We are extremely grateful to Peter Knight for helpful discussions and critical review of the manuscript. We would like to thank all of the members of Neil Ranson&#39;s and Stephen Muench&#39;s labs and the Astbury Biostructure Laboratory staff for helpful discussions. This work was funded by the European Research Council (FP7/2007-2013)/ERC Grant Agreement 322408. C-protein was produced using resources provided by a British Heart Foundation grant (BHF PG/13/83/30485). We also thank the Wellcome Trust for equipment funding to support Electron Microscopy in Leeds (090932/Z/09/Z and 094232/Z/10/Z). CS is funded by a Wellcome Trust ISSF Grant.

1. Preparation of EM Grids
<ol>
	<li>Carbon Sheet Method
	<ol>
		<li>Prepare a freshly cleaved mica sheet.
		<ol>
			<li>Gently insert a precision syringe needle or a razor blade at one corner of the mica sheet, a few mm in between the layers. Insert the tool as close to the vertical centre of the sheet as possible to produce two pieces of approximately equal thickness.</li>
			<li>Carefully prise apart the two halves of the mica sheet. Do this can by eye, or under a dissecting microscope.</li>
			<li>Cut off one of the corners of each of the newly cleaved mica sheets. In the event that the sheet turns over in the carbon evaporator during vacuum release, the carbon coated side of the sheet can be identified.</li>
		</ol>
		</li>
		<li>Place the cleaved mica sheet/s in the chamber of a carbon evaporator, with the freshly-cleaved surface facing up.</li>
		<li>Make sure the carbon evaporator is set-up correctly with a properly prepared carbon electrode. 
		NOTE: The method of preparing the carbon rods will vary depending on the specifications of the carbon evaporator. A protocol for one instrument is as follows until step 1.1.5.
		<ol>
			<li>Sharpen a carbon rod with a sharpener to have a sharp tip and then polish it with a paper towel to remove any rough burrs.</li>
			<li>Using fine sandpaper flatten the end of a second rod and again polish it smooth with a paper towel.</li>
			<li>Place the two carbon rods into the evaporator according to manufacturer&#39;s instructions. Ensure the sharpened end on the first rod makes firm contact with the flattened face of the second rod.</li>
		</ol>
		</li>
		<li>Place a small piece of clean, dry filter paper partially under the mica if the carbon thickness is going to be gauged visually. Alternatively, place a white frosted microscope slide with a small dab of vacuum grease alongside the mica to gauge carbon thickness.</li>
		<li>Deposit carbon onto the mica according to manufacturer&#39;s instructions.
		<ol>
			<li>Pump the vacuum down and wait until it is at 10-5 mbar. Set the voltage of the electrode to 4.0 V (up to 5 V may be required depending on the carbon rod source).</li>
			<li>Run multiple short pulses of approximately 3.5 s in duration through the electrode to deposit 1-2 nm thick evaporations of carbon on the mica surface. 
			NOTE: As current is applied to the carbon rod it will glow red and then white. Do not stare at the bright light as this could damage your eyes.</li>
			<li>Allow the carbon to deposit on the mica until the desired thickness is reached, as measured by the carbon evaporator&#39;s thickness gauge or by visual observation of the carbon deposited on the filter paper or microscope slide. Ensure that the final carbon layer is 5-10 nm thick.
			<ol>
				<li>If the carbon thickness is gauged visually compare the frosted part of the microscope slide coated with vacuum grease to the exposed area, it will become darker as more carbon is deposited. 
				NOTE: There is no quantitative method to determine carbon thickness when using this method.</li>
			</ol>
			</li>
		</ol>
		</li>
		<li>Vent the vacuum chamber and remove the carbon-coated mica from the carbon evaporator. 
		NOTE: The carbon-coated mica can be left to settle overnight before proceeding with subsequent steps</li>
		<li>Use one of the two water containers to float the carbon film onto the EM grids: a container with a drain valve at the bottom so water can be drained out and the carbon layer lowered on to the awaiting grids or a lifting rig that the grids can be set on under the water surface, which can subsequently raise the grids up to the carbon film at the water&#39;s surface.</li>
		<li>Fill the container with ultrapure distilled water so that the surface of the water is approximately 5 mm from the top. Clean the surface of water by dragging a sheet or two of lens tissue over the surface to remove any floating particulates.</li>
		<li>Place a piece of clean stainless-steel mesh (1 inch by 2.5 inches is an appropriate size) under the surface of the water.</li>
		<li>Using a pair of fine tweezers, lay clean, dry EM grids face up (according to the manufacturer&#39;s description) on the stainless-steel mesh. Pack the grids together as tightly as possible, but do not allow them to overlap.</li>
		<li>Once the grids are arranged, firmly grip the carbon coated mica sheet in a pair of tweezers or film developing tongs.</li>
		<li>Introduce the mica sheet into the water. Ensure that this is done at a very shallow angle (~10 degrees). 
		NOTE: The mica should break through the water surface and submerge, whilst the carbon film should separate from the mica and float on the water surface. This step should not be performed directly over the grids, to avoid damage and/or contamination.
