Field Emission Scanning Electron
Microscopy and Visualization of
the Cell Interior
Resolution in scanning electron microscopy (SEM)
has improved dramatically in recent years so that for
the majority of biological material, no significant differences
exist in resolution between SEM and conventional
transmission electron microscopy (TEM). High
brightness sources (field emission) and novel final lens
configurations have resulted in instrument resolutions
of 0.5 to l nm, allowing direct, in situ
visualization of surface detail at molecular
resolution. As all this technology relies on field emission
sources of the electron beam, either by "cold"
field emission or thermally assisted "Schottky" field
emission, we refer to it as FESEM.
Surface imaging allows bulk samples to be examined
without limitation of specimen thickness. Visualization
of intracellular surfaces requires some means
of access, such as isolation of cell fractions or macromolecules,
or in situ
, via fracture, or sectioning
techniques. Cell-free systems, e.g., in vitro
formation, allow biological interfaces such as developing
nuclear envelopes to be imaged directly (Goldberg et al.
, 1992). True three-dimensional (3D) surface
visualization can be achieved by tilting the specimen
to make stereo pairs, and accurate surface measurements
can be made from computerized 3D reconstructions.
The surfaces can be characterized further by
immunogold labeling, which can be unequivocally
localized by the strong backscatter signal of the gold
probes. For specimens that are thin enough to allow
electron penetration, a scanning TEM (STEM) image
can also be obtained readily and displayed simultaneously
alongside the secondary electron image,
producing complementary information from the
transmitted beam/specimen interactions. The use of
low accelerating voltages in FESEM has also been
shown to be of advantage, reducing charging and
penetration of the electron beam, but maintaining a
high-resolution information content. High-pressure
freezing, freeze substitution, and examination of cryohydrated
specimens may all be used for FESEM
(Muller and Hermann, 1990; Walther, 2003) but can be
considered specialized and are not covered in this
article, although we do describe the use of cryoultramicrotomy
and cryoabrasion as techniques to access
internal surfaces within the cell prior to conventional
imaging by FESEM. Basically, we deal with techniques
that rely on chemical preservation, followed by dehydration,
critical point drying, and coating. Conventional
SEM coating (up to 20nm thickness) with
sputtered gold completely obscures fine surface detail
in HRSEM and must be replaced by high-resolution
coating. We routinely coat with a 1- to 2-nm film of
chromium or tungsten, which has a grain size of 0.3 to
0.5 nm (Apkarian et al.
II. MATERIALS AND
- Glutaraldehyde (Agar Scientific)
- Tannic acid (TAAB Laboratories)
- TCH (Cat. No. T-2137. Sigma)
- Osmium textroxide (Agar Scientific)
- Uranyl acetate (Agar Scientific)
- Molecular sieve (Merck Ltd.)
- Arklone (trichlorotrifluoroethane) Taab Labs UK
- HEPES (Sigma)
- PIPES (Sigma)
- Phosphate-buffered saline (PBS)
- EDTA (Sigma)
- Phenylmethylsulfonyl fluoride (PMSF, Sigma)
- Percoll (Sigma)
- Sorensen's phosphate buffer
- Poly-L-lysine HBR, MW 150,000-300,000 (Cat.
No. P-1399 Sigma)
- Glass coverslips, 5-7mm diameter
- Silicon chips, 5 × 5 mm2 (Agar Scientific Ltd.)
- Carbon-coated support grids
- Fine forceps for handling
- Microcentrifuge to spin suspended material
- Microcentrifuge tubes, 1.5ml, half-filled with
polymerized EM resin, ideal for supporting coverslips,
chips, and grids during specimen deposition by
- High-resolution scanning EM. Conventional
"pinhole" final lens instruments with field emission
sources will allow subcellular imaging, as will conventional
transmission instruments with scanning
attachments. To date, the highest resolution achieved
has been in field emission instruments with the facility
to position the specimen in, or very close to,
the final lens. Recent technology has significantly
improved the resolution of field emission instruments
at lower accelerating voltages (1 kV), from around 4 to
1.5 nm, facilitating imaging without the need for metal
coating. The microscope should also be equipped with
a suitable high-resolution backscatter detector for
immunogold labeling. The main suppliers for these
instruments are Hitachi, Jeol, Philips, and Leo.
