Negative Staining
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.
Methylamine Tungstate
Make up a 4 % solution in water. It’s difficult to dissolve so you need to vortex.
Centrifuge before use to pellet insoluble particulates.
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 µl) 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
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.
Negative staining
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.
Other Negative Staining procedures
Techniques for the preparation of negative stain specimens are simple and direct. The
essential aim of the procedure is to embed the specimen in a uniformly thin deposit of
stain. Resolution of molecular features is only accomplished at the stain-specimen
boundary where there is maximum contrast. This result is only achieved if the deposition
of buffer salts or other materials with densities less than the stain at that boundary are
severely limited; otherwise the specimen molecules will be imaged at low resolution and
will appear as nondescript blobs.
The specimen sample is usually applied directly to the surface of the support film where a
population of specimen particles becomes adsorbed. Attachment of the molecules is
usually secure enough that they are not removed by subsequent rinsing and staining
operations which do remove most of the buffer salts. Since different specimens often
have different affinities for the particular support film being used, some measure of
control over how much specimen attaches to the film may be effected by adjustment of
the specimen and buffer concentrations, adsorption time, etc. Appropriate conditions
must be established by experiment for each new specimen. For protein solutions, typical
concentrations range between 50-500 µg/ml and adsorption times from as little as 1-5
seconds to several minutes. Stains are usually applied in a range of concentration from
0.25-4.0%. Adjustment of stain concentration provides some control over the thickness of
the deposit.
It does NOT follow that a procedure that is successful with one type of specimen is also
suitable for another, so various modifications should always be tried until good contrast
and spreading conditions are achieved.
There are two common procedures for preparing negatively-stained specimens on EM
grids:
Adhesion (drop) method (Figs. II.42 and II.43)
A droplet of specimen is placed on the surface of the grid support film, making sure it
sufficiently wets the surface. After an appropriate time interval, excess specimen is
wicked away by touching a piece of filter paper to the edge of the grid surface. Without
letting the grid dry, a droplet of rinse or stain solution is applied to the grid. Rinsing is
necessary if the specimen preparation contains high concentrations of buffer salts or other
solutes which may interfere with deposition of stain. The nature of an appropriate rinse
depends on the conditions that the specimen can tolerate. Many viruses, for example, can
withstand rinsing with distilled water. In some instances the stain solution can itself act as
a suitable rinse. After rinsing and staining, excess fluid is wicked from the grid, leaving a
thin aqueous film on the surface which is left to dry, usually in air.
Fig. II.42. Preparation of a specimen
from particles in aqueous
suspension. (From Hall, p.290)
Fig. II.43. Washing a specimen.
(From Hall, p.290)
The specimen can also be applied to the support film by floating the grid on top of a
droplet of the specimen solution. The grid is then transferred to droplets of rinse and stain
solutions and then dried as before.
An additional variation of the usual adhesion method is to apply the sample to a holey
support film in the same way as is done on regular films. The sample dries in a thin layer
of stain stretched out over the holes, thereby giving maximum contrast since there is no
plastic and/or carbon support. Also, the stain tends to be more evenly distributed around
the particle although the particle often undergoes distortions (shrinkage and flattening)
due to the surface tension forces created as the layer of stain dries. The stain layer also
has a tendency to break either before or after it is exposed to the electron beam. The layer
of stain can be stabilized with a thin layer of evaporated carbon. Another advantage of
this technique over the usual method is that, if small enough, the specimen particles will
be randomly oriented in the stain layer. On regular support films, particles often settle on
the surface of the film in one or a few preferred orientations, thus limiting the possible
views of the specimen.
Spray droplet technique(Fig. II.44)
The normal adhesion method of preparing a particulate suspension may lead to erroneous
conclusions about the relative proportion of particles since different particles are likely to
have different affinities for the substrate. Also the microscopist may select fields
attractive to the eye but which are not representative. The only reliable way of preparing
specimens without introducing a bias is to dry a drop of the original sample in its entirety.
Non-volatile salts and buffers must be removed by centrifugation and washing or by
dialysis so they don't obscure the particles under study or alter the structure when the salt
concentrates in the last stages of drying. The entire residue from the drop must be
examined, thus it is necessary to obtain very small drops. The suspension is atomized as a
fine mist and the droplets are allowed to impinge on the substrate.