		<ol>
			<li>To minimize the possibility of the carbon film will not separating from the mica sheet, score around the edge of the mica sheet with a razor blade or cut one corner off with small scissors before introducing it into the water.</li>
		</ol>
		</li>
		<li>Once the carbon film has detached, remove the mica sheet or let it fall to the bottom of the container.</li>
		<li>Using fine tipped tweezers, apply very gentle pressure and with slow movements guide the carbon film over the top of the grids.</li>
		<li>Bring the carbon sheet in contact with the surface of the grids either by slowly draining the water or raising the lifting ring, depending on the type of apparatus used.</li>
		<li>Carefully lift the stainless-steel mesh (now with carbon-coated grids) from the apparatus and wick away some of the excess water using a piece of filter paper. Ensure that this is done by touching the filter paper to the very edge of the steel mesh but not coming in contact with the grids or carbon film.</li>
		<li>Place the mesh of grids in a petri dish containing a dry piece of filter paper and allow to it dry completely. 
		NOTE: This is best affected by drying overnight at room temperature, but the step can be expedited by placing the grids in an oven at approximately 60 &#176;C.</li>
	</ol>
	</li>
	<li>Float and coat (direct carbon deposition). This method has been described in detail previously16
	<ol>
		<li>Completely fill a clean large glass bowl to the brim with distilled water so a meniscus forms at the top.</li>
		<li>Apply a single drop of collodion solution (nitrocellulose in amyl acetate) to the surface of the water using a clean pasteur pipette, allow the droplet to spread out and dry completely. Once dry a thin layer of collodion floating on top of the water surface will be visible.</li>
		<li>Gently remove the collodion layer using a toothpick to remove dust or other contamination from the surface of the water.</li>
		<li>Apply a second collodion droplet to the water and allow it to spread out and dry for 2-3 minutes. 
		NOTE: Repeat steps 1.2.3-1.2.4 until a flat and wrinkle free sheet of collodion is obtained.</li>
		<li>Using a pair of fine tweezers place EM grids face down (according to the manufacturer&#39;s description) on the floating collodion sheet. Pack the grids together tightly in a hexagonal array, but do not allow them to overlap. 
		NOTE: If a grid is misplaced or placed upside down it is generally best to leave it in place rather than risk damaging the collodion sheet when trying to move it.</li>
		<li>Once all of the grids are placed, gently lay a sheet of filter paper over them. Allow the paper to become saturated by capillary action. 
		NOTE: Any size or thickness of filter paper is appropriate if it completely covers the grids.</li>
		<li>Use a toothpick to remove any collodion film that extends beyond the filter paper.</li>
		<li>Grip the filter paper at the edge and peel it from the water surface. 
		NOTE: The grids should stay adhered to the paper.</li>
		<li>Place the paper flat and collodion-face up in a petri dish and allow it to dry completely.</li>
		<li>Place the filter paper with the grids in the chamber of a carbon evaporator with a properly prepared carbon electrode as detailed in 1.1.2.</li>
		<li>Follow the carbon evaporation procedure as described for the carbon sheet method in</li>
	</ol>
	</li>
	<li>Allow several seconds between pulses to avoid overheating and damaging the nitrocellulose sheet. 
	NOTE: If desired the polymer layer can be removed after the grids have been carbon coated, although this step is rarely necessary. Place the grids carbon side up on a fresh piece of filter paper and put several drops of acetone on the paper near, but not on, the grids. Allow the acetone to spread out under the grids and dissolve and absorb the polymer layer.</li>
</ol>
2. Preparation of Negative Staining Reagents
<ol>
	<li>Preparation of Uranyl Acetate
	<ol>
		<li>Bring a small volume of ultrapure water to a boil and allow it to boil for 10 min to thoroughly degas. Allow it to cool slightly, and then use it to dissolve uranyl acetate (UA) at 1-2 % (w/v). 
		NOTE: Perform this procedure in a fume cupboard and with appropriate personal protective equipment.</li>
		<li>After the solution has cooled, filter through a 0.2 &#181;m syringe filter or filter paper.</li>
		<li>Store the UA protected from light and at 4 &#730;C. The solution is stable for up to 1 year.</li>
	</ol>
	</li>
	<li>Preparation of Uranyl Formate from Powder. This Method has been Described in Detail Previously8
	<ol>
		<li>Dissolve 20 mg uranyl formate (UF) powder in 2 mL of boiled degassed ultrapure water (as in step 2.1.1) by stirring.</li>
		<li>While continuing to stir, add 8 &#181;L of 5 M NaOH, the solution should change to a darker yellow colour, but no precipitate should form.</li>
		<li>Filter the solution through a 0.2 &#181;m syringe filter.</li>
		<li>Store the UF stain protected from light. Discard the stain should if any precipitate or brown discoloration is observed. The solution is only stable for 1-2 days.</li>
	</ol>
	</li>
	<li>Preparation of Uranyl Formate from Uranyl Acetate
	<ol>
		<li>Precipitate 1 mL of 1% (w/v) UA stain by adding 100 &#181;L of 1 M NaOH.</li>
		<li>Centrifuge the mixture for 2 min at maximum speed in a benchtop centrifuge.</li>
		<li>Discard any supernatant and dissolve the precipitate in 100 &#181;L of 5% (v/v) formic acid by vigorous vortexing.</li>