- Critical-point drier, with high-purity CO2 (<5 ppm water). Liquid CO2 should be passed through
a filter to remove water (Tousimis Research Corp.,
- Coating units. Oxygen, hydrocarbons, and
water vapor all adversely affect the grain size of
chromium or tantalum deposited by sputter coating
(Apkarian et al., 1990). We use an Edwards Auto 306
12-in. coating unit with cryopump, magnetron head,
and suitable power source (Edwards High Vacuum
International). Similar configurations using Denton
HiVac and Balzers equipment with cold-trapped turbopumping
have also been successful. Any system
should use high-purity argon and have a shutter and
a specimen table that tilts and rotates. Film thickness
monitoring of coating deposition is an advantage.
Several "benchtop" systems are currently on the
market, many untested by the highest resolution. In
our own very recent experience, the provision of a suitably
performing coating system has often proved to be
a limitation of the perceived performance of a newly
acquired microscope, and suitable resources for a highresolution
coating system should be incorporated in
any application for a field emission instrument. If
possible, get in touch with an established group and
get advice before committing to any particular coating
It is crucial to visit manufacturers' demonstrations
with the material that will be investigated to ascertain
that suitable performance can be assured from the
III. PROCEDURES A. Exposing Surfaces within the Cell
1. Subcellular Fractionation
Organelles and macromolecules can be isolated by
standard procedures, possibly requiring subsequent
modifications in the light of HRSEM visualization,
which are beyond the scope of this article. Basically,
the specimens must be undamaged by osmotic shock,
proteolysis, or unsuitable isolation buffers. They must
also be clean. The surface of organelles should, for
instance, be free of attached cytoskeletal remnants or
cytoplasmic contamination. Where the specimens are
available as purified macromolecules or viruses, they
may be deposited on carbon-coated TEM grids in the
conventional manner and viewed by HRSEM. In this
situation, TEM negative staining will usually be
replaced by fixation for SEM and air drying replaced
by critical-point drying followed by chromium
coating. If a STEM detector is available, the virus/macromolecule can be recognized as a transmitted
"reference image" after this protocol and compared
directly with the secondary electron (SEM) image. The
thin coating of chromium applied for the secondary
electron imaging does not interfere with the STEM
Adhering Sample to Support
Many cell components naturally adhere to glass
coverslips, silicon chips, or carbon support film on
grids. Glass coverslips may be a useful initial prepar ative substratum, as they can be checked in the phasecontrast
microscope for the density and distribution of
specimens and for the progression of various protocols
such as detergent extraction of cytoplasm. Once the
isolation protocol is established, coverslips should be
replaced with silicon chips as specimen substrates, as
silicon is a conductive substratum in contrast to glass,
an insulator, which can generate problems with charging
in the SEM. Tissue culture cells will grow in identical
fashion on silicon as they do on glass or plastic,
and isolated cytosol or organelles will also adhere naturally
to silicon in the same way as they do to glass. If
samples are fixed in suspension it may be necessary to
coat the support with poly-L
-lysine to facilitate adherence.
Different samples may require slight modification,
but the basic technique is as follows (Fig. 1).
|FIGURE 1 Xenopus nuclear assembly egg extract spun onto a silicon chip, fixed, frozen, and sandpapered
while frozen showing a section through the edge of a nucleus where the two membranes of the nuclear envelope
can be seen, as well as the chromatin and nucleoskeletal fibres on the nuclear interior. Bar: 125 nm.
: Make a fresh 1-mg/ml solution of
-lysine in sterile distilled water; use within 24h.
2. "In Situ" Exposure of Intracellular Surfaces
- Mark the surface of chip or coverslip with identification
number using a diamond marker.
