Method Article
An erratum was issued for: Variations on Negative Stain Electron Microscopy Methods: Tools for Tackling Challenging Systems. An author name was updated.
One of the authors' names was corrected from:
Matthew G. Iadaza
to:
Matthew G. Iadanza
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 contrast enhancing stain reagent. Although limited in resolution to a maximum of ~18 - 20 Å, 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.
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' 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's resolution being limited to ~18 Å 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 'halo' 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 Å)9 and provide higher resolution images over other stains such as phospho-tungstates (8 - 9 Å 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.
1. Preparation of EM Grids
2. Preparation of Negative Staining Reagents
3. Adsorbing Samples to the Carbon Substrate and Staining
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 Å 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.
| Reagent | Concentration | pH | Type |
| Ammonium Molybdate | 1 - 2 % | 5 – 7 | Anionic |
| Erbium acetate (ErAc) | 1 – 2% | 6 | Cationic |
| Gadolinium Acetate (GdAc) | 1 – 2% | 6 | Cationic |
| Methylamine tungstate | 2% | 6 – 7 | Anionic |
| Samarium acetate (SmAC) | 1% | 6 | Cationic |
| Sodium silicotungstate | 1 – 5 % | 5 – 8 | Anionic |
| Sodium phosphotungstate | 1 -3 % | 5 – 8 | Anionic |
| Thulium Acetate (TmAc) | 1 – 2% | 6 | Cationic |
| Uranyl Acetate (UA) | 1 – 3% | 3 – 4 | Cationic |
| Uranyl Formate (UF) | 0.75 – 1% | 3 – 4 | Cationic |
Table 1: Some common negative staining reagents.
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. Please click here to view a larger version of this figure.
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 Å. Please click here to view a larger version of this figure.
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. Please click here to view a larger version of this figure.
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.
The authors declare no competing financial interests.
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's and Stephen Muench'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.
| Name | Company | Catalog Number | Comments |
|---|---|---|---|
| 200 mesh copper EM grids | Sigma-Aldrich | G4776-1VL | Other materials and/or mesh sizes can also be used |
| Ammonium Molybdate | Sigma-Aldrich | 277908 | |
| Carbon evaporator | Ted Pella Inc. | 9620 | Cressington 208 or equivalent |
| Collodion solution 2% in amyl acetate | Sigma-Aldrich | 9817 | |
| Dumont #5 negative pressure tweezers | World Precision Instruments | 501202 | Or other tweezers as preferred |
| Erbium Acetate | Sigma-Aldrich | 325570 | |
| Gadolinium Acetate | Sigma-Aldrich | 325678 | |
| Mica Sheets. 75x25x0.15mm. | AGAR Scientific | AGG250-1 | |
| Microscope slides, white frosted | Fisher Scientific | 12607976 | Or equivalent |
| Parafilm | Fisher Scientific | 10018130 | Or equivalent |
| Pasteur pipette (glass) | Fisher Scientific | 10343663 | Or equivalent |
| Razor blade | Fisher Scientific | 11904325 | Or equivalent |
| Sandpaper | Hardware store | Wet and dry sandpaper with grit finer that 200 (600 suggested) | |
| Samarium Acetate | Sigma-Aldrich | 325872 | |
| Sodium Hydroxide | Sigma-Aldrich | 1.06462 | |
| Sodium Phosphotungstate | Sigma-Aldrich | P6395 | |
| Stainless Steel Mesh, 150x150 mm (cut to size). | AGAR Scientific | AGG252 | |
| Thulium Acetate | Sigma-Aldrich | 367702 | |
| Two Step Carbon Rod Sharper, for 1/4" rods | Ted Pella Inc. | 57-10 | Or equivalent for carbon evaporator used |
| Ultra pure water | |||
| Uranyl Acetate | Electron Microscopy Sciences | 22400 | |
| Uranyl Formate | Electron Microscopy Sciences | 22450 | |
| Vacuum grease | Fisher Scientific | 12719406 | Or equivalent |
| Whatman #1 Filter paper. | Fisher Scientific | 1001 090 | Or equivalent |
| Whatman #40 filter paper | Fisher Scientific | 10674122 | Or equivalent |
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