		<li>Dilute to a final volume of 1 mL with 900 &#181;L ultrapure water to yield the UF stain in 0.5% (v/v) formic acid.</li>
		<li>Store the UF stain protected from light. Discard the stain if any precipitate or brown discoloration is observed.</li>
	</ol>
	</li>
	<li>Preparation of Other Staining Reagents
	<ol>
		<li>Preparation of Lanthanide Acetate Stains
		<ol>
			<li>Dissolve Samarium Acetate (SmAc), Gadolinium Acetate (GdAc), Thulium Acetate (TmAc), or Erbium Acetate (ErAc) at 1-2% (w/v) in ultrapure water. 
			NOTE: If samples show positive staining or poor adherence to the grid when using these stains, they can be acidified with up to 0.5% (v/v) formic acid. Positive staining results in the sample appearing as a dark object surrounded by a white halo. Poor adherence to the grid will result in fewer molecules than expected being observed on the grid.</li>
		</ol>
		</li>
		<li>Preparation of Ammonium Molybdate and Sodium Phosphotungstate
		<ol>
			<li>Dissolve the stain at 1-3% (w/v) in ultrapure water. Adjust the pH to 7.0 using 5 M NaOH if desired.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
3. Adsorbing Samples to the Carbon Substrate and Staining
<ol>
	<li>Preparation of the Grid Surface for Sample Application by Rendering it Hydrophilic
	<ol>
		<li>Place the grid facing up on a microscope slide in a glow discharge unit.</li>
		<li>Treat the grid for a minimum of 30 s at 10 mA. 
		NOTE: The exact method of glow discharge will depend on the specifications of the particular piece of equipment used.</li>
		<li>Alternatively this may be accomplished by UV irradiation for 10 minutes using a benchtop UV lamp4.</li>
	</ol>
	</li>
	<li>Side Blotting Method. This Method has been Described in Detail Previously8
	<ol>
		<li>Grip the edge of the grid with a pair of negative pressure tweezers, and apply 3-5 &#181;L of sample to the support surface.</li>
		<li>Allow the sample to adsorb to the grid surface for 10 s to 1 min. Optimize the adsorption time must for individual samples.</li>
		<li>Touch the edge of the grid to a sheet of filter paper and allow capillary action to pull off the liquid.</li>
		<li>Optional: Wash the grid. Place 50 &#181;L drops of ultrapure water or appropriate volatile buffer solution on a sheet of laboratory film. Gently touch the carbon surface of the grid to the drop and lift off a small droplet onto the surface of the grid. Touch the edge of the grid to a sheet of filter paper and allow capillary action to pull off the liquid.</li>
		<li>Repeat this wash step as many times as desired.</li>
		<li>Place two 50 &#181;L drops of staining reagent on a sheet of laboratory film.</li>
		<li>Gently touch the carbon surface of the grid to the drop and lift off a small droplet onto the top surface of the grid. 
		NOTE: If the stain migrates to the back of the grid then the grid should be discarded.</li>
		<li>Touch the edge of the grid to a sheet of filter paper and allow capillary action to draw off the liquid. Perform this staining step twice.</li>
		<li>Allow the grid to air dry or dry under an incandescent lamp.</li>
	</ol>
	</li>
	<li>Flicking Method
	<ol>
		<li>Grip the edge of the grid with a pair of negative pressure tweezers, and apply 3-5 &#181;L of sample to the support surface.</li>
		<li>Holding the tweezers in one hand, so that the grid is angled at approximately 45&#176; facing away, rapidly flick the wrist of that hand to &#39;flick off&#39; the majority of the droplet that is on top of the grid.</li>
		<li>Optional: Using a glass Pasteur pipette apply a drop of wash solution to the support surface and &#39;flick off&#39; as in 3.2.2. Repeat as necessary.</li>
		<li>Using a glass Pasteur pipette apply a drop of staining reagent to the support surface and &#39;flick off&#39; as in 3.2.2. Repeat 1-3 times dependent on stain depth required for visualization of specimen. 
		NOTE: This is not the only factor that attributes to final stain depth (see discussion).</li>
		<li>Remove excess stain by touching the torn edge of a piece of filter paper to the edge of the grid.</li>
		<li>Allow the grid to air dry or dry under an incandescent lamp.</li>
	</ol>
	</li>
	<li>Rapid Flushing Method
	<ol>
		<li>Draw 30-70 &#181;L of stain (1 % UA usually used) into the tip of a 200 &#181;L pipette, turn the volume dial to draw up 5 &#181;L of air, then draw up wash/mixing reagent (5-30 &#181;L), if required, followed by another small air gap and then draw up 5 &#181;L of sample.</li>
		<li>Grip the edge of a grid with a pair of negative pressure tweezers, holding the tweezers so that the grid is angled at approximately 45&#176; facing away from the researcher, eject the entire contents of the pipette tip across the face of the carbon-coated EM grid.</li>
		<li>Remove excess stain by touching the torn edge of a piece of filter paper to the edge of the grid.</li>
		<li>Allow the grid to air dry or dry under an incandescent lamp. 
		NOTE: For all methods it is advisable to slide the torn edge of a sheet of filter paper along the forceps until it reaches the grid as this removes solution trapped between the two sides of the forceps, which can pull the dried grid into the jaws of the forceps once they are opened. The grid in the tweezers can also be placed on the edge of a fume hood to dry. The constant airflow can help produce more even staining.</li>
	</ol>
	</li>
</ol>