Fig. II.44 A simple hand-held nebulizer
from which the sample (s) is sprayed
onto mica (m). (From Willison and
Rowe, p.69)
The spray droplet technique is particularly useful for examining specimens that adsorb so
poorly to the support film that application and removal of rinsing and stain solutions also
removes the specimen. Appropriate volumes of the specimen and stain solutions are
mixed and sprayed in small droplets onto a wetable support surface. If the solution itself
has the propensity to wet and spread over the surface, uniformly thin deposits of
negatively-stained specimen result. The resultant aqueous film will be of uniform depth,
and the mass of stain deposited per unit area of support film tends to be constant. When
the specimen appears to have dried, some water may still be present in the stain bed, and
a rearrangement of the stain deposit could result from its rapid vaporization if the
specimen is suddenly placed in the vacuum of the microscope. Thus, the specimen is
usually allowed to dry (sometimes over a desiccant) for at least 10 minutes.
Drying of the aqueous film proceeds from the edges, the central area covered by the
droplet being last to dry. Minor solutes tend to be held in solution until the last stages of
drying and are deposited in highest concentrations in the central area. As a result, the
specimen in this area ordinarily is of inferior quality.
Fig. II.45. Drop pattern. The small particles
are tomato bushy stunt virus, and the large
spheres are polystyrene latex particles
2,600 Å in diameter. The wedge-shaped
sector has been magnified and
superimposed. (From Hall, p.359)
Note that, using the adhesion drop method, there may be preferential adherence of
particles so relative particle distribution counts cannot be made. A major advantage of the
spray technique over the adhesion method is that preferential adherence to the grid of one
type of particle over another type of particle in a mixture cannot occur. Thus, this is the
method of choice in quantitative studies where relative concentrations of particles in the
sample need to be determined. By knowing the volume of the original drop (from adding
known concentrations of polystyrene spheres) a count of the number of particles in a drop
pattern provides immediately the number of particles per unit volume (Fig. II.45: Note
that this figure shows a metal-shadowed specimen). This, together with the mass per unit
volume obtained by weighing the dried residue from a measured volume can be used to
calculate a value for the mean molecular weight of the particles.
High resolution negative staining
(From Valentine et al, 1968. Biochemistry 7:2143-52)
Rationale: For the highest resolution with negative staining, there should be little or no
support film, but some support is necessary to hold the protein. In this method the
proteins are supported by a carbon film and "embedded" in a film of uranyl acetate. The
film is cast on mica, which provides the cleanest possible surface for the carbon.
1. Freshly cleave a piece of mica, coat with a carbon film using the vacuum
evaporator.
2. Put ~30 mg/ml protein solution in a small vessel.
3. Cut the mica to 3-4 mm2 pieces. Hold a piece with forceps and push into the
solution of protein at a 30-45 degree angle. Do not let the film detach completely
from the mica.
4. Let the film sit on the protein solution for 20-40 seconds.
5. Pull the mica back and allow the film to sit on the mica.
6. Then slide the mica and film onto a solution of 1-2% uranyl acetate or uranyl
sulfate. Pick up the film with a copper grid, dry, and examine in the electron
microscope.
Variations: A more stable film can be obtained if you pick up the film on a grid coated
with a holey formvar film.
A wrinkled carbon film may be better than a smooth one, as it seems to keep pools of
uranyl acetate around the protein better than smooth films.
Try varying the protein concentration for optimal staining. The proteins should be closely
spaced to collect the stain but if they are too crowded proteins will be overlapped and will
not be easily resolved.
The carbon support film should be as thin as possible (3-10 nm) and should be prepared
on freshly cleaved mica. Once the carbon coated mica has been prepared it can be stored
in a dessicator almost indefinitely. Thicker support films will lower image contrast.
Stain solution should be removed from below the surface of a stock stain solution to
avoid contaminants from precipitates that may exist at the surface or the bottom of the
stock solution. The precipitates do not affect the overall quality of the staining procedure
but will result in local heating of the support film when exposed to the electron beam,
which in its most benign form will result in drift, and at its worst will result in a ripping
and curling of the support film.
The carbon support film should only be exposed to proteins that exist in the bulk solution
and must not be exposed to denatured material that is usually present in the meniscus.
The greatest potential source of contamination and the one most often ignored is the
contamination that arises from the tweezers and the support grid. At no time should the
tweezers or the support grid be dipped into either the protein solution or the negative
staining solution.
Procedures for the Preparation of Carbon Support
Films
Norm Olson
Last Updated March 2000
A. Preparation of normal carbon support films
1. Copper grids should be pre-cleaned by sonicating for 10 sec. in acetone followed by 10
sec. of sonication in ethyl alcohol. Allow grids to dry on filter paper in a dust-free
environment before use.
2. Add 0.12g of Formvar powder to 50 ml of ethylene dichloride and mix well on a
magnetic stirrer until dissolved. Pour the solution into a clean coplin jar.