- Place a 50-µl drop of poly-L-lysine on coverslip
or chip; allow to stand for at least 60min in a moist
chamber to avoid drying. Rinse in sterile distilled
water. The surface will retain its adhesive properties
for up to 2 weeks in a refrigerator.
- Place a 50-µl drop of suspended material (fixed
and rinsed) on coverslip/chip. Allow to settle and
attach at 4°C and unit gravity in a moist chamber (1 h
to overnight). Alternatively, spin directly (see isolation
of nuclei) onto silicon chip.
- Allow bulk of drop to run off, put chip/coverslip
back in fixative, and continue as for fresh tissue.
Unfixed living samples (e.g., whole cells) may be distorted
by poly-L-lysine. To spin down materials from
suspension, use minicentrifuge tubes half-filled with
polymerized EM resin to support the coverslip/silicon
This is a simple but extremely effective way of
exposing internal surfaces in both tissues and cells.
After fixation, dehydration, and critical-point drying, merely gently press the surface of the specimen to a
square of double-sided tape mounted to a second
silicon chip and pull away without shearing, coat both
chips as normal, and examine in the SEM. The fracture
will remove material on the surface of the adhesive
and leave fractured material "in situ
.'" This technique
may be enhanced by pretreatment with detergent
(0.5% Triton X-100, 2-3min for tissue culture cells),
either alone or mixed with the primary fixative (2.0%
paraformaldehyde and 0.1% glutaraldehyde), and
subsequently refixed as described (Allen et al.
These methods involve sectioning of embedded
specimens followed by exposure of internal surfaces
by removal of the supporting material. This may vary
among epoxy resins, various waxes, and even ice.
Resins that require corrosive solvents for removal will
tend to be prone to surface etching. A mixture of 50%
propylene oxide and 50% sodium methoxide (dissolve
2g NaOH pellets in 100ml absolute methanol) will
remove most resins.
3. Cryo Methods to Expose Internal
Surfaces for FESEM
|FIGURE 2 Low-power image of cryo-planed block face
of a mitochondrial
pellet where the internal cristae
structure has been
exposed. Scale bar: 667 nm.
Surface imaging by FESEM may be achieved by isolating
the cellular component of interest, such as mitochondria,
but this approach does not allow access to
the interior structure of such an organelle (see later).
One way to expose such surfaces is to freeze the cells
or tissue and cut cryo sections, which themselves can
be viewed in the SEM, to "cryoplane" and expose the
whole blockface in the SEM, or to "cryoabrade" the
surface of the frozen sample and expose surface features
in a different way. Samples are then thawed and
processed for FESEM as normal. This gives a crosssectional
view but with much greater depth of information
than in a resin-embedded thin section viewed
in the TEM because the sections can be very thick,
they are resin free, and there is a greater depth of focus.
Information can also be gathered quite simply in 3D
simply by taking stereo pairs, which also allows computerized
3D reconstruction and measurement of the
surface. This method is also compatible with immunogold
Cryomicrotomy is an adaptation of the widely
used "Tokyasu" technique (Tokyasu, 1986, 89) for
4% paraformaldehyde in PBS
sucrose in PBS
Cryo ulramicrotome (e.g., Leica Ultracut R with FCS
- Fix samples (e.g., pellet of organelles) with 4%
paraformaldehyde in PBS at room temperature for
- If possible, trim to a cube of about 1 mm3.
- Transfer to 2M sucrose in PBS overnight at 4°C.
- Mount sample on cryo stub for ultramicrotome
and wick off excess sucrose.
- Plunge into liquid nitrogen.
- Mount into a cryo ultramicrotome, which has been
precooled to -100°C.
- Cut sections of 100-300 nm thickness.
- As in the Tokyasu technique, pick up sections off
the knife on a drop of sucrose suspended from a
loop, where they thaw, and then touch the sucrose
drop to a silicon chip where the sections adhere.
