Negative Staining for Transmission Electron Microscopy

 

Introduction

Specimens that are to be examined in the transmission electron microscope (TEM) usually have to be thin, dry and contain contrast (usually from a heavy metal stain. One of the easiest ways of preparing biological samples, of small size, for the TEM is by negative staining. This preparation method is useful for visualizing suspensions of small particles, which includes viruses, purified proteins, liposomes and small vesicle fractions.

Negative staining is a simple technique for routine examination of structure. It does not allow for high resolution examination of samples - more technically demanding methods, using sample vitrification (or rapid freezing) are used for this.

As with most techniques, there are levels of complexity which yeild increasingly better results. The major differences between the techniques is in the choice of sample support.

 

Staining samples on formvar-carbon coated grids

The sample is suspended (or diluted) into water (if possible) and adsorbed onto a carbon-coated formvar film which is attached to a metal specimen grid. The carbon surface of the grid becomes contaminated when stored, and thus hydrophobic, so it is best to glow discharge the grid surface (making it hydrophilic) prior to use. This is usually done in a vacuum chamber of a vacuum evaporator.

Once the specimen has been adsorbed onto the film surface, the excess sample is blotted off and the grid is covered with a small drop (5 ?l) of stain solution (see below). This is left on the grid for a few minutes and then blotted off. The sample is dried and examined in the TEM.

If carbon-formvar films are prepared on specimen grids with small mesh size, the films, with adsorbed sample, can usually be immunolabeled without them breaking. This protocol is described in more detail in the section on immunolabeling. Briefly, the grids are floated onto drops of diluted antibody. washed by floating on drops of buffer, and then floated on drops of diluted visualization probe. We normally use colloidal gold coupled to staphylococcal protein A (protein A-gold) or other affinity markers such as antibodies.

 

Methods using carbon films alone

I. Carbon-coated grids

The grids are prepared by first preparing formvar-carbon coated grids and then removing the formvar support. This is done by placing the grids in an atmosphere of solvent vapour, which dissolves the formvar (which is a plastic). The grids are placed on a wire mesh in a glass perti dish, the solvent (chloroform or carbon tetrachloride) is placed in the dish below the wire mesh and the dish is closed by replacing the lid. If the vapour alone does not remove the film (which should take a few hours), the process can be accelerated by dipping the coated grids into the solvent prior to closing the dish and placing them on the mesh to soak. Allow the solvent to completely evaporate before removing your grids, which will now have only carbon on them. Remember that the carbon must be thick enough to be self supporting. Advantages include high stability in the electron beam and high resolution examination of the adsorbed specimen. Disadvantages include a hydrophobic surface which is not of uniform thickness.

II. Carbon films

Negative staining

• For this technique, carbon film is evaporated onto a freshly cleaved mica surface and the sample is applied to the surface of the carbon film attached to the mica. The carbon film and the sample is then picked up onto a specimen grid and examined in the TEM. The carbon films can be stored on the mica for long periods without them becoming contaminated, the films are thin but tough so can be easily examined in the TEM without any noticable specimen movement. However, they are fragile and must be manipulated with care. They are more stable if supported on specimen grids with small mesh size (e.g. 400 mesh). Hexagonal grids seem to offer a more stable support.

  
  Making the carbon film

  Take a piece of mica and cut off a square or rectangular piece that is approximately 2 x 3 cm. Cleave the mica with a razor blade or scalpel that has been cleaned with acetone. Once the mica has begun to separate along a cleavage plane, forceps may be used to pull the mica completely apart. It is better to do this than to scratch the surface of the mica with the razor blade. Mica, as purchased from most EM supply companies, is rather thick and a new piece may be split around 4 times. Attach the mica to a filter paper, with sticky tape, with the freshly cleaved plane facing upward. Place the filter paper in a vacuum evaporator for carbon coating and deposit a film of carbon onto the mica surface. This mica should be cleaved immediately before coating as the freshly cleaved surface is clean and hydrophilic, but it becomes contaminated (and thus hydrophobic) over time time. After coating, the filter paper should be light gray in color (compare the filter paper behind the mica with that which was exposed, and therefore carbon coated). This is a good indicator of the film thickness.

  Prepare the sample (the staining works best if the sample is in water) by making several serial dilutions of the preparation. The best negative stain preparations are those that have a single layer of individual, separated particles adsorbed onto the film. This is achieved by dilution of the sample.

  With a pair of scissors, cut off a small square of mica, about 4 x 5 mm, about twice the size of an EM grid. Put the square on a piece of parafilm, carbon side up. With a pipetman, gently squirt 5 - 10 ml of the sample under the carbon by placing the pipette tip to the side of the mica square. The sample will do one of two things: 1) It will flow between the carbon and mica, in which case you will need only a small amount of sample. 2) It will flow between the mica and the parafilm, in which case just keep applying sample to the side of the square until it goes under the carbon. Place a 400 mesh specimen grid onto the carbon. (Wash the grid in 0.5% acetic acid and then acetone prior to use) Break the carbon film to free the specimen grid, lift the grid and place it on a drop of stain solution for about 30 seconds (sample side down). Blot dry, and examine in the TEM when completely dry. Staining and washing times can be varied. A better support for the carbon film can be obtained by using specimen grids coated with a holey film of formvar. This is a film containing lots of small holes which are covered by the carbon film. There are many ways to produce these films but one quick, simple way is to breath onto a film of formvar before it dries onto the glass slide. The moisture droplets will displace the film, leaving small holes. Althernatively, the wet film on the glass slide can be placed in a cloud of steam.
  Adenovirus particles

• 

  An example of negative staining. These adenovirus particles have been adsorbed onto a carbon film that was deposited onto a freshly cleaved mica surface. The film was picked up onto a clean, 200 mesh specimen grid coated with a holey formvar film. The preparation was stained for 1 minute with neutral 1% aqueous phosphotungstic acid and photographed in a transmission electron micrograph.
  Negative stain solutions

  Aqueous Uranyl Acetate

  A 1% to 3% solution of uranyl acetate dissolved in water can be used to negatively stain many samples. The stain has a low pH so this solution is not recommended for particles that are unstable in acid conditions.

  Neutral Phosphotungstic Acid

  A 1% to 3% solution of phosphotungstic acid is made up in water and the pH is adusted to 7 using sodium hydroxide. This is a useful stain for many samples but is especially good for viruses that dissociate at low pH. The stain produces less contrast than the uranyl acetate.

  Ammonium Molybdate

  Make up a 1% solution of ammonium molybdate in water. This solution has also been used to negatively stain thawed, thin cryosections of fixed cells.