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.

Transcript

The overall goal of this video is to demonstrate several different variations on techniques for preparing negative stain transmission electron microscopy samples. Negative stain is the best way for rapidly assessing the quality of an EM sample. It's important to recognize that different grid preparation techniques work better for some samples.

The technique which works best for any given sample must be determined by trial and error. To begin, prepare the EM grids using the carbon sheet method and prepare the negative staining reagents as described in the accompanying text protocol. Place a carbon sheet coated EM grid facing up on a microscope slide.

Set the slide into a glow discharge unit and treat the grid for a minimum of 30 seconds at 10 milliamps to render the grid hydrophilic. When finished, remove the sample from the discharge unit. Then, use a pair of negative pressure tweezers to grip the edge of the grid.

To stain using the side blot method, first apply three to five microliters of the sample to the support surface. Allow the sample to adsorb to the grid surface for 10 seconds to one minute, then touch the edge of the grid to a sheet of filter paper and allow capillary action to pull off the liquid. Next, place 50 microliter drops of ultrapure water or an 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. Then, touch the edge of the grid to a sheet of filter paper and allow capillary action to pull off the liquid. Next, place two 50 microliter drops of staining reagent 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 top surface of the grid. If the stain migrates to the back of the grid, then the carbon film has been broken and it must be discarded. After 10 to 15 seconds, touch the edge of the grid to a sheet of filter paper and allow capillary action to draw off the liquid.

Repeat this staining step one more time and then allow the grid to air dry. To stain the sample using the flicking method, grip the edge of the grid with a pair of negative pressure tweezers and apply three to five microliters of sample to the support surface. Holding the tweezers in one hand, angle the grid so that it is at approximately 45 degrees facing away and rapidly flick the wrist to flick off the majority of the droplet that is on top of the grid.

Then, use a glass Pasteur pipette to apply a drop of wash solution to the support surface. Flick off the drop and repeat a few times to wash the sample. Next, apply a drop of staining reagent to the support surface and again flick off the droplet.

Repeat this staining step one to three times, depending on the stain depth that is required for visualization of the specimen. Remove excess stain by touching the torn edge of a piece of filter paper to the edge of the grid and then allow the grid to air dry. To stain the sample using the rapid flushing method, first draw 30 to 70 microliters of stain into the tip of a 200 microliter pipette.

Turn the volume dial to draw up five microliters of air then draw up five to 30 microliters of the wash reagent, followed by another small air gap, before another five microliters of sample. Grip the edge of a grid with a pair of negative pressure tweezers and angle the grid approximately 45 degrees facing away. In one swift motion, eject the entire contents of the pipette tip across the face of the carbon coated EM grid.

Remove excess stain by touching the torn edge of a piece of filter paper to the edge of the grid and allow the grid to air dry. The ideal choice for a negative stain is highly sample dependent. The most commonly used stain is uranyl acetate, but lanthanide stains can also be used.

For deeply embedded samples, lanthanide-based stains, thulium acetate and erbium acetate, produced negative staining of equivalent quality to uranyl acetate, as judged by the apparent contrast and sharpness of the stained particles, with thulium acetate producing clearer and crisper images than erbium acetate. The large grain size of thulium acetate becomes apparent at high magnification, shown here when a sample of tobacco mosaic fibrous particles were stained with 1%thulium acetate. Using thulium acetate, the 23 angstrom repeat of the tobacco mosaic fibrous particle is clearly visible by eye.

None of the other lanthanide stains tested were able to resolve this feature. Another difficult sample to image is the muscle-derived C protein. This sample produces significantly different images depending on the staining method.

When stained with a side blot method, it appears as a globular, collapsed wing-like structure. When grids are prepared by the flicking method, the C protein is observed as a series of domains that resemble beads on a string. This better represents the actual C protein structure.

Once mastered, it only takes about a minute to prepare a negatively stained sample. It is important to remember that each sample responds differently to each staining technique. For new samples, the best approach must be determined experimentally.

Don't forget that uranyl staining reagents are toxic and slightly radioactive. Make sure they're disposed of in a proper waste stream.

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Journal (Biochemistry)

Journal (Biochemistry)

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

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