Electron Microscopy of Extracted
Cytoskeletons: Negative Staining,
Cryoelectron Microscopy, and
Correlation with Light Microscopy
Our understanding of the function of the cytoskeleton
and its associated proteins has been much
advanced by developments in imaging methods for
light microscopy. With the advent of probes, such as
green fluorescent protein, the localisation of gene
products in living cells under different physiological
conditions is now readily feasible (e.g., Vignal and
Resch, 2003). However, to extend our understanding
of the functional interactions taking place, parallel
information about cellular ultrastructure is essential.
Here, electron microscopy (EM) is required to provide
information about cell architecture at the nanometre
Three major filament systems make up the
cytoskeleton: actin filaments, microtubules, and intermediate
filaments. When not in organised bundles,
actin filaments are poorly preserved after dehydration
and embedding in plastic. This fact, taken together
with the open, three-dimensional nature of the
cytoskeleton networks, has limited the information
obtainable by conventional EM methods based on thin
sectioning. The methods of choice for studying the
cytoskeleton of cultured cells have therefore centered
on "whole mount" procedures, involving the culturing
of cells on plastic films or coverslips and the preparation
of entire cytoskeletons, or their replicas, for
electron microscopy. Methods that deliver useful
structural information about cytoskeleton organisation
include quick freezing and deep etching (Heuser and
Kirschner, 1980), critical point drying followed by
metal shadowing (Svitkina et al.
, 1995), and negative
staining (reviewed in Small, 1988; Small et al.
More recently, advances in cryoelectron microscopy
have opened the way to the study of cytoskeletal structures
under conditions that obviate processing steps
that could produce artefactual distortions of filament
networks (Resch et al.
, 2002; Medalia et al.
, 2002). In
particular, these latter approaches, involving cryoelectron
tomography, promise to deliver new information
about ultrastructural interactions in three-dimensional
space unobtainable by other means.
This article describes the application of negative
staining and cryo-EM to studies of the cytoskeleton;
taking advantage of the developments in light
microscopy, we also highlight the feasibility of correlating
information on the living cell with electron
microscopy (see also Svitkina and Borisy, 1999) for
delivering meaningful structural information.
II. MATERIALS AND
Chloroform (Cat. No. 1.02445.1000), disodium
hydrogen phosphate (Cat. No. 1.06580.1000), ethanol
(Cat. No. 1.00983.2511), glucose (Cat. No. 1.04074.1000), sodium chloride (Cat. No. 1.06404.1000),
and sodium dihydrogen phosphate (Cat. No.
1.06345.1000) are from Merck EGTA (Cat. No. E3889),
MES (Cat. No. M2933), taxol ("paclitaxel," Cat. No.
T7402), and Triton X-100 (Cat. No. X100) are from
Sigma. Phalloidin was a generous gift from Professor
H. Faulstich (Heidelberg) and is obtainable from
Sigma (Cat. No. P2141). EM-grade glutaraldehyde
(Cat. No. R1020) and Formvar (Cat. No. R1201) are
from Agar Scientific, Stanstead, UK; nickel 200 mesh
hexagonal or nickel H2 finder grids are from Graticules
Ltd., Tonbridge, UK; and steel and titanium
forceps (number 5) are from Dumont & Fils., Switzerland.
Thirteen-millimeter-diameter plastic coverslips
(Cat. No. 174950) are from Nunc, Rochester. Further
materials required include Dow Corning siliconebased
high vaccum grease and Whatman Qualitative
No. 1 filter paper. The following are from local suppliers:
glassware (50-ml bottle, measuring cylinder, a
glass through minimum 7cm deep, a rod or pipette),
object slides with frosted ends, Parafilm, transfer
pipets, scalpels, and blades. Additionally, cell culture
equipment (media, incubators, inverted phase contrast
microscope), a basic workshop setup (for producing
coverslips with holes), and a routine transmission electron
microscope are required.
B. Negative Staining
Sodium silicotungstate (Cat. No. R1230) is from
Agar Scientific, Stanstead, UK. Minisart 0.2-µm filters
are from Sartorius.