3. Clean a glass slide with water and detergent. Rinse well to make sure that all of the
detergent is removed and finally rinse in de-ionized water before drying with a paper
towel. Blow off any lint on the slide with compressed air. Place the slide in a dry, dustfree environment such as on filter paper under an upturned beaker. If there are problems
in getting the plastic film to be released from the slide (Step 5), using a slide that has not
been as thoroughly cleaned might help.
4. Dip the cleaned slide into the Formvar solution (Fig. 1-1) and touch edge to filter paper
to drain off the excess fluid (Fig 1-2). Dry upright in a dust-free environment (this
requires 5 to 10 min.).
5. Score the edges of the Formvar film with an acetone-cleaned razor blade (Fig. 1-3).
Breathe on the slide to loosen the film, and slowly slide off onto a clean water surface by
immersing the slide into the water at a ~15° angle (Fig. 1-4). Place grids, dull/rough
surface down, onto good (uniform, gray color, un-wrinkled, etc.) areas of the film. Place
a small piece of clean, white office paper onto the surface of the grids and film and allow
the paper to soak up water. Pick up the paper, grids and film and place in a covered petri
dish to dry.
6. Carbon coat film according to directions (see Sec.C) to desired thickness (A lightbrown color indicates a thickness of <100Å.).
7. Place the paper and coated grids onto a piece of filter paper that is soaked with
ethylene dichloride in a covered petri dish. One half hour should be sufficient time to
dissolve the Formvar film and not damage the carbon support. Remove the grids and
paper and allow them to dry in a dust-free area.
B. Preparation of perforated carbon support films
1. Copper grids should be pre-cleaned by sonicating for 10 sec. in acetone followed by 10
sec. of sonication in ethyl alcohol. Allow grids to dry on filter paper in a dust-free
environment before use.
2. Add 0.17 g of Formvar powder to 50 ml of chloroform and mix well on a magnetic
stirrer until dissolved. Pour the solution into a clean coplin jar.
3. Clean a glass slide with water and detergent. Rinse well to make sure that all of the
detergent is removed and finally rinse in de-ionized water before drying with a paper
towel. Blow off any lint on the slide with compressed air. Place the slide in a dry, dustfree environment such as on filter paper under an upturned beaker. If there are problems
in getting the plastic film to be released from the slide (Step 6), using a slide that has not
been as thoroughly cleaned might help.
4. Add about 50 drops of a 50% glycerol/water solution to the surface of the Formvar
solution. Place the tip of a probe sonicator onto the surface of the solution and sonicate
until mixed. Sonication intensity should be great enough to "violently" cause the solution
to bubble. This often requires not much more than about 5 seconds. This should produce
numerous holes that are 1-2 µm in diameter and suitable for use with frozen-hydrated
samples. Sonicating for longer periods of time produces smaller holes in the film.
5. Immediately after sonicating, dip the cleaned slide into the Formvar solution (Fig. 1-1)
and touch edge to filter paper to drain off the excess fluid (Fig 1-2.). Dry upright in a
dust-free environment for about 5 to 10 min.
6. Score the edges of the Formvar film with an acetone-cleaned
razor blade (Fig. 1-3). Breathe on the slide to loosen the film,
and slowly slide off onto a clean water surface by immersing
the slide into the water at a ~15° angle (Fig. 1-4). Place grids,
dull/rough surface down, onto good (uniform, gray color,
unwrinkled, etc.) areas of the film. Place a small piece of clean,
white office paper onto the surface of the grids and film and
allow the paper to soak up water. Pick up the paper, grids and
film and place in a covered petri dish to dry.
7. Place the paper with the film and grids onto a methanol-soaked piece of filter paper in
a covered petri dish for about 30 minutes. This should perforate any pseudo-holes (these
occur when a small drop of glycerol was present but it was not enough to perforate the
film) that may be in the films. After allowing the paper and film to dry, the grids may be
examined in a light microscope under phase contrast to determine the quality of the films.
8. Carbon coat film according to directions (see Sec.C) to desired thickness (A lightbrown color indicates a thickness of < 100Å.).
9. Place the paper and coated grids onto a piece of filter paper that is soaked with
ethylene dichloride in a covered petri dish. One half hour should be sufficient time to
dissolve the Formvar film and not damage the carbon support. Remove the grids and
paper and allow them to dry in a dust-free area.
C. Use of the Ladd shadow evaporator for carbon coating plastic films
1. Turn shadow evaporator on: Turn both the main and mechanical pump switches on.
Move the black-knobbed, manifold valve handle downwards to "backing" position. Open
the air inlet valve and CAREFULLY remove the implosion shield and bell jar. Set the
bell jar upside down on the rest on the adjacent cabinet
2. Set up carbon coating apparatus: Plug one lead to ground ("E") and the other to "1"
(See lower diagram). Remove the cylindrical glass shield. Release the tension spring that
holds the right carbon rod in place and remove the rod. File the edge of the left carbon
rod flat with a piece of emery cloth. Replace the right rod with a fresh one or sharpen it
by the procedure described below.