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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+introduction+to+sample+preparation+and+imaging+by+cryo-electron+microscopy+for+structural+biology.">An introduction to sample preparation and imaging by cryo-electron microscopy for structural biology.</a> Methods. 100, 3-15 (2016).</li><li>Ha, J. Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Molecular+architecture+of+the+complete+COG+tethering+complex.">Molecular architecture of the complete COG tethering complex.</a> Nat Struct Mol Biol. 23 (8), 758-760 (2016).</li><li>Burgess, S. A., Walker, M. L., Thirumurugan, K., Trinick, J., Knight, P. 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The authors declare no competing financial interests.

This manuscript describes multiple methods for negative staining of samples for electron microscopy using a variety of staining reagents, including two novel lanthanide reagents (TmAc and ErAc). Many of the steps of the negative staining process must be optimized for individual samples including the choice of stain, amount of washing required if any, and the blotting technique. This manuscript thus provides a basis for microscopists to develop their own workflows for tackling the negative-staining of challenging systems.
The choice of stain is highly sample dependent. Samples that are especially sensitive to low pH may be degraded by UA and/or UF, despite the fixative properties of these stains19. In these cases, lanthanide based stains such as TmAc or ErAc may be more appropriate, although the overall pH of the preparation must be kept below the isoelectric point of the sample protein to help prevent positive staining. This can be accomplished by acidifying the stain with acetic acid if necessary. For especially low pH sensitive samples, anionic tungstate or molybdate stains may be more effective. Although these stains have been found to induce the formation of artifacts in some cases, such as the formation of rouleaux in lipoprotein samples20. Again, the pH of the stain may need to be adjusted, this time to above the isoelectric point of the sample, to prevent positive staining.
Washing of the sample prior to staining may be necessary if the buffer in which the specimen is maintained has a high salt or phosphate component. In many cases, washing can be performed with ultrapure water but for more sensitive samples, which may degrade or undergo structural changes when exposed to water alone, washing may need to be performed with a volatile buffer of low ionic strength8. Even under carefully controlled conditions, washing can result in some structural rearrangement on the carbon surface21.
The method by which a grid is prepared in terms of sample adsorption, blotting and staining can also significantly affect what is observed. The most appropriate method is thus, again, highly sample dependent. C-protein, for example, is observed as a globular ring-like structure following side-blot staining, but this appears to be an artifact of the staining process, as revealed when grids are prepared by the flicking method (or by the rapid flushing method) (Figure 3). In the flicking and rapid flushing methods, the time the sample has to interact with the carbon support surface before fixation is minimized15. The sample also experiences fewer forces from the receding meniscus upon blotting before fixation. This means that structural changes in the specimen that could occur upon prolonged absorption time on the carbon film or through capillary action are minimized. The rapid flushing method can also be used for time-resolved analysis of specimens. The sample can be mixed with a ligand or additive within a pipette tip for a set period of time before application to a grid or only momentarily on the grid surface before fixation within milliseconds.
The depth of stain required to provide optimal images of a particular specimen is again sample dependent2. If the stain is too shallow, molecules can be damaged by the electron beam but if the stain is too thick structural features can be lost. Stain depth is influenced by multiple factors such as hydrophilicity of the grid surface, evenness of the carbon layer, the amount of stain applied to the grid, the length of time stain is in contact with the grid prior to blotting, the extent of blotting and the time it takes for the grid to completely dry. A grid will never have an even distribution of stain across its entirety and therefore areas of the grid appropriate for imaging need to be selected carefully. Indeed, grids often vary in quality even when prepared on the same day under the same conditions. A good example of how variation in stain depth affects the appearance of molecules and the appropriate stain depth for imaging is provided by Burgess et al5.
Despite negative staining being a very versatile, quick, and simple method, not all biological specimens are amenable to visualization by this method. Fragile assemblies can collapse or disassemble upon adsorption, staining or drying on the EM grid22. Negative staining can also lead to flattening of molecules and induce preferred orientations of molecules on the carbon support film7.
Negative stain is a valuable tool for assessment of specimens in its own right and also prior to cryo-EM analysis but many of the physical forces the sample encounters during the process are poorly understood. Therefore, the best approach to use is highly sample dependent and must be determined by trial-and-error rather than taught following a fixed protocol.

An erratum was issued for:&#160;<a href="https://www.jove.com/video/57199/variations-on-negative-stain-electron-microscopy-methods-tools-for">Variations on Negative Stain Electron Microscopy Methods: Tools for Tackling Challenging Systems</a>. An&#160;author&#160;name&#160;was updated.
One of the authors&#39; names was corrected from:
Matthew G. Iadaza
to:
Matthew G. Iadanza

An erratum was issued for:&#160;<a href="https://www.jove.com/video/57199/variations-on-negative-stain-electron-microscopy-methods-tools-for">Variations on Negative Stain Electron Microscopy Methods: Tools for Tackling Challenging Systems</a>. An&#160;author&#160;name&#160;was updated.

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

Despite recent attention to the resolution revolution resulting from significant advances in cryo-electron microscopy1 (cryo-EM), negative stain EM remains a powerful technique and a crucial component of electron microscopists&#39; toolbox. Negative staining still remains the best method for rapid assessment of a sample before optimizing cryo-grid conditions2. The high contrast and speed of grid preparation of negative stained samples makes it ideal for assessing sample purity, concentration, heterogeneity, and conformational flexibility3. Many biologically informative structures have resulted from negative stain reconstructions, despite the technique&#39;s resolution being limited to ~18 &#197; resolution4,5,6, and some samples yield better results in stain than cryo-EM for a variety of reasons7.
In negative stain EM, the particle of interest is adsorbed onto the surface of an EM grid and enveloped by an amorphous matrix of electron dense stain compound. A high relative contrast is produced between the background and the particle of interest, with the particle being less electron dense than the surrounding stain8. The particles appear as light areas because of their low electron scattering power relative to the dense surrounding stain, which scatters the electrons more and appears darker. Substructural features of particles can be deduced from the detailed examination of resultant images as stain will penetrate into any crevice and produce irregular contrast detail9.
The negative staining process begins with the preparation of a support substrate on which the sample particles are captured, and the layer of dried stain supported. The most commonly used support substrate is a layer of amorphous carbon, sometimes supported by a thin layer of polyvinyl (e.g. Formvar) or nitrocellulose (e.g. Collodion) polymer. These substrates can be purchased commercially or prepared in-house using the protocols described below.
After the support substrate is prepared, the sample can be applied, and the excess solution blotted off. Samples should be suspended in a suitable buffer for negative-staining. It is best to avoid the use of phosphate buffer and high salt concentrations, which can give rise to crystalline precipitates that can obscure the specimen. Reducing agents, detergents, sucrose, glycerol, and high concentrations of nucleotide should also be avoided as they also affect stain quality4. When the buffer composition cannot be changed, washing the surface of the EM grid with water or a more suitable buffer after adsorption and prior to staining may reduce the formation of buffer related artifacts and generally improve the stain background. If buffer artifacts are suspected, it can be informative to stain a buffer-only grid to determine if the buffer components are the source of the observed artifacts.
After the sample is adsorbed, and blotted and washed if necessary, a staining reagent is applied. A variety of reagents have been found to be effective negative stains (Table 1), but the stain must be chosen to suit the sample. A &#39;halo&#39; of stain forms around the particle due to both the displacement of the stain molecules by the hydrophobic regions of the protein and repulsion by charged groups. Therefore, the stain must be chosen so that the protonation state of any potential charged groups on the protein is the same as the stain at the working pH. Opposite charges on the surface of the protein can contribute to a positive staining effect, which although a useful technique in its own right10 is not in the scope of this paper. The most commonly used negative staining reagents are uranyl acetate and uranyl formate. These stains have a relatively fine grain size (4 - 5 &#197;)9 and provide higher resolution images over other stains such as phospho-tungstates (8 - 9 &#197; grain size)9,11, ammonium molybdate11, and some lanthanide-based stains12. Uranyl acetate and formate also act as a fixative, preserving many protein-protein interactions on a millisecond time scale13, although the low pH of the stain and its propensity to precipitate at physiological pH may be detrimental to some samples14. Despite their utility, the uranyl salts also present logistical challenges as they are both toxic and mildly radioactive, which can require special handling, storage, and disposal requirements, which leads some users to seek non-radioactive alternatives.
There are a large variety of methods described for substrate preparation, sample application, and staining of EM grids. The most appropriate method to use is sample dependent and can be difficult to ascertain when tackling a new system. This manuscript describes two methods of substrate preparation and three blotting methods; side blotting, flicking5,and rapid flushing15. Side-blotting is the simplest of the methods described. Both the flicking method and the rapid flushing method are more difficult to implement but limit the contact time of the sample with the support film before fixation and have been shown to ameliorate formation of stain artifacts for some samples5. The goal of this manuscript is thus to provide an initial workflow for tackling the visualization of challenging systems by negative-stain EM.

<table><tbody><tr><td>200 mesh copper EM grids</td><td>Sigma-Aldrich</td><td>G4776-1VL</td><td>Other materials and/or mesh sizes can also be used</td></tr><tr><td>Ammonium Molybdate</td><td>Sigma-Aldrich</td><td>277908</td><td></td></tr><tr><td>Carbon evaporator</td><td>Ted Pella Inc.</td><td>9620</td><td>Cressington 208 or equivalent</td></tr><tr><td>Collodion solution 2% in amyl acetate</td><td>Sigma-Aldrich</td><td>9817</td><td></td></tr><tr><td>Dumont #5 negative pressure tweezers</td><td>World Precision Instruments</td><td>501202</td><td>Or other tweezers as preferred</td></tr><tr><td>Erbium Acetate</td><td>Sigma-Aldrich</td><td>325570</td><td></td></tr><tr><td>Gadolinium Acetate</td><td>Sigma-Aldrich</td><td>325678</td><td></td></tr><tr><td>Mica Sheets. 75x25x0.15mm.</td><td>AGAR Scientific</td><td>AGG250-1</td><td></td></tr><tr><td>Microscope slides, white frosted</td><td>Fisher Scientific</td><td>12607976</td><td>Or equivalent</td></tr><tr><td>Parafilm</td><td>Fisher Scientific</td><td>10018130</td><td>Or equivalent</td></tr><tr><td>Pasteur pipette (glass)</td><td>Fisher Scientific</td><td>10343663</td><td>Or equivalent</td></tr><tr><td>Razor blade</td><td>Fisher Scientific</td><td>11904325</td><td>Or equivalent</td></tr><tr><td>Sandpaper</td><td>Hardware store</td><td></td><td>Wet and dry sandpaper with grit finer that 200 (600 suggested)</td></tr><tr><td>Samarium Acetate</td><td>Sigma-Aldrich</td><td>325872</td><td></td></tr><tr><td>Sodium Hydroxide</td><td>Sigma-Aldrich</td><td>1.06462</td><td></td></tr><tr><td>Sodium Phosphotungstate</td><td>Sigma-Aldrich</td><td>P6395</td><td></td></tr><tr><td>Stainless Steel Mesh, 150x150 mm (cut to size).</td><td>AGAR Scientific</td><td>AGG252</td><td></td></tr><tr><td>Thulium Acetate</td><td>Sigma-Aldrich</td><td>367702</td><td></td></tr><tr><td>Two Step Carbon Rod Sharper, for 1/4&quot; rods</td><td>Ted Pella Inc.</td><td>57-10</td><td>Or equivalent for carbon evaporator used</td></tr><tr><td>Ultra pure water</td><td></td><td></td><td></td></tr><tr><td>Uranyl Acetate</td><td>Electron Microscopy Sciences</td><td>22400</td><td></td></tr><tr><td>Uranyl Formate</td><td>Electron Microscopy Sciences</td><td>22450</td><td></td></tr><tr><td>Vacuum grease</td><td>Fisher Scientific</td><td>12719406</td><td>Or equivalent</td></tr><tr><td>Whatman #1 Filter paper.</td><td>Fisher Scientific</td><td>1001 090</td><td>Or equivalent</td></tr><tr><td>Whatman #40 filter paper</td><td>Fisher Scientific</td><td>10674122</td><td>Or equivalent</td></tr></tbody></table>

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.

All of the staining reagents tested produced negative staining to some degree, with UF yielding the samples with the greatest contrast and sharpest, most detailed particles. For deeply embedded samples (Figure 1) lanthanide based stains ErAc and TmAc produced negative staining of equivalent quality to UA as judged by the apparent contrast and sharpness of the stained particles, with TmAc producing clearer, more crisp images than ErAc. Although the larger grain size of TmAc becomes apparent at high magnification, when Tobacco Mosaic Virus (TMV) particles were stained with 1% TmAc the ~23 &#197; repeat of the TMV particle17 was still clearly visible by eye and as a meridional layer line in the Fourier transform of the raw image. None of the other lanthanide stains tested, ErAc, SmAc, or GdAc, were able to resolve this feature. Class averages were generated by extracting overlapping segments from TMV particles where the helical repeat was visible. The extracted segments were then aligned and classified using RELION18 to better visualize the periodic feature (Figure 2).
Some samples are especially sensitive to the method of staining, such as the muscle derived C-protein. C-protein, which consists of a flexible string of Ig and Fn-like domains, produces significantly different images by negative-stain EM dependent on the method of staining used (Figure 3). When using the side-blotting method, collapsed ring-like structures are observed, whereas when stained by the rapid flushing or flicking methods, C-protein is observed as a series of domains that resemble beads on a string.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Reagent</td>
			<td>Concentration</td>
			<td>pH </td>
			<td>Type</td>
		</tr>
		<tr>
			<td>Ammonium Molybdate</td>
			<td>1 - 2 %</td>
			<td>5 &#8211; 7</td>
			<td>Anionic</td>
		</tr>
		<tr>
			<td>Erbium acetate (ErAc)</td>
			<td>1 &#8211; 2%</td>
			<td>6</td>
			<td>Cationic</td>
		</tr>
		<tr>
			<td>Gadolinium Acetate (GdAc)</td>
			<td>1 &#8211; 2%</td>
			<td>6</td>
			<td>Cationic</td>
		</tr>
		<tr>
			<td>Methylamine tungstate</td>
			<td>2%</td>
			<td>6 &#8211; 7</td>
			<td>Anionic</td>
		</tr>
		<tr>
			<td>Samarium acetate (SmAC)</td>
			<td>1%</td>
			<td>6</td>
			<td>Cationic</td>
		</tr>
		<tr>
			<td>Sodium silicotungstate</td>
			<td>1 &#8211; 5 %</td>
			<td>5 &#8211; 8</td>
			<td>Anionic</td>
		</tr>
		<tr>
			<td>Sodium phosphotungstate</td>
			<td>1 -3 %</td>
			<td>5 &#8211; 8</td>
			<td>Anionic</td>
		</tr>
		<tr>
			<td>Thulium Acetate (TmAc)</td>
			<td>1 &#8211; 2%</td>
			<td>6</td>
			<td>Cationic</td>
		</tr>
		<tr>
			<td>Uranyl Acetate (UA)</td>
			<td>1 &#8211; 3%</td>
			<td>3 &#8211; 4</td>
			<td>Cationic</td>
		</tr>
		<tr>
			<td>Uranyl Formate (UF)</td>
			<td>0.75 &#8211; 1%</td>
			<td>3 &#8211; 4</td>
			<td>Cationic</td>
		</tr>
	</tbody>
</table>
Table 1: Some common negative staining reagents.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57199/57199fig1.jpg" /> 
Figure 1: Example micrographs of Tobacco Mosaic Virus stained with various negative stain reagents (A) 1% UF (B) 2.5% TmAc (C) 2.5% ErAc. (D) 1% UA (E) 2.5% GdAc and (F) 2.5% SmAc. Scale bars are 100 nm. Representative images from multiple replicates with multiple areas imaged per replicate. <a href="https://cloudflare.jove.com/files/ftp_upload/57199/57199fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57199/57199fig2.jpg" /> 
Figure 2: Staining Tobacco Mosaic Virus with Thulium acetate (A) High magnification of area from a micrograph of TMV stained with 1% TmAc. Scale bar is 20 nm. (B) Class average of extracted TMV segments. (C) Fourier transform of the image in panel A showing layer line reflections at ~23 &#197;. <a href="https://cloudflare.jove.com/files/ftp_upload/57199/57199fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/57199/57199fig3.jpg" /> 
Figure 3: Effects of blotting method on the conformation of C-protein. (A) C-protein stained with UA using the side blot method and (B) flicking method. Upper panel scale bar is 50 nm, lower panel scale bar is 20 nm. Representative images from multiple replicates with multiple areas imaged per replicate. <a href="https://cloudflare.jove.com/files/ftp_upload/57199/57199fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Variations on Negative Stain Electron Microscopy Methods: Tools for Tackling Challenging Systems at JoVE.com

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

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 ...

variations-on-negative-stain-electron-microscopy-methods-tools-for

Research

JoVE Journal

Biochemistry

32.6K Views.  University of Leeds. Negatieve vlek EM is een krachtige techniek voor het visualiseren van macromoleculaire structuur, maar verschillende kleuring technieken wisselende resultaten kunnen produceren in een monster afhankelijke wijze. Hier zijn verschillende negatieve kleuring benaderingen in detail te bieden een eerste workflow voor het aanpakken van de visualisatie van uitdagende systemen beschreven.

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

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.

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 ...

Environment

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 ...

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

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 ...

Utilization of Capsules for Negative Staining of Viral Samples within Biocontainment

Transmission electron microscopy (TEM) is used to observe the ultrastructure of viruses and other microbial pathogens with nanometer resolution. Most biological materials do not contain dense elements capable of scattering electrons to create an image; therefore, a negative stain, which places dense heavy metal salts around the sample, is required. In order to visualize viruses in suspension under the TEM they must be applied to small grids coated with a transparent surface only nanometers thick. Due to their small size and fragility, these grids are difficult to handle and easily moved by air currents. The thin surface is easily damaged, leaving the sample difficult or impossible to image. Infectious viruses must be handled in a biosafety cabinet (BSC) and some require a biocontainment laboratory environment. Staining viruses in biosafety levels (BSL)-3 and -4 is especially challenging because these environments are more turbulent and technicians are required to wear personal protective equipment (PPE), which decreases dexterity. 
In this study, we evaluated a new device to assist in negative staining viruses in biocontainment. The device is a capsule that works as a specialized pipette tip. Once grids are loaded into the capsule, the user simply aspirates reagents into the capsule to deliver the virus and stains to the encapsulated grid, thus eliminating user handling of grids. Although this technique was designed specifically for use in BSL-3 or -4 biocontainment, it can ease sample preparation in any lab environment by enabling easy negative staining of virus. This same method can also be applied to prepare negative stained TEM specimens of nanoparticles, macromolecules and similar specimens.

This protocol provides instruction for negative staining virus samples which can easily be used in BSL-2, -3, or -4 laboratories. It includes the use of an innovative processing capsule, which protects the transmission electron microscopy grid and provides the user easier handling in the more turbulent environments within biocontainment.

Transmission electron microscopy (TEM) is used to observe the ultrastructure of viruses and other microbial pathogens with nanometer resolution. Most ...

Immunology and Infection

Analysis of Brain Mitochondria Using Serial Block-Face Scanning Electron Microscopy

Human brain is a high energy consuming organ that mainly relies on glucose as a fuel source. Glucose is catabolized by brain mitochondria via glycolysis, tri-carboxylic acid (TCA) cycle and oxidative phosphorylation (OXPHOS) pathways to produce cellular energy in the form of adenosine triphosphate (ATP). Impairment of mitochondrial ATP production causes mitochondrial disorders, which present clinically with prominent neurological and myopathic symptoms. Mitochondrial defects are also present in neurodevelopmental disorders (e.g. autism spectrum disorder) and neurodegenerative disorders (e.g. amyotrophic lateral sclerosis, Alzheimer&#39;s and Parkinson&#39;s diseases). Thus, there is an increased interest in the field for performing 3D analysis of mitochondrial morphology, structure and distribution under both healthy and disease states. The brain mitochondrial morphology is extremely diverse, with some mitochondria especially those in the synaptic region being in the range of &#60;200 nm diameter, which is below the resolution limit of traditional light microscopy. Expressing a mitochondrially-targeted green fluorescent protein (GFP) in the brain significantly enhances the organellar detection by confocal microscopy. However, it does not overcome the constraints on the sensitivity of detection of relatively small sized mitochondria without oversaturating the images of large sized mitochondria. While serial transmission electron microscopy has been successfully used to characterize mitochondria at the neuronal synapse, this technique is extremely time-consuming especially when comparing multiple samples. The serial block-face scanning electron microscopy (SBFSEM) technique involves an automated process of sectioning, imaging blocks of tissue and data acquisition. Here, we provide a protocol to perform SBFSEM of a defined region from rodent brain to rapidly reconstruct and visualize mitochondrial morphology. This technique could also be used to provide accurate information on mitochondrial number, volume, size and distribution in a defined brain region. Since the obtained image resolution is high (typically under 10 nm) any gross mitochondrial morphological defects may also be detected.