1,2-Dichlorethane (Cat. No. 8.22346.2500) and
methanol (Cat. No. 1.06009.2511) are from Merck, and
87% glycerol (Cat. No. G7757) is from Sigma. Additional
laboratory equipment required for producing
holey films is a microprobe sonicator and a
vacuum evaporator for carbon coating (e.g., Edwards
A freeze-plunging device, either home made (drawings
and materials list available upon request) or a
commercial apparatus housed in a chemical extractor
hood, is needed. At the base of the plunger is a polystyrene
box for liquid nitrogen and into this sits a small
container for condensing ethane (see Fig. 3). Frost-free
liquid nitrogen, gaseous ethane, and a regulator valve
with a length of silicon tubing, 4mm inner diameter
and 6 mm outer diameter (Tygon 3603, Norton Performance
Plastics Corporation, Akron, OH), with one end
fitted with a pipette tip with its end cut at a 45° angle, are needed. Note: Remove any sources of ignition from the
working area as ethane is highly flammable and explosive
Safety glasses for eye protection or a Plexiglas full face
shield, which additionally helps prevent inadvertent
breathing onto the cold surfaces, a cause of ice contamination,
Use Whatman qualitative No. 1 filter paper (Cat.
No. 1001055), cut to rectangular strips of approximately
1 × 4cm. Grid box (Agar Scientific, Cat. No.
G276A) cut into small squares of 15 mm so that there
are four grid holes arranged symmetrically within the
square, fitted with a clear plastic lid, 13mm circular
diameter and 1mm thick, with a 4-mm square notch
cut out to allow access to one grid hole at a time is
required. This is fitted with metal or nylon screw
attached through the center to the grid box base so it
can be rotated from one hole to the next and fixed tight
for storage. Obtain 50-ml polypropylene conical tube
with two approximately 8-mm-diameter holes in the
lid to allow nitrogen to enter the tube and for retrieval
using long tweezers. Small, portable industrial quality
vacuum dewars (do not
use houshold vacuum flasks
as they can dangerously implode) or a polystyrene
container for transporting grids in the grid box is
needed. Purchase a large vacuum dewar for long-term
storage; it is useful to have a dewar that has numbered
and colour-coded holders of the correct size to hold
Additional tools needed are large tweezers (200mm
long; Bochem, Weilberg, Germany) and small screwdrivers.
At least two sets of tools that contact liquid
nitrogen should be available, as they will frost after
they have been cooled in the nitrogen and then
removed and left on a bench. If you then reintroduce
them back into the liquid nitrogen, they will contaminate
the nitrogen with the ice crystals.
For electron microscopy, a transmission electron
microscope fitted with cryo blades to protect the frozen
grid from ice contamination, as well as a cryo holder,
cryo transfer station, and temperature controller
(Gatan, Pleasanton, USA), are needed.
A. Negative Staining
1. Preparation of Formvar-Coated Grids
Thin plastic films on grid supports, as used for conventional
EM, can be used as a growth substrate for
cultured cells. For alternative protocols, see Small and
Herzog (1994) and Small and Sechi (1998).
Dissolve 0.4 g Formvar in 50ml chloroform and stir
vigorously in a tightly closed bottle for at least 1 h. To
produce films without holes, this solution and all
glassware used should be kept as dry as possible.
2. Cell Culture
- Clean an object slide with a frosted end first with
distilled water and then with 70% ethanol and allow
to dry completely.
- Transfer the Formvar solution into a measuring
cylinder wide enough for the slide and submerge the
slide, holding it on the frosted end by a clamp or peg
attached to a thread, to just below the frosted area. After
30 s, lift the slide smoothly out of the solution and let it
dry for 3 min hanging just above the solution surface.
- Drain the excess solution at the bottom of the
slide with filter paper and cut the film at the bottom,
the left and the right edges, around 1-2 mm inside the
edge of the slide. (Do the same for the back side before wetting the film, if both films are to be floated off
- Prepare a large vessel filled to the lip with distilled
water, clean the surface with a glass rod from
dust, and float off either one or both of the films simultaneously
at an angle of 45° resp. 90° onto the surface.
A black background below the vessel improves visualisation
for this and the following steps. (Note: If difficulties
are encountered with floating off the film, try
another brand of slides or a different washing protocol;
see also Small and Herzog, 1994.)
- Place 200 mesh Ni grids with the dull side downwards
onto the film. For easy handling of the magnetic
nickel grids, the use of titanium forceps is recommended.
Retrieve the film from the water surface with
a piece of Parafilm that is put on top of the floating
grids/film assembly and then lifted up carefully from
one end. (An alternative way to do this is to use a
microscope slide: with an address sticker adhered to
one side and trimmed at the the edges, hold the slide
at a 45° angle (paper side down), touch the film a few
millimetres within one of the narrow ends, and slowly
submerge the slide into the water. The film with the
grids will follow the slide and stick very well.)
- Remove excess water by blotting carefully with
filter paper and allow the filmed grids to dry completely.
To separate them without damaging the film,
cut the film around each individual grid with the tip
of a pair of forceps.
Cells are plated onto the grids the same as for
coverslips, and the density is chosen to give one or two
cells per grid square after attachment and spreading. In
general, cells spread more slowly on plastic film than
on glass; to encourage attachment and spreading, the
filmed grids may be coated with matrix molecules or
serum (depending on cell type) prior to plating. Check
for adequate cell spreading in an inverted microscope
with phase-contrast optics prior to further processing.
In order to render the film more stable in the electron
beam, an additional layer of carbon deposited
onto the grid can be used. However, under these conditions,
we experience a decrease in quality of the negative
stain so that we usually apply a thin carbon layer after
staining (see later).
To avoid the problem of loose, floating grids, it is an
advantage to immobilise them in one way or another
(see also Small and Sechi, 1998). To immobilise single
grids, we sandwich them between a plastic coverslip
with a 2.5- to 2.7-mm hole in the center of a petri dish.
Dots of vacuum grease are used to fix the plastic
coverslip to the petri dish.
- Prepare plastic coverslips >12mm diameter with
a central hole of 2.5-2.7 mm; this is best done by fixing
a stack of them tightly in a holder prepared separately
for this purpose (Fig. 2a) before drilling. Clean the
coverslips by washing them twice with 70% EtOH and
once with water, both of them heated in a microwave
oven until boiling.
- For cell culture over periods of more than 12h,
sterilise grid sets under UV light in an open petri dish
for 5 rain. Air bubbles get trapped easily in the mesh,
which makes observation of the spreading cells (e.g.,
in phase contast) difficult. This can be avoided by
incubating the grids in sterile water in the fridge
overnight before use. This step should be performed
individually for each grid in grid boxes (volume per
slot approximately 15µl) to avoid the grids adhering
to each other.
- Transfer the grids, with the filmed side up, to
sterile (and dry) petri dishes and fasten them with the
plastic coverslips: Apply small spots of high vacuum
grease onto the edge of the coverslips and press them
down into the petri dishes, with the grid trapped
below the hole. Only coverslips with a regular hole
and fiat edge should be used.
- After they are mounted, the films can be protein
coated and cell culture can be carried out as usual.
- Prewarmed phosphate-buffered Saline (PBS, 150mM NaCl, 3 mM NaH2PO4, 8 mM Na2HPO4, pH 7.4)
- Prewarmed extraction buffer [0.25% Triton X-100
(diluted from a 20% stock) and 0.5% EM-grade glutaraldehyde
(diluted from a 25% stock) in CB: 10
mM MES, 150 mM NaCl, 5 mM EGTA, 5 mM MgCl2,
5 mM glucose, pH 6.1]
- Fixation buffer at room temperature (1.0% EMgrade
glutaraldehyde from a 25% stock in CB)
- Phalloidin, 1mg/ml in MeOH, stored at -20°C
- Taxol, 10 mM in dimethyl sulfoxide, stored at -20°C
- Remove cover glasses and transfer grids to a
small petridish with prewarmed PBS for a brief wash.
To avoid bending in the transfer step, a magnetised
steel forceps is useful for initially lifting the grid so that
it can be grabbed more easily with another pair of (titanium)
- Aspirate the PBS carefully, do not allow to dry,
and replace it with prewarmed extraction buffer; incubate
- Replace the extraction buffer with fixation buffer
and fix for >20min up to overnight at 4°C; to improve
the preservation of actin and microtubules, 10µg/ml
phalloidin or 10µg/ml taxol can be added.
- Negative Staining and Electron Microscopy
|FIGURE 1 Extracted cytoskeletal networks in thin regions
visualised by (a) negative staining with sodium
(b) cryoelectron microscopy, a, actin;
ab, actin bundle; mt, microtubule;
filaments; r, ribosome aggregation. Bar:
2% sodium silicotungstate in ddH2
O, adjusted to
pH 7 with NaOH (equilibration of pH can take a long
time) and filtered through a 0.20-µm filter.
- Clamp the grid in a pair of forceps using a large
paper clip over the shaft to hold the tips together; do
not allow the sample to dry at this step!
- Rinse with a few drops of the negative stain from
a transfer pipette. After a few seconds, drain the stain
from the entire back side of the grid with filter paper
to remove it as completely as possible. On the front,
drain only from the edges to leave a thin film so as not
to damage the sample. Special care has also to be taken
to drain excess stain from between the tips of the
forceps: Bring the edge of the filter paper into contact
with the edge of the grid at the contact point with the
- Allow to air dry and observe in a routine TEM
(Fig. 1a). To avoid drift of the plastic film upon exposure
to the electron beam, a thin layer of carbon can be
deposited onto the film in a vacuum evaporator.
This section describes the method from Resch et al.
(2002) in more detail.
1. Preparation of Holey Films
For unstained specimens, the support film itself
contributes a significant background noise in the electron
microscope image. To improve contrast conditions
in the cryo-EM, we have therefore used holey
films to support cells and have selected areas for structural
analysis where parts of cells span the holes. The
method for making holey films was derived from that
described in Hodgkinson and Steffen (2001); it was
optimised for a hole/film ratio that was significantly
lower than for films used for molecular suspensions to
allow the cells to attach and spread.
Dissolve 0.25 g Formvar in 50 ml chloroform and stir
in a tightly closed bottle for at least 1 h. After the
Formvar is dissolved, add 150µl of 50% glycerol in
water; shake the solution vigorously for 1 min.
|FIGURE 2 (a) Device used for mounting the plastic
drilling consists of a plastic body (1) with an
elongated grove (here
horizontal) and a central round
indentation (2) in which a stack of
cover glasses sit. They
are mounted by a Plexiglas bar (3) on top of
which is pressed down firmly onto them by two screws
at its end (4). Drilling is done via a central hole (5) in the
bar. (b) Perforated carbon film as typically used
for cultivating cells
for cryo-EM on a 200 mesh hexagonal
Ni grid. (c) Two setups to
mount grids for correlative
LM/EM in petri dishes with a coverslip
inset in the bottom
on an inverted microscope; samples are covered
medium, the objective is coming from below. (Left)
setup: The grid with cells up (1) is mounted by a
with a central hole (2) that is attached via
high vaccuum grease (3).
(Right) Inverted setup for
fluorescence microscopy: The grid with
cells on it facing
downwards (1) is put on top of two other nickel
(4) that serve as spacers for the cells and fixed with the
as described earlier.
2. Cell Culture
- Prepare a clean object slide as described
- Sonicate the Formvar/glycerol solution for
- Rapidly transfer the solution to a staining jar, dip
the slide into it, remove, and allow to air dry for a few
- Float off both sides of the film as described earlier
and place 200 mesh Ni grids with the dull side facing
the film on top. In this case, the grids should not
be placed too close together on the film, as the filter
paper used to retrieve the grids will not adhere
- Retrieve the grids and the film with a piece of
filter paper, not with Parafilm or a slide, and allow to
- Prepare a stack of filter paper in a large glass
petri dish saturated with MeOH and incubate the grid
set on this, with the film up, for 10 min. This serves to
perforate the pseudoholes. Remove and allow to air
dry completely before proceeding.
- At this stage, we recommend checking the film
with a phase-contrast microscope for the size and distribution
of holes, which is affected by the amount of
glycerol as well as by the power and duration of the
sonication. If neccessary, vary these factors.
- Coat the grids with a thick (dark grey) layer of
carbon in a high vacuum evaporator.
