Electron microscopy
The value of the electron microscope lies in its great resolving power. Transmission Electron
Microscopes today are capable of resolving objects only 0.2 nanometers apart (one nanometre
is a millionth of a millimetre) - just five times the diameter of a hydrogen atom, while in
comparison the bright-field light microscope, has a resolution of approximately 0.2
micrometers and a useful magnification of X 2,000.
The greatest obstacle to examine biological material with the electron microscope is the
unphysiological conditions to which specimens must be exposed.
Since the material must be exposed to a very high vacuum (
to
Torr) when being
examined, it must be dried at some stage in its preparation. The biological specimen must be
stabilized (or fixed) so that its ultrastructure is as close to that in the living material when
exposed to the vacuum.
The limited penetrating power of electrons means that the specimens must be very thin or
must be sliced into thin sections (50 - 100 nm) to allow electrons to pass through. Because
ultra thin sections have little contrast, they must be stained with heavy metal salts to provide
contrast necessary to reveal details of the cells ultrastructure.
Specimen preparation for TEM
FIXATION
Fixation, in itself, is an artifactual process that disorts cell structure from its living state. Cells
are never completely free of fixation artifact. The goal is to minimize fixation artifact to the
point that is as close to the living state as possible.
I. Chemical fixation:
Tissues can be exposed to primary fixatives in two ways: a) immersion fixation or b) vascular
perfusion fixation. Immersion fixation, for most tissues, is less desirable than vascular
perfusion fixation. The animal´s own vascular system is ideal to disseminate fixative to tissue
as compared with relatively slow penetration of fixative in tissue fragments that have been
mechanically disturbed by excision. There are numerous reasons to employ perfusion fixation
whenever possible, however, the size of animal being perfused often determines the
accesibility to the vascular system. Other large animals cannot be perfused because their
vascular system would require an enormous volume of fixative.
Postfixation with osmium is important to obtain quality electron micrographs. The rate of
penetration of osmium is very slow (less than 0.5 mm in 1-2 hours depending on the
compactness of the tissue).
Cell cultures can be fixed in suspension and immediately pelleted by gentle centrifugation.
For cells grown on culture dishes or in flasks, the fixation is performed by the addition of
double strenght fixative to culture medium. Then the cells are scraped off and centrifuged into
a pellet. If the pellet is loose or is falling apart, it will be necessary to embed in gelatin or
agarose. The last step is to slice the embedded pellet into small blocks (1 mm3); the same
method of cutting for the tissue samples. The cells grown on cover slips are already ready to
be processed.
A number of factors affect the quality of fixation including pH, temperature, osmolarity,
fixation time, and sample size - these are likely to have to be optimised for the sample. The
most commonly used fixation protocol involves primary fixation with glutaraldehyde (1-2 %)
followed by secondary fixation with OsO 4 (2-4%). For immuno-studies, paraformaldehyde
(4-8 %) is often preferred as the primary fixative, sometimes with low concentrations of
glutaraldehyde (0.1-0.5 %).
Paraformaldehyde
Formaldehye reacts with many functional groups on proteins including amine, thiol, hydroxyl,
imidazoyl and phenolic groups. Importantly, the majority of reactions are reversible and so
extensive wash steps should be avoided. Formaldehyde does not cross-link lipids but unlike
glutaraldehdye, can cross-link DNA.
Glutaraldehyde
The best fixative for preserving fine structure, cross-links proteins rapidly and irreversibly.
Targets protein amino groups, lysine is the most important component of proteins although
glutaraldehdye also reacts with other amino acids including cysteine, histidine, tyrosine and
tryptophan. It probably also reacts with free amino groups on some lipids (eg. PS and PE)
although may not prevent lipid extraction during subsequent processing for resin embedding
unless secondary OsO4 fixation is also performed. Stock (25 %) glutaraldehyde solution is
relatively stable at 4°C or -20°C however, shelf-life is limited to a few weeks once added to a
buffer.
Osmium tetroxide
Most important features are its ability to cross-link lipids and the electron density of reduced
osmium that can provide a scaffold for further staining by lead to increase contrast. Primarily
used as a secondary fixative because of its slow tissue penetration rate and poor ability to crosslink proteins and carbohydrates. The most likely lipid targets are unsaturated fatty acids. OsO4
reacts with certain protein side chains including thiol, hydoxyl, phenolic, carboxyl and amino
groups; cyteine and methioine appear to be the most reactive amino acids towards osmium
tetroxide.
In details:
Reaction of formaldehyde with proteins
The aldehyde group can combine with nitrogen and some other atoms of proteins, or with two such atoms if they
are very close together, forming a cross-link -CH2- called a methylene bridge. Studies of the chemistry of
tanning indicate that the most frequent type of cross-link formed by formaldehyde in collagen is between the
nitrogen atom at the end of the side-chain of lysine and the nitrogen atom of a peptide linkage (Fig. 2), and the
number of such cross-links increases with time. The tanning of collagen to make leather is comparable to the
hardening of a tissue by a fixative. The fixative action of formaldehyde is probably due entirely to its reactions
with proteins. Initial binding of formaldehyde to protein is largely completed in 24 hours but the formation of
methylene bridges proceeds much more slowly. Substances such as carbohydrates, lipids and nucