boundaries of these “clouds”. An example of step-by-step zoom-in one “cloudy” stain region (left middle panel) show the high-magnification images presenting with high-resolution and high contrast images of particles (right middle panel). Selected images of particles show the structure details of proteins (rightmost panel). Bars: 30nm. <a href="https://www.jove.com/files/ftp_upload/51087/51087fig6highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="EM staining procedure diagram with grid incubation, workstation setup, and blotting process steps." fo:content-width="5in" src="/files/ftp_upload/51087/51087fig7highres.jpg" width="500" data-altupdatedat="1747911167000" data-alt="Figure 7"> 
Figure 7. A schematic diagram of OpNS procedures. (A) EM grid incubation station, simple station to incubate the sample solution on the glow-discharged grid. (B) Staining workstation, designed to maintain water washing droplets, and stain droplets above an ice bed, while minimizing stain exposure to light. (C) Overview of staining procedure, illustrating: blotting direction, 3x water rinsing, 3x UF stain exposure, Staining incubation with inverted sample grid on stain droplets, and final backside sample blotting to ensure a thin layer of sample stain solution prior to drying by nitrogen air. <a href="https://www.jove.com/files/ftp_upload/51087/51087fig7highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Cryo-PS analysis with low-pass filtering, masked zoom-in, and crystal structures; electron microscopy resultados." fo:content-width="5in" src="/files/ftp_upload/51087/51087fig8highres.jpg" width="500" data-altupdatedat="1747911167000" data-alt="Figure 8"> 
Figure 8. High-resolution images of 53kDa small protein CETP revealed from a modified OpNS method, cryo-positive-staining (cryo-PS). Five select high-resolution and circular shape soft masked raw images of CETP particles (with reverse contrast) and particle shape masked raw particles (manually reduced noise surrounding raw particle images’ edges, but without modifying any density within the particles) to visualize easily. All-atom and ribbon crystal structure comparison to masked raw particles aligned to similar orientations of each particle to EM images. The high-resolution details of the raw image areas display certain similarity with the crystal structure, while not all crystal structure features can be resolved from raw data. Remarkably, these raw images show similarity in both terminal ends having nearby small circular holes seen in manually reduced noise images (triangles); moreover, the strands in the C-terminal regions within particle images complement crystal structure β-sheets (arrows) and finally, protruding loops on the CETP C-terminal region are akin to in the crystal structure (diamonds). (A) Five select high-resolution and circular shape soft masked raw images of CETP particles (with reverse contrast). (B) Low pass filtering to 10Å. (C) Higher Low pass filtering to 20Å. (D) Masked manually noise reduced images. (E) Crystal structure comparison by ribbon structure. (F) Crystal structure comparison by all atom representation. Bar: 5nm. Part of this work was published in Nature Chemical Biology6. <a href="https://www.jove.com/files/ftp_upload/51087/51087fig8highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Phosphotungstic acid (PTA) concentration effects on virus-like particles; electron microscopy." fo:content-width="5in" src="/files/ftp_upload/51087/51087fig9highres.jpg" width="500" data-altupdatedat="1747911167000" data-alt="Figure 9"> 
Figure 9. Electron micrographs of liposomes showing rouleau formation by conventional NS protocol (PTA stain) under differing salinity. (A) High salinity (~0.5 MNaCl). (B) Regular salinity (~0.25 M NaCl). (C) Low salinity (~0.1 M NaCl). (D) Pure water. Micrographs (above panel) and select windowed particles (below panel) shown. Bars: 100 nm. Window size: 80 nm. The Journal of Lipid Research originally published this work30. <a href="https://www.jove.com/files/ftp_upload/51087/51087fig9highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Optimized Negative Staining: a High-throughput Protocol for Examining Small and Asymmetric Protein Structure by Electron Microscopy at JoVE.com

Optimized Negative Staining: a High-throughput Protocol for Examining Small and Asymmetric Protein Structure by Electron Microscopy

More than half of proteins are small proteins (molecular mass &#60;200 kDa) that are challenging for both electron microscope imaging and three-dimensional reconstructions. Optimized negative staining is a robust and high-throughput protocol to obtain high contrast and relatively high resolution (~1 nm) images of small asymmetric proteins or complexes under different physiological conditions.

Structural determination of proteins is rather challenging for proteins with molecular masses between 40 - 200 kDa. Considering that more than half of ...

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43.4K Views.  The Molecular Foundry. More than half of proteins are small proteins (molecular mass 

Video: Optimized Negative Staining: a High-throughput Protocol for Examining Small and Asymmetric Protein Structure by Electron Microscopy

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 ...

Biochemistry

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

Variations on Negative Stain Electron Microscopy Methods: Tools for Tackling Challenging Systems

Negative stain electron microscopy (EM) allows relatively simple and quick observation of macromolecules and macromolecular complexes through the use of contrast enhancing stain reagent. Although limited in resolution to a maximum of ~18 - 20 &#197;, negative stain EM is useful for a variety of biological problems and also provides a rapid means of assessing samples for cryo-electron microscopy (cryo-EM). The negative stain workflow is straightforward method; the sample is adsorbed onto a substrate, then a stain is applied, blotted, and dried to produce a thin layer of electron dense stain in which the particles are embedded. Individual samples can, however, behave in markedly different ways under varying staining conditions. This has led to the development of a large variety of substrate preparation techniques, negative staining reagents, and grid washing and blotting techniques. Determining the most appropriate technique for each individual sample must be done on a case-by-case basis and a microscopist must have access to a variety of different techniques to achieve the highest-quality negative stain results. Detailed protocols for two different substrate preparation methods and three different blotting techniques are provided, and an example of a sample that shows markedly different results depending on the method used is shown. In addition, the preparation of some common negative staining reagents, and two novel Lanthanide-based stains, is described with discussion regarding the use of each.

Negative stain EM is a powerful technique for visualizing macromolecular structure, but different staining techniques can produce varying results in a sample dependent manner. Here several negative staining approaches are described in detail to provide an initial workflow for tackling the visualization of challenging systems.

Negative stain electron microscopy (EM) allows relatively simple and quick observation of macromolecules and macromolecular complexes through the use of ...

Expression and Purification of the Human Lipid-sensitive Cation Channel TRPC3 for Structural Determination by Single-particle Cryo-electron Microscopy

Transient receptor potential channels (TRPCs) of the canonical TRP subfamily are nonselective cation channels that play an essential role in calcium homeostasis, particularly store-operated calcium entry, which is critical to maintaining proper function of synaptic vesicle release and intracellular signaling pathways. Accordingly, TRPC channels have been implicated in a variety of human diseases including cardiovascular disorders such as cardiac hypertrophy, neurodegenerative disorders such as Parkinson&#39;s disease, and neurologic disorders such as spinocerebellar ataxia. Therefore, TRPC channels represent a potential pharmacologic target in human diseases. However, the molecular mechanisms of gating in these channels are still unclear. The difficulty in obtaining large quantities of stable, homogeneous, and purified protein has been a limiting factor in structure determination studies, particularly for mammalian membrane proteins such as the TRPC ion channels. Here, we present a protocol for the large-scale expression of mammalian ion channel membrane proteins using a modified baculovirus gene transfer system and the purification of these proteins by affinity and size-exclusion chromatography. We further present a protocol to collect single-particle cryo-electron microscopy images from purified protein and to use these images to determine the protein structure. Structure determination is a powerful method for understanding the mechanisms of gating and function in ion channels.

This protocol describes techniques used to determine ion channel structures by cryo-electron microscopy, including a baculovirus system used to efficiently express genes in mammalian cells with minimum effort and toxicity, protein extraction, purification, and quality checking, sample grid preparation and screening, as well as data collection and processing.

Transient receptor potential channels (TRPCs) of the canonical TRP subfamily are nonselective cation channels that play an essential role in calcium ...

Single Particle Cryo-Electron Microscopy: From Sample to Structure

Cryo-electron microscopy (cryoEM) is a powerful technique for structure determination of macromolecular complexes, via single particle analysis (SPA). The overall process involves i) vitrifying the specimen in a thin film supported on a cryoEM grid; ii) screening the specimen to assess particle distribution and ice quality; iii) if the grid is suitable, collecting a single particle dataset for analysis; and iv) image processing to yield an EM density map. In this protocol, an overview for each of these steps is provided, with a focus on the variables which a user can modify during the workflow and the troubleshooting of common issues. With remote microscope operation becoming standard in many facilities, variations on imaging protocols to assist users in efficient operation and imaging when physical access to the microscope is limited will be described.

Structure determination of macromolecular complexes using cryoEM has become routine for certain classes of proteins and complexes. Here, this pipeline is summarized (sample preparation, screening, data acquisition and processing) and readers are directed towards further detailed resources and variables that may be altered in the case of more challenging specimens.

Cryo-electron microscopy (cryoEM) is a powerful technique for structure determination of macromolecular complexes, via single particle analysis (SPA). The ...

Single Particle Electron Microscopy Reconstruction of the Exosome Complex Using the Random Conical Tilt Method

Single particle electron microscopy (EM) reconstruction has recently become a popular tool to get the three-dimensional (3D) structure of large macromolecular complexes. Compared to X-ray crystallography, it has some unique advantages. First, single particle EM reconstruction does not need to crystallize the protein sample, which is the bottleneck in X-ray crystallography, especially for large macromolecular complexes. Secondly, it does not need large amounts of protein samples. Compared with milligrams of proteins necessary for crystallization, single particle EM reconstruction only needs several micro-liters of protein solution at nano-molar concentrations, using the negative staining EM method. However, despite a few macromolecular assemblies with high symmetry, single particle EM is limited at relatively low resolution (lower than 1 nm resolution) for many specimens especially those without symmetry. This technique is also limited by the size of the molecules under study, i.e. 100 kDa for negatively stained specimens and 300 kDa for frozen-hydrated specimens in general.
For a new sample of unknown structure, we generally use a heavy metal solution to embed the molecules by negative staining. The specimen is then examined in a transmission electron microscope to take two-dimensional (2D) micrographs of the molecules. Ideally, the protein molecules have a homogeneous 3D structure but exhibit different orientations in the micrographs. These micrographs are digitized and processed in computers as "single particles". Using two-dimensional alignment and classification techniques, homogenous molecules in the same views are clustered into classes. Their averages enhance the signal of the molecule's 2D shapes. After we assign the particles with the proper relative orientation (Euler angles), we will be able to reconstruct the 2D particle images into a 3D virtual volume.
In single particle 3D reconstruction, an essential step is to correctly assign the proper orientation of each single particle. There are several methods to assign the view for each particle, including the angular reconstitution1 and random conical tilt (RCT) method2. In this protocol, we describe our practice in getting the 3D reconstruction of yeast exosome complex using negative staining EM and RCT. It should be noted that our protocol of electron microscopy and image processing follows the basic principle of RCT but is not the only way to perform the method. We first describe how to embed the protein sample into a layer of Uranyl-Formate with a thickness comparable to the protein size, using a holey carbon grid covered with a layer of continuous thin carbon film. Then the specimen is inserted into a transmission electron microscope to collect untilted (0-degree) and tilted (55-degree) pairs of micrographs that will be used later for processing and obtaining an initial 3D model of the yeast exosome. To this end, we perform RCT and then refine the initial 3D model by using the projection matching refinement method3.

This article describes a standard method to get a three-dimensional (3D) reconstruction of biological macromolecules using negative staining electron microscopy (EM). In this protocol, we explain how to get the 3D structure of the Saccharomyces cerevisiae exosome complex at medium resolution using the random conical tilt reconstruction method (RCT).

Single particle electron microscopy (EM) reconstruction has recently become a popular tool to get the three-dimensional (3D) structure of large ...

Biology

Optimizing Sample Preparation for Cryogenic Electron Microscopy

Cryogenic electron microscopy (cryo-EM) has revolutionized structural biology by enabling the study of macromolecular structures in near-native conditions, suspended in vitreous ice. This technique allows for the high-resolution visualization of proteins and other biomolecules without the need for crystallization, offering significant insights into their function and mechanism. Recent advancements in single-particle analysis, coupled with improved computational data processing, have made cryo-EM an indispensable tool in modern structural biology. Despite its growing adoption, cryo-EM faces persistent challenges that can limit its effectiveness, particularly uneven particle distribution. This issue often leads to poor resolution and reduced accuracy in reconstructed protein structures. This article outlines a simple, practical approach to address this challenge, using the small heat-shock protein from Methanocaldococcus jannaschii (MjsHSP16.5) as an example. The method optimizes sample preparation to minimize preferential adsorption, ensuring more homogeneous particle distribution and higher-quality protein cryo-EM structures. This technique offers valuable guidance for researchers aiming to overcome similar challenges in structural studies.

This protocol presents a practical method using Methanocaldococcus jannaschii (MjsHSP16.5) as an example to address uneven particle distribution through sample preparation optimization, providing a reference for researchers to efficiently elucidate macromolecular structures using cryogenic electron microscopy (cryo-EM).

Cryogenic electron microscopy (cryo-EM) has revolutionized structural biology by enabling the study of macromolecular structures in near-native ...

Nano-fEM: Protein Localization Using Photo-activated Localization Microscopy and Electron Microscopy

Mapping the distribution of proteins is essential for understanding the function of proteins in a cell. Fluorescence microscopy is extensively used for protein localization, but subcellular context is often absent in fluorescence images. Immuno-electron microscopy, on the other hand, can localize proteins, but the technique is limited by a lack of compatible antibodies, poor preservation of morphology and because most antigens are not exposed to the specimen surface. Correlative approaches can acquire the fluorescence image from a whole cell first, either from immuno-fluorescence or genetically tagged proteins. The sample is then fixed and embedded for electron microscopy, and the images are correlated 1-3. However, the low-resolution fluorescence image and the lack of fiducial markers preclude the precise localization of proteins. 
Alternatively, fluorescence imaging can be done after preserving the specimen in plastic. In this approach, the block is sectioned, and fluorescence images and electron micrographs of the same section are correlated 4-7. However, the diffraction limit of light in the correlated image obscures the locations of individual molecules, and the fluorescence often extends beyond the boundary of the cell. 
Nano-resolution fluorescence electron microscopy (nano-fEM) is designed to localize proteins at nano-scale by imaging the same sections using photo-activated localization microscopy (PALM) and electron microscopy. PALM overcomes the diffraction limit by imaging individual fluorescent proteins and subsequently mapping the centroid of each fluorescent spot 8-10. 
We outline the nano-fEM technique in five steps. First, the sample is fixed and embedded using conditions that preserve the fluorescence of tagged proteins. Second, the resin blocks are sectioned into ultrathin segments (70-80 nm) that are mounted on a cover glass. Third, fluorescence is imaged in these sections using the Zeiss PALM microscope. Fourth, electron dense structures are imaged in these same sections using a scanning electron microscope. Fifth, the fluorescence and electron micrographs are aligned using gold particles as fiducial markers. In summary, the subcellular localization of fluorescently tagged proteins can be determined at nanometer resolution in approximately one week.

We describe a method to localize fluorescently tagged proteins in electron micrographs. Fluorescence is first localized using photo-activated localization microscopy on ultrathin sections. These images are then aligned to electron micrographs of the same section.

Mapping the distribution of proteins is essential for understanding the function of proteins in a cell. Fluorescence microscopy is extensively used for ...

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 ...

The CryoAPEX Method for Electron Microscopy Analysis of Membrane Protein Localization Within Ultrastructurally-Preserved Cells

Key cellular events like signal transduction and membrane trafficking rely on proper protein location within cellular compartments. Understanding precise subcellular localization of proteins is thus important for answering many biological questions. The quest for a robust label to identify protein localization combined with adequate cellular preservation and staining has been historically challenging. Recent advances in electron microscopy (EM) imaging have led to the development of many methods and strategies to increase cellular preservation and label target proteins. A relatively new peroxidase-based genetic tag, APEX2, is a promising leader in cloneable EM-active tags. Sample preparation for transmission electron microscopy (TEM) has also advanced in recent years with the advent of cryofixation by high pressure freezing (HPF) and low-temperature dehydration and staining via freeze substitution (FS). HPF and FS provide excellent preservation of cellular ultrastructure for TEM imaging, second only to direct cryo-imaging of vitreous samples. Here we present a protocol for the cryoAPEX method, which combines the use of the APEX2 tag with HPF and FS. In this protocol, a protein of interest is tagged with APEX2, followed by chemical fixation and the peroxidase reaction. In place of traditional staining and alcohol dehydration at room temperature, the sample is cryofixed and undergoes dehydration and staining at low temperature via FS. Using cryoAPEX, not only can a protein of interest be identified within subcellular compartments, but also additional information can be resolved with respect to its topology within a structurally preserved membrane. We show that this method can provide high enough resolution to decipher protein distribution patterns within an organelle lumen, and to distinguish the compartmentalization of a protein within one organelle in close proximity to other unlabeled organelles. Further, cryoAPEX is procedurally straightforward and amenable to cells grown in tissue culture. It is no more technically challenging than typical cryofixation and freeze substitution methods. CryoAPEX is widely applicable for TEM analysis of any membrane protein that can be genetically tagged.

This protocol describes the cryoAPEX method, in which an APEX2-tagged membrane protein can be localized by transmission electron microscopy within optimally-preserved cell ultrastructure.

Key cellular events like signal transduction and membrane trafficking rely on proper protein location within cellular compartments. Understanding precise ...