- Prepare a bed of filter paper saturated with 1,2-
dichlorethane. Place the set of grids, film side up, onto
the filter paper and leave for 2 h to dissolve the plastic
film. If the filter paper bends up at the edges, either cut
it into smaller pieces or use small pieces of glass to
hold it down.
- After final drying, check a few grids in the EM
(see Fig. 2b) and store them individually in grid boxes.
For preparation of cells for cryo-EM, the same
procedure is used as for negative staining. However,
making the cells attach and spread on perforated films
might require additional steps, including coating with
matrix molecules (for general strategies, see also Resch et al.
, 2002; and Vignal and Resch, 2003).
It can save a lot of time and effort prior to preparing
the cryo-frozen grids if they are screened beforehand
in a phase-contrast microscope. This allows the
possibility to roughly assess the quality and number of
cells grown on the surface. The proximity of likely cell
candidates for the electron microscopy can also be
mapped at this stage (see later).
For cryo-EM as outlined here, removal of the membrane
and cytosolic protein is necessary to visualise the
cytoskeletal networks and actin filament subunit structure
clearly. It is also necessary to fix the cytoskeletons
lightly during this extraction process to stabilise the
networks. Medalia et al.
(2002) have applied cryoelectron
tomography to unextracted cells, in which case
filament networks are visible only after complex
Here, the extraction/fixation protocol is the same as
for negative staining, except that postfixation seems
not to be neccessary.
4. Cryo Plunging of Specimen Grids
The objective of plunge freezing is to obtain a well
distributed, quick frozen specimen in a homogeneously
thin layer of vitrified ice uncontaminated with
frost. For this, the blotting regime is critical. The
amount of blotting required can be sample/buffer
dependent so this might have to be adapted for different preparations. It is important to remain consistent
but because in some cases even subsecond differences
in blotting times can cause variability with ice
thickness, differences between grids are unavoidable.
In order to counter this, make several grids at each
If a washing buffer is to be used, it is important that
it is at the same osmolarity as the cell culture media,
as the integrity of the cells could be compromised.
|FIGURE 3 Working setup as typically used for freeze
(1) Gaseous ethane bottle, (2) regulator valve
with Tygon tubing and
pipette tip, (3) full face shield, (4)
filter paper strips, (5) polystyrene
box for liquid nitrogen,
(6) small container (sitting on an aluminum
condensing ethane, (7) plunger arm, fixed with a pair of
forceps holding the grid (arrowhead), (8) 50-ml conical
tube for grid
box storage, (9) tools (fine and large forceps,
screwdrivers), and (10)
sample transport dewar.
details, see text.
5. Electron Microscopy
- Set up your workspace with all the required tools
close at hand prior to freezing (Fig. 3). It is important
to have good light and it is useful to have additional
light (e.g., equipoise lamp) that can be positioned
facing toward the grid for observing the blotting step
when plunging. Frost in the liquid nitrogen and ethane
can be a source of contamination on your grids so
always try to minimise exposure to water vapour.
Always keep the nitrogen containers sealed with lids
and do not breathe onto the nitrogen or ethane surfaces.
Precool all tools that come into contact with the
grid or grid box, as they may heat the specimen,
causing cubic or hexagonal ice to form.
- Set up the plunger. Check the alignment of the
forceps in the plunging arm so that the forceps will
plunge the grid a few millimetres below the surface of
the liquified ethane. Do not adjust the drop height so
that the grid gets too near the base of the ethane container as ethane ice forms there, which will damage
the grid if it comes into contact with it.
- Put on eye protection or a face shield and fill the
polystyrene container with liquid nitrogen. Also fill the
portable transfer dewar with liquid nitrogen and submerge
the polypropylene conical tube into this. Keep
a lid on the transfer dewar to minimise contamination.
- Precool the small ethane container with liquid
nitrogen. Make sure all liquid nitrogen is removed
from the inside of the container as it interferes with the
liquifying of the ethane. Place one of the grid storage
boxes into the liquid nitrogen.
- Place the pipette tip from the ethane supply into
the bottom of the small ethane container and slowly
open the regulator valve. After a short time, the ethane
gas will start to condense on the bottom of the container.
This is normally accompanied by a high-pitched
squeal. Keeping the pipette tip just below the surface
of the liquid ethane and gently moving it around the
inner walls allow the level to rise to the top of the
ethane container. Keep the tip moving in the ethane so
that the tip does not ice up and block. Do not allow the
liquid ethane to spill over into the nitrogen. You can
use a double-walled ethane container with a small gap
between the walls to avoid this. Note: Liquid ethane can
cause serious burns if it comes into contact with your skin
or eyes so always exercise caution and regulate the ethane
to a slow and gentle flow.
- After a short time, the liquified ethane will start
to solidify. If this happens, you must deice by inserting
the pipette tip into the ethane. Slowly open the
ethane gas supply as you do not want to splash ethane
into the nitrogen. Gently move the pipette tip around
the edges of the ethane container until the ethane
- Pick up a grid by the edge, sample side up with
forceps. The samples are usually washed briefly,
immediately prior to plunging. Cut a piece of Parafilm,
place on a flat surface, and apply 2 × 50-µl drops of
washing buffer, e.g., prewarmed PBS. Keeping the grid
in the tweezers, blot the grid side on with a piece of
filter paper. Apply the grid, sample side down to the
surface of the washing droplet, blot, and place onto
the second drop. Remove the grid; a droplet from the
washing buffer should remain on the grid surface.
Place the forceps into the clamp of the plunger arm
with the sample (and droplet) facing towards you.
Using a strip of the Whatman qualitative No. 1
filter paper, carefully blot the surface of the grid. Use
a visual assessment to roughly gauge the degree of
blotting (Fig. 4): When first touching the grid, the filter
paper will pull onto the droplet by surface tension and
you should see the outline image of the grid, through
the paper. Once most of the surface liquid is adsorbed by the paper you will reach a point where the grid
outline will suddenly disappear and the paper will
also pull away. This is the time when the sample
should be plunged rapidly into the ethane. Waiting
even a few seconds after the pull-away point can result
in excess removal of liquid, sample damage, and
|FIGURE 4 Blotting of the grid immediately prior to freeze
(a) when first touching the grid and droplet with
a strip of filter
paper, an outline of the grid is clearly visible
through the paper and
(b) as the liquid is absorbed by the
filter paper, the droplet imprint
will increase in diameter
until most of the liquid is removed from
the grid surface.
The grid outline will disappear and the paper will
away from the grid. At this moment, plunge the grid
For more details, see text.
- While keeping the grid under the surface of the
ethane, remove the tweezers from the plunging arm.
Raise the plunging arm away from the tweezers. Carefully
remove the tweezers and grid from the ethane
and immediately hand plunge the grid into the liquid
nitrogen. Make sure the grid stays under the cold
nitrogen vapour cloud to keep it from warming and
frosting. This can be achieved by having sides on the
polystyrene liquid nitrogen box that are higher than
the top level of the liquid ethane container.
- Transfer the grid, keeping it submerged under
the liquid nitrogen, into one of the storage slots in the
grid box. After all samples have been loaded, close the
lid firmly with a screwdriver. Place the liquid nitrogenfilled
transfer dewar containing the polypropylene
tube close to the plunger. Using a large pair of tweezers,
quickly transfer the grid box from the plunger into
the tube. Screw the lid onto the tube.
- Move the tube and grid boxes holding the
samples into a long-term storage dewar. As long as the
nitrogen is kept topped off the grids can remain here
for long periods (sometimes years) without deterioration
until you are ready to view them in the microscope.
We totally dry our dewars once a year with a
stream of dry nitrogen gas to remove any frost that
may accumulate with time. Transfer all grids to
another frost-free container while drying the original
6. Image Acquisition
- Using a precooled pair of large tweezers, remove
the grid storage box holding the frozen grids and place
in a liquid nitrogen-filled transfer dewar.
- At the electron microscope, fill the anticontaminator
and cryo-blade dewars.
- If the cryo holder has been pumped (it is recommended
to continually vacuum pump the cryo holder
dewar when not being used), remove it from the
vacuum pump stand. Insert the holder carefully into
the loading station. Connect the cable for temperature
measurement between the cryo holder and the temperature
- Remove the clip ring from the cryo holder with
the clip ring tool.
- Put on eye protection or a face shield.
- Fill the specimen holder dewar and the loading
station dewar with liquid nitrogen. After each filling
with liquid nitrogen, replace the Plexiglas lid on the
loading station dewar to minimise frost contamination.
Keep the cap on the holder dewar, as it keeps the
outer sides from cooling. Make sure you open and
close the tip shutter as you cool as this prevents the
shutter from jamming. You will need to keep topping
off the level so that nitrogen sits just below the tip
of the sample holder. Wait for the temperature controller
to read at least -160°C before loading your
- Placing the transfer dewar as close as possible to
the specimen holder dewar, rapidly transfer the grid
box holding the frozen specimens to the specimen
holder dewar. Use a precooled pair of large tweezers.
- Precool a screwdriver and loosen the lid of the
- Check that the level of the liquid nitrogen is just
below the tip of the holder; cool a pair of No. 5 forceps
and the clip ring tool in the loading station dewar.
- Using the forceps, remove a grid from the grid
box. Making sure you keep the grid within the cold
nitrogen vapour, place the grid into the recess in the
cryo holder tip. Centre the grid in the recess and fasten
with the clip ring. Immediately close the holder tip
- Replace the Plexiglas lid on the loading station
dewar and lift the loading station and cryo holder and
place onto the console desk of the microscope. We
15rotect the console surface from nitrogen damage with
a metal splash guard (Gowen and Burger, 1998).
- Cycle the roughing pump on the microscope so
that the pump lines to the microscope sample stage
(e.g., compustage on an FEI machine) are under good
vacuum before insertion of the cryo holder.
- Remove the holder from the loading station and
carefully insert into the microscope stage for preevacuation.
(It can be useful to pretilt the stage to around
-60° so that the holder dewar neck is positioned at
"3 o'clock" instead of "6 o'clock" to lessen nitrogen
spillage when the holder is introduced.) Wait for the
vacuum to be restored and then insert the holder all
the way into the microscope.
- Allow at least 20min for the temperature to
stabilise and the vacuum to recover. Temperature variations
cause the holder to drift. Connect the cable from
the temperature controller and monitor the temperature.
During the transfer the temperature should not
warm to above about -150°C. Normally the holder will
stabilise at around -180°C. Do not leave the cable connected
to the holder as this can also cause drift effects.
To reduce damage when viewing cryo grids in the
electron microscope, it is necessary to have low-dose
software, which allows you to minimise the electron
dose applied to the sample. Always use the beam
blanker when you are not observing the specimen. The
microscope should be well aligned and set up for lowdose
conditions. To lower the electron intensity at the
sample, you can use a small condenser aperture, small
spot size, and low emission settings.
- Once the holder temperature has equilibrated
and drift minimised, the grid should be screened at
low magnification (usually at around 3000x or less) to
find promising cell areas for imaging.
- Images can be acquired either digitally using a
CCD camera system or taken onto plate film. If
required, images on plate film can be scanned digitally
using a suitable high-resolution film scanner.
- In some instances, micrographs may appear to
have an uneven intensity difference across the image
due to local variations in ice thickness. Applying a
high-pass filter helps even the intensity, which allows
subsequent contrast adjustments to be made more
easily (Fig. 1b).
C. Correlation with Light Microscopy
|FIGURE 5 Correlative light and electron microscopy as
by a B16 mouse melanoma cell: (a) Cell
stained with fluorescently
labelled phalloidin and visualised
microscopy, (b) corresponding view of
the same cell afer negative
staining, (c and d) higher
magnification insets of the previous
Bars (a and b) = 10 µm, (c) = 1 µm, (d) = 250 nm.
For situations in which the cell shape cannot be
visualised clearly (e.g., frozen hydrated samples) or in
which the function cannot be deduced solely from the
morphology but only from a marker protein (e.g., GPFVASP
for protruding lamellipodia; Rottner et al.
correlative light and electron microscopy on the
same cell can be very helpful in establishing the
structure-function link as discussed earlier. This correlation should be relatively straightforward for the
classical methods, where cells are grown directly on
glass coverslips with etched finder patterns; appealing
examples can be found in Svitkina and Borisy (1999).
For negative staining and cryo-EM, where cells are
grown directly on grids, finder grids with indexed grid
holes have to be used (Fig. 5).
- Prepare grids with a Formvar or a holey carbon
film as described earlier; use H2 Ni finder grids instead
of hexagonal 200 mesh grids to be able to relocate your
- Spread cells on these grids. For correlative
LM/EM, it is essential to remove any air bubbles
trapped in the mesh by preincubating them in water
as described previously.
- Mount the grids individually for light
microscopy; how exactly this is done depends on the
setup and the cell line used. As a general guide, care
has to be taken to (1) mount the grid flat (e.g., by using
the method described earlier with coverslips with a
hole drilled in the center), (2) have the cells as close to
the objective to avoid problems with focus, and (3)
mount grids with a carbon film upside down for epifluorescence
to avoid loss of intensity. Two sample
setups using petri dishes with a glass coverslip at the
bottom are shown in Fig. 2c.
- For the light optical observation of the cells, there
are several possibilities available: (1) living cells in
phase contrast, (2) living cells expressing a fluorescent
protein (conjugate), and (3) fixed cells after staining
with a fluorescent probe. For the latter method, it is
essential to minimise the damage of the substructure by prolonged incubation steps in the staining protocol,
e.g., fluorescently labelled phalloidin can be included
directly in the fixative.
- Record the position of each cell observed in a
map of the finder grid; such a map for the H2 grids is
provided by the authors at ftp://cellix.imba.oeaw.ac.
- Proceed as described earlier for negative staining
The authors thank Johanna Prast, Institute of Molecular
Biology, Salzburg, for her continuous support,
Marietta Schupp, Photo Lab, EMBL Heidelberg, for her
contributions (Figs. 3 and 4), and Johann Diendorfer,
Institute of Molecular Biology, Salzburg, for his help
with producing coverslips with holes. GPR is supported
by project P-15710 of the Austrian Science
Gowen, B. E., and Burger, L. (1998). Cryo-TEM liquid nitrogen
splash guard. J. Micros
Heuser, J. E., and Kirschner, M. W. (1980). Filament organization
revealed in platinum replicas of freeze-dried cytoskeletons. J. Cell
Hodgkinson, J. L., and Steffen, W. (2001). Direct labeling of components
in protein complexes by immuno-electron microscopy. Methods Mol. Biol
Medalia, O., Weber, I., Frangakis, A. S., Nicastro, D., Gerisch, G., and
Baumeister, W. (2002). Macromolecular architecture in eukaryotic
cells visualized by cryoelectron tomography. Science 298
Resch, G. P., Goldie, K. N., Krebs, A., Hoenger, A., and Small, J. V.
(2002). Visualisation of the actin cytoskeleton by cryo-electron
microscopy. J. Cell Sci
Rottner, K., Behrendt, B., Small, J. V., and Wehland, J. (1999). VASP
dynamics during lamellipodia protrusion. Nature Ceil Biol
Small, J. V. (1988). The actin cytoskeleton. Electr Microsc. Rev
Small, J. V., and Herzog, M. (1994). Whole-mount electron
microscopy of the cytoskeleton: Negative staining methods. In
"Cell Biology: A Laboratory Handbook"
(J. E. Celis, ed.), Vol. 2, pp.
135-139. Academic Press, San Diego.
Small, J. V., Rottner, K., Hahne, P., and Anderson, K. I. (1999). Visualising
the actin cytoskeleton. Microsc. Res. Tech
Small, J. V., and Sechi, A. (1998). Whole-mount electron microscopy
of the cytoskeleton: Negative staining methods. In "Cell Biology:
A Laboratory Handbook"
(J. E. Celis, ed.), 2nd ed., Vol. 3, pp.
285-291. Academic Press, San Diego.
Svitkina, T. M., and Borisy, G. G. (1999). Arp2/3 complex and actin
depolymerizing factor/cofilin in dendritic organization and
treadmilling of actin filament arrays in lamellipodia. J. Cell Biol
Svitkina, T. M., Verkhovsky, A. B., and Borisy, G. G. (1995). Improved
procedure for electron microscopic visualization of the cytoskeleton
of cultured cells. J. Struct. Biol
Vignal, E., and Resch, G. P. (2003). Shedding Light and Electrons on
the Lamellipodium: Imaging the Motor of Crawling Cells.