Mitochondrial visualization and analysis from mammalian brain tissue is a challenging task. Here, we describe how three dimensional (3D) reconstruction analysis from the serial block-face scanning electron microscopy (SBFSEM) can be used to gain insights on the morphological and volumetric analysis of this critical energy generating organelle.

Human brain is a high energy consuming organ that mainly relies on glucose as a fuel source. Glucose is catabolized by brain mitochondria via glycolysis, ...

Neuroscience

Biological Sample Preparation by High-pressure Freezing, Microwave-assisted Contrast Enhancement, and Minimal Resin Embedding for Volume Imaging

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the imaging modality in a focused ion beam scanning electron microscope (FIB-SEM), which is used to obtain stacks of sequential images for 3D reconstruction and modelling. High-pressure freezing (HPF) allows close to native structural preservation, but the subsequent freeze substitution often does not provide sufficient contrast, especially for a bigger specimen, which is needed for high-quality imaging in the SEM required for 3D reconstruction. Therefore, in this protocol, after the freeze substitution, additional contrasting steps are carried out at room temperature. Although these steps are performed in a microwave, it is also possible to follow traditional bench processing, which requires longer incubation times.The subsequent embedding in minimal amounts of resin allows for faster and more precise targeting and preparation inside the FIB-SEM. This protocol is especially useful for samples that require preparation by high-pressure freezing for a reliable ultrastructural preservation but do not gain enough contrast during the freeze substitution for volume imaging using FIB-SEM. In combination with the minimal resin embedding, this protocol provides an efficient workflow for the acquisition of high-quality volume data.

Here we present a protocol for combining two sample processing techniques, high-pressure freezing and microwave-assisted sample processing, followed by minimal resin embedding for acquiring data with a focused ion beam scanning electron microscope (FIB-SEM). This is demonstrated using a mouse tibial nerve sample and Caenorhabditis elegans.

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the ...

Biology

Microscopy Techniques for Interpreting Fungal Colonization in Mycoheterotrophic Plants Tissues and Symbiotic Germination of Seeds

Structural botany is an indispensable perspective to fully understand the ecology, physiology, development, and evolution of plants. When researching mycoheterotrophic plants (i.e., plants that obtain carbon from fungi), remarkable aspects of their structural adaptations, the patterns of tissue colonization by fungi, and the morphoanatomy of subterranean organs can enlighten their developmental strategies and their relationships with hyphae, the source of nutrients. Another important role of symbiotic fungi is related to the germination of orchid seeds; all Orchidaceae species are mycoheterotrophic during germination and seedling stage (initial mycoheterotrophy), even the ones that photosynthesize in adult stages. Due to the lack of nutritional reserves in orchid seeds, fungal symbionts are essential to provide substrates and enable germination. Analyzing germination stages by structural perspectives can also answer important questions regarding the fungi interaction with the seeds. Different imaging techniques can be applied to unveil fungi endophytes in plant tissues, as are proposed in this article. Freehand and thin sections of plant organs can be stained and then observed using light microscopy. A fluorochrome conjugated to wheat germ agglutinin can be applied to the fungi and co-incubated with Calcofluor White to highlight plant cell walls in confocal microscopy. In addition, the methodologies of scanning and transmission electron microscopy are detailed for mycoheterotrophic orchids, and the possibilities of applying such protocols in related plants is explored. Symbiotic germination of orchid seeds (i.e., in the presence of mycorrhizal fungi) is described in the protocol in detail, along with possibilities of preparing the structures obtained from different stages of germination for analyses with light, confocal, and electron microscopy.

This protocol aims to provide detailed procedures for collecting, fixing, and maintaining mycoheterotrophic plant samples, applying different microscopy techniques such as scanning and transmission electron microscopy, light, confocal, and fluorescence microscopy to study fungal colonization in plants tissues and seeds germinated with mycorrhizal fungi.

Structural botany is an indispensable perspective to fully understand the ecology, physiology, development, and evolution of plants. When researching ...

Capsule-Based Negative Staining of Virus Samples on TEM Grids: A Heavy Metal Staining Technique to Visualize the Ultrastructure of Enveloped Viruses

Source: Blancett, C. D., et al. <a href="https://www.jove.com/v/56122">Utilization of Capsules for Negative Staining of Viral Samples within Biocontainment.</a> J. Vis. Exp. (2017)
In this video, we demonstrate the negative staining of enveloped virus samples using a processing capsule within the biocontainment environment. The negatively stained viruses can be visualized using transmission electron microscopy to investigate viral structure and morphology with nanometer resolution.

Source: Blancett, C. D., et al. Utilization of Capsules for Negative Staining of Viral Samples within Biocontainment. J. Vis. Exp. (2017)
In this video, ...

JoVE Encyclopedia of Experiments

Biological Techniques

Staining Techniques

Biomolecule staining

Negative Staining of Purified Exosomes: A Procedure to Visualize Exosome Morphology Using Transmission Electron Microscopy

Source: Jung, M. K. et al. <a href="https://www.jove.com/video/56482" target="_blank">Sample Preparation and Imaging of Exosomes by Transmission Electron Microscopy.</a> J. Vis. Exp. (2018)
This video describes a procedure for negative staining of purified exosomes. The negatively stained exosomes are imaged using transmission electron microscopy. This technique helps visualize the morphology of exosomes.

Source: Jung, M. K. et al. Sample Preparation and Imaging of Exosomes by Transmission Electron Microscopy. J. Vis. Exp. (2018)
This video describes a ...

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

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