- Place chip in PBS to wash off sucrose.
- This can then be immunogold labeled, refixed, and
processed for FESEM.
These problems are associated with the Tokuyasu
- Poor infiltration of sample with sucrose leading to
- Curling of sections.
- Small sample size.
When processing the sample remaining on the pin
for SEM, the specimen always detaches from the pin.
This leaves a very small specimen that is easily lost
during the dehydration and CPD steps. An additional
problem is the attachment of the sample to a silicone
chip after CPD. The sample is very fragile and it is not
always easy to identify the "planed" side. Adhere the
sample to the chip using a thin smear of silver dag. It
is possible for the sample to flip over during this
process. Also, because of the irregular shape of the
sample, ensuring good contact and therefore a good
earth path from specimen to chip can be tricky. It is
easy to submerge the specimen in too much dag.
Specific Protocol for Mitochondrial Isolation and
Exposure of Internal Structure by FESEM
- Centrifuge organelles onto a silicon chip.
- Fix (e.g., 4% paraformaldehyde in PBS, room temperature
- Transfer to 2M sucrose in PBS for 2h to overnight
- Remove from sucrose and place on filter paper to
dry the back of the chip.
- Wick off most of the sucrose from the sample,
leaving as thin a film as possible without drying.
- Plunge into liquid nitrogen.
- The sucrose step can be avoided if the sample can be
frozen ultra-rapidly by plunging into liquid propane
or ethane. The chip can then be held under liquid
nitrogen or on a liquid nitrogen-cooled platform
(such as the Leica EM CPC) while it is abraded with
fine sandpaper (400-600 grade wet and dry abrasive
as per automobile body and paintwork).
- Thaw into fix and process for FESEM.
Isolate mitochondria using the differential centrifugation
method (Gottlieb et al.
, 2003). Harvest and place
(2.5-5 × 108
) on ice for 15min, centrifuge at 500g
for 5 min at 4°C, wash with ice-cold PBS, and subsequently
wash with ice-cold mitochondrial isolation
buffer (MIB) (200mM
HEPES, 0.5 mg/ml BSA; pH 7.4). Resuspend
cells in ice-cold MIB and then homogenize in a
syringe-driven cell disruptor. Spin the lysate at 800g
for 10min at 4°C. Remove supematants and spin at
for 10min at 4°C. Add fixative (3% glutaraldehyde)
to the pellets and keep samples at 4°C for 1h. Remove the fixative carefully and infiltrate
the pellet with sucrose/PVP solution overnight
(Tokuysau, 1989). During this process, leave the pellet undisturbed. Then carefully excise small (>1mm2
pieces of sample from the pellets and mount onto aluminium
plunge freezing pins (Leica Microsystems,
Milton Keynes). Mount the pins into a plunge freeze
unit (Leica CPC) and freeze in liquid propane at a temperature
of -182°C. Transfer the frozen specimen and
pin under LN2 into a cryo ultramicrotome (Leica
Ultracut S with FCS attachment). Using a diamond
trimming knife (Diatome Cryotrim 45), trim several
semithin sections (350 nm) from the sample in order to
remove surface sucrose. Cut further semithin sections
(350-400nm) from the sample using a diamond cryo
knife. Collect each section on a sucrose loop according
to the "Tokuyasu" technique (Tokuyasu, 1986). Thaw
the frozen sections onto 5-mm silicone chips and
process for SEM as follows.
- Transfer the chips with attached sections using a
metal loop (2-3 mm diameter) and invert such that the
chip is floated, section side down, in a plastic Petri dish
(35-mm Falcon) containing double-distilled water.
- Wash 3x over 15min in order to rinse out the
- Fix by floating in 1% OsO4 in double-distilled
water for 1h.
- Wash in double-distilled water 3x for 5 min each.
- Then dehydrate the sections and critical point
dry as described later.
Aligning the cryo abrasive pad with the specimen
is very tricky and must be done with care. It is very
easy for the protruding abrasive shards of the wet and
dry paper to embed themselves into the frozen block.
Also, if the section advance is too great, the sample
block can be literally ripped from the specimen pin. A
few micrometres must be shaved off the sample face
in order to ensure that all surface sucrose has been
removed. This only leaves a few micrometres of wellfrozen
vitrified sample to work with. Once again, the
sample size is very small and is easily damaged or lost
in subsequent processing and mounting steps.
Identifying the abraded face can be tricky even
under a stereomicroscope. The swirled pattern of the
specimen pin that has been embossed into the underside
of the sample can look very similar to the abraded
face, leading to the specimen being mounted pin
All fixatives are ideally made up just before use
or at least the same day; both glutaraldehyde and glutaraldehyde-tannic acid solutions should be filtered
before use through a 0.22-µm filter. The 1% aqueous
uranyl acetate should be stored in a brown bottle.
Osmium tetroxide is made by breaking the glass
ampoules in which the crystals are delivered, having
previously washed them free of label and adhesive
under the tap, in a fume cupboard. The ampoules plus
crystals are dropped in the correct amount of buffer
or distilled water where the osmium dissolves to give
the appropriate final concentration. (Note
: Osmium is
extremely hazardous and appropriate precautions
must be observed.) Thiocarbohydrazide or tannic acid
solutions should also be made just prior to use (Allen et al.
Always use glutaraldehyde of EM-grade quality
from a high stock concentration (50%) stored in a
freezer. Low concentration stock solutions and storage
in large bottles at room temperature will reduce the
cross-linking properties of the glutaraldehyde.
1. Isolated Proteins and Nucleoproteins
2. Small and Easily Preserved Structures
- Proteins and nucleoproteins on carbon support
films on TEM grids may be floated on top of drops
(25-50µl) of the appropriate solutions spread on
- Place in 1% glutaraldehyde in appropriate buffer for
- Wash in double-distilled water for 5 min.
- Transfer to 1% uranyl acetate for 5 min.
- Transfer to 100% ethanol for 1-2 min.
- Air dry or critical point dry (see later).
3. Large and/or Fragile Structures (e.g., Whole Cells,
Organelles, Cytoskeletal Preparations, Isolated
Cells, or Nuclear Membranes)
- Fix in 3% glutaraldehyde in Sorensen's phosphate
buffer for 30min.
- Wash in Sorensen's for 5 min.
- Postfix in 1% OsO4 (in Sorensen's for 30 min).
- Wash in double-distilled water for 5 min.
- Dehydrate through ethanol series for 5 min each.
- Place in Arklone for 5 min.
- Critical point dry (see later).
- Attach whole cells to specimen supports such as
silicon chips and handle by changing the solutions
in 35-mm-diameter petri dishes.
- Fix in 2% glutaraldehyde, 0.2% tannic acid, and
0.1% HEPES, pH 7.4, for 10min.
- Wash in double-distilled water for 5 min.
- Postfix in 0.1% OsO4 in water for 10min.
- Wash in water for 5 min.
- Stain with 1% aqueous uranyl acetate for 10min.
- Dehydrate through ethanol series and Arklone and
critical point dry.
- Isolation of Nuclei from Tissue Culture Cells
- Take approximately 10 million tissue culture cells
(usually from suspension culture), cool to 4°C and
pellet in a swing-out centrifuge (1000g for 10min).
- Wash the pellet in PBS buffer and then resuspend
in 8ml ice-cold swelling/shearing buffer (50mM Tris-HCl, pH 7.4, 5mM MgCL, 1.3mM EDTA, and
5mM phenylmethylsulfonyl fluoride added shortly
before use) in which nuclei are allowed to swell for
- Mechanically homogenize the cells using a
plunger-type tissue grinder (e.g., Kontes Dounce),
checking the number of strokes required for nuclear
release by phase-contrast light microscopy (usually
10-20 strokes). Precool the plunger in ice prior to
use. Alternatively, nuclei may be isolated by a
single passage through a 26-G3/8 syringe needle
- Prepare a Percoll gradient as follows: use a 13.5-ml
Beckman centrifuge tube filled with 0.86ml Percoll,
density 1.130g/ml, 2.74ml 10mM Tris-HCl (pH 7.4),
and 0.40ml 2.5M sucrose. Add density marker beads
to monitor gradient, red beads (1.12g/ml) in Percoll
containing 0.25M sucrose, and yellow beads
(1.049 g/ml) in Percoll containing 0.25 M sucrose. Spin
for 30 min at 30,000g at 4°C with a 60° angle head rotor;
red beads will form a line 5 mm from the bottom of
the tube, with the yellow beads a further 12 mm above.
- Gently layer the homogenate on top of the gradient
and spin for 10min at 30,000g, which generates
two bands from the homogenate. Damaged nuclei and
whole cells are found in the upper band 7mm above
the red beads, whereas the pure nuclear fraction is
found 0.5 mm below the red beads. Remove this fraction
and wash gently in 150mM Tris-HCl (pH 7.4) for
5 min at 4°C.
- Spin the isolated nuclei onto 5-mm poly-L-lysine-coated silicon slips. Spin the silicon chips in
1.5-ml Eppendorf tubes previously half-filled with polymerized
resin. Overlay the Si chips with 0.5 ml of freshly
isolated nuclear suspension and spin for 5 min at 1000g.
- Fix the whole chip (+nuclei) in 6% glutaraldehyde
in 0.15M Sorensen's buffer for 20min, rinse gently in buffer, and postfix in 1% osmium tetroxide
in 0.15M Sorensen's buffer for 1h. Dehydrate, critical
point dry, and coat with 3-4nm of tantalum or
5. Preparation of in Vitro-Assembled
Organelles for FESEM
- Low yield of nuclei. Dounce tissue homogenizers
are produced with different clearance distances
between the polished tube and the pestle. Some
homogenizers are designed just to disrupt tissues as a
necessary step prior to homogenization of nonsupension
cells. Make sure that the clearance distance of the
Dounce tissue homogenizer for the final release of the
nuclei is small enough to disrupt whole cells. Some
manufactures offer pestles with two different clearance
distances to allow tissue disruption and release of the
nuclei within one and the same tube. For certain cell
types, making the buffer more hypotonic can help
increase the yield of nuclei.
- Enzymatic degradation of structures. It is good
practice to ensure cooling for every preparation step
prior to the fixation of nuclei on silicon chips. Moreover,
the homogenate should be processed without delay.
Therefore, preparation of the Percoll gradient before
homogenization is recommended. To ensure a suitable
ratio between biological material and buffer, a homogenizer
with a sufficient capacity (for 8ml) should be
selected. Do not reduce the number of cells significantly
to keep this ratio in a homogenizer with lower capacity,
as this might cause problems with recognition of the
nuclear layer after gradient centrifugation.
Organelles, such as nuclei, endoplasmic reticulum
(ER), and Golgi, can be assembled in cell-free extracts.
Extracts made from frog eggs are a particularly powerful
system for studying the assembly, dynamics, and
functions of these organelles. Organelles can be isolated
cleanly from the extract and their surfaces can be
examined by FESEM. In vitro
-assembled nuclei, as well
as ER, can be prepared for FESEM as follows.
- Membrane wash buffer (MWB): 250mM sucrose,
50mM KCl, 50mM HEPES-NaOH (pH 8.0), 1µg/ml
aprototin, and 1 µg/ml leupeptin
- Fix buffer: 150mM sucrose, 80mM PIPES-KOH
(pH 6.8), 1 mM MgCL, 2% paraformaldehyde, and
C. Critical-Point Drying
- Prepare Xenopus egg extracts (Newmeyer and
Wilson, 1991) and incubate extract with demembranated Xenopus sperm chromatin to assemble nuclei.
ER and Golgi will also assemble in the same extract.
- After the required time, remove a 4-µl extract,
place in a 1.5-ml Eppendorf tube, and resuspend very
gently in 1 ml of MWB. At this stage, centrifugation of in vitro nuclei and organelles onto 5-mm2 silicon chips
requires a simple modification of the 1.5-ml Eppendorf
centrifuge tubes as follows. Remove lid from tube and
cut tube with a sharp knife at a level where the cut end
fits tightly into the lid, thus creating a "flat-bottomed"
tube. Snap the cut end into the lid, having placed a
silicon chip in the lid first.
- Pipette the 1ml of MWB containing the extract
into the modified tube, spin in a swing-out rotor, inside
a 10-ml centrifuge tube (with a single tissue as cushion),
and spin for 10min at 4°C or room temperature
at 2000g. Some leakage of the suspension at the joint
between the cut end of the tube is not a problem at this
- Pipette off most of the buffer, break open the
tube, and remove the chip. Place the chip in 5ml
of fix buffer in a small petri dish for 10min at room
- Wash chip in 0.2M cacodylate (pH 7.4), place in
1% OsO4 in 0.2M cacodylate for 10min, wash twice in
distilled water, and place in 1% aqueous uranyl acetate
for 10min (at room temperature) and then dehydrate,
critical point dry, etc.
All traces of water should be removed from ethanol,
Arklone, and CO2
. Let 100% ethanol and Arklone stand
over molecular sieve for more than 24h prior to use.
High-purity liquid CO2
(less than 5 ppm water) should
be used and passed through a water filter as a
- Exchange Arklone for CO2.
- Flush six times.
- Leave in CO2 for 30min.
- Flush six times.
- Raise temperature to 40°C.
- Release gas slowly (over about 15-20min).
- Transfer to coating unit as soon as possible.
Critical-point-dried samples should be transferred
immediately into the sputter coater to avoid rehydration,
and coated samples are best viewed in the microscope
directly. However, if it is known that the
microscope cannot be accessed, it is better to pause preparations after critical-point drying and store preparations
D. Sputter Coating
- Pump specimen to at least 5 × 10-7mbar.
- Introduce high-purity argon to a pressure of
8 × 10-3mbar.
- Start specimen rotation (60rpm).
- Sputter at 50-100mA current (voltage 450V) and
60rpm; specimen table should be tilted at 30°. Presputter onto the shutter for 20-60s to remove
the chromium oxide layer from the target.
- Open shutter and deposit 2nm chromium as indicated
by a film thickness monitor (usually 20-30s).
- Examine in microscope as soon as possible, preferably
within a day or two. Coatings are variable
according to the specimen, but a general rule is that
they deteriorate with time, usually over a few days
to about 2 weeks.
F. Immunogold Labeling
- A liquid nitrogen-cooled decontaminator (if
present) should always be used; this is more likely to
be fitted on an "in-lens" electron optical column configuration.
Many recent field emission instruments are
of conventional "pinhole configuration," but with very
short working distances to optional decontamination
- Spot size and apertures should be as small as
possible, consistent with a sufficient signal to visualize
high resolution of specimen at photographic collection
rates (e.g., 40-s scans).
- An appropriate accelerating voltage must be
selected. High-resolution scanning electron microscopes
usually work in the range of 1-30kV. Instrument
resolution decreases with decreasing accelerating
voltage; however, at high voltage there may be problems
with charging and specimen penetration, leading
to a nonspecific signal from below the specimen
surface. At low voltage, penetration and charging are
reduced, but so are resolution and signal. Signal is generated
almost completely from the surface at 1.0 kV so
there is no problem of a "bulk" signal from underlying
structures. In general, we use high kilovolts for thin
and conductive specimens and lower kilovolts for
bulky or less conductive specimens; however, a wide
range of kilovolts should be experimented with for
each type of sample. The more recently produced field
emission instruments have vastly improved low kilovolt
performance, and specimens that have some inherent
conductivity as a result of osmium fixation can be
viewed uncoated at low kilovolts, without compromising
signal and resolution.
The basics of specimen preparation for immunogold
labeling are beyond the length limits for this
article and are adequately covered elsewhere (see
article by Roos et at. for additional information). For
immunogold labeling for HRSEM, the following
points are important.
1. Size of Probe
The choice of probe size is a compromise between
sensitivity and subsequent detection. Very small gold
probes (around l nm) have minimal steric hindrance
and consequently label with maximum sensitivity.
One-nanometer gold has been visualized by backscatter
imaging in HRSEM (Hermann et al.
, 1991), but this
is at the limits of resolution and is best increased
in diameter in situ
by silver or gold enhancement to a
size at which it can be visualised more easily (around
5-10nm). We have used both 5- and 10-nm gold as a
good compromise between sensitivity and localization.
Because most modern instruments will discriminate
easily between 5- and 10-nm labeling, these can be used
together successfully for double-labeling studies.
Using gold probes obviously prohibits gold coating
for SEM. In the past, gold-labelled specimens have
been coated with carbon, mainly to inhibit charging,
but carbon produces a severely limited secondary electron
signal and, consequently, little topographical
information. We have found that a 1.5-nm chromium
coating provides the ideal solution, retaining the full
secondary electron-generated surface information,
without compromising the detection of gold by
backscattered electron detection (Allen and Goldberg,
In this situation, having found that "mixed"
imaging of SE and BSE signals was not satisfactory, we
have chosen to collect each signal separately (but
simultaneously) and then to superimpose the gold BSE
signal onto the secondary signal (retaining register) in
Adobe Photoshop, often altering the colour to improve
the appearance of label against the monochrome background.
In modern instruments with good low kilovolt
performance, uncoated or carbon-coated imaging
will generate such a strong signal from gold probes
that they are observed easily in secondary electron
Although field emission SEM has been available for
some time, it is still a relatively new technique in cell
biology. The procedures given here may need to be
modified to optimize the preservation of some structures.
Probably the most difficult step is exposing
recognizable and undamaged intracellular surfaces.
Isolation of organelles offers the possibilities of further
characterization by other methods, but gives no "in
situ" information and may involve extensive biochemical
protocols. Resinless sections and dry fracture give in situ
information, but only after some initial extraction
of the cell. Freeze fracture, followed by frozen
hydrated coating and visualization, may alleviate
these problems but is limited by the plane of fracture,
as the structure of interest may not be exposed. It is
also technically difficult and expensive. Osmium
etching results in spectacular images of intracellular
membranes, but the uncertainty of what is removed
makes interpretation difficult. Direct visualization of
biological interfaces in cell-free systems (e.g., in vitro
nuclear formation) is a particularly promising area
(Goldberg et al.
, 1992, 1997). Considerable fresh structural
information has also been demonstrated for
nuclear pore complexes and associated structures
(Ris, 1991; Goldberg and Allen, 1992, 1996; Kiseleva et al.
T. D. Allen, S. Rutherford, and S. Murray are supported
by CRUK and M. W. Goldberg is supported by
a Wellcome Lectureship. The mitochondrial pellets
were supplied by Dr. E Gottlieb (Beatson Institute).
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along microtubules in vitro
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structure of human metaphase chromosomes determined by
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(K. W. Adolph, ed.), Vol. 11, pp. 52-70. CRC Press, Boca Raton,
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Kiseleva, E., and Goldberg, M. W. (1998). Three dimensional
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electron microscopy of the nuclear envelope: Demonstration
of a new regular, fibrous lattice attached to the baskets of the
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and lamina: Three dimensional structures and interactions determined
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field emission in-lens scanning electron microscope to study the
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Dimples, pores, star rings and thin rings on growing nuclear
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and cytochrome c release during apoptosis. Cell Death
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immunostaining electron microscopy using Fab fragments
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Kiseleva, E., Goldberg, M. W., Daneholt, B., and Allen, T. D. (1996).
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