3. Carbon rod sharpening procedure: Place the carbon rod in the chuck of the
sharpener. Pull the rod out until its edge is aligned with the edge of the aligning arm and
then tighten the chuck. Turn on the sharpener and run the first sharpener tool against the
rod until a conical point is formed. Then run the other sharpener tool against the rod until
a narrow point is formed. Turn off the sharpener and clean off all carbon dust. Put the
newly sharpened rod in the chuck of the carbon coater and tighten. Replace the tension
spring and then the glass shield.
4. Set up grids: Place the grids and paper support on a piece of filter paper on top of the
base of the carbon coating apparatus (See diagram below). Place a thumbtack along side
the slide. This provides a "shadow" on the filter paper and helps you determine the
relative thickness of the carbon coating.
5. Diffusion pump warm up: Replace the bell jar and the implosion shield. Close the air
inlet, and move the manifold valve handle slowly upwards to the roughing position.
Allow the vacuum to reach 0.04 Torr on the bell jar gauge and then move the handle
downwards to backing. IMPORTANT: Turn on the water supply. The water supply-line
valve is located on the wall behind the shadow evaporator. Turn on the diffusion pump
switch and allow the pump to warm up for 15 minutes before continuing.
6. Obtaining a high vacuum: Move the manifold valve handle slowly upwards to the
roughing position and allow the vacuum to reach 0.04 Torr on the bell jar gauge. While
waiting for the vacuum to recover, fill the baffle with liquid nitrogen. When the bell jar
vacuum has reached 0.04 Torr, move manifold valve handle down to the backing
position. Depress the metal guard beneath the red mains valve knob and move the knob
handle upwards to the open position Allow the vacuum to reach a minimum of 2x10-5
Torr or better.
7. Carbon coating: Turn the electrode selector to #1. Turn the electrode switch on.
Slowly turn the electrode current control knob until there is a slight glow at the point
where the two carbon rods meet. Slowly increase the current until the rods become white
hot. The proper current setting should be just before the point where the carbon starts to
sputter. Frequently monitor the thickness of the carbon by turning down the current,
checking the darkening of the filter paper and then turning the current back up again.
8. Diffusion pump cool down: Turn down the electrode current control knob and turn
off the electrode switch. Make sure the manifold valve is set to the backing position and
close the mains valve. Open the air inlet, remove the implosion shield and bell jar and
remove the grids. Then replace the bell jar and implosion shield, close the air inlet and
move the manifold valve handle to the roughing position. Allow the vacuum to reach
0.04 Torr on the bell jar gauge, move the manifold valve handle to the backing position,
turn off the diffusion pump, and allow the pump to cool for 20
minutes.
9. Turn Shadow Evaporator off: Close the manifold and turn
off the mechanical pump and main power switches. Turn off the
cooling water.
D. Glow discharging carbon films
1. Note: Place the very edge of your carbon coated grids along the edge of a piece of
double-sided tape on a glass slide. This will help to prevent your grids from flying around
inside the shadow evaporator when the air release switch is opened.
2. Turn shadow evaporator on: Turn the main power switch on, turn on the mechanical
pump and move the manifold valve handle (black knob) downwards to the backing
position. Open the air inlet. CAREFULLY remove the implosion shield and bell jar.
3. Set up glow discharge unit: Plug the lead into the proper receptacle (bnc connector).
Place the glass slide with your grids on the unit and replace the bell jar and implosion
shield. Close the air inlet, turn the butterfly switch by the current gauges to glow
discharge and move the manifold valve handle slowly upwards to the roughing position.
Allow the vacuum to reach 0.2-0.15 Torr on the bell jar gauge. The manifold valve may
be turned to the closed position if the vacuum rises above 0.10 Torr.
4. Glow discharging: Turn the electrode selector to position #1 and turn the electrode
switch on. Slowly turn up the electrode current until there is a bright purple glow
surrounding the glow discharge unit. Maintain this setting for approximately 10 seconds
while monitoring vacuum. Turn off the electrode current control knob and the electrode
switch. Move the manifold valve handle to the backing position. Turn the butterfly switch
back to the evaporator setting.
5. Turn shadow evaporator off: Slowly open the air inlet to prevent your grids from
being blown around the bell jar. Remove the grids, replace the shields and then close the
air inlet. Move the manifold valve to the roughing position. Allow the vacuum to reach
0.04 Torr on the bell jar gauge before moving the manifold valve handle to the horizontal
(closed) position. Turn off the mechanical pump and the main switch.
Prepared by Ricardo A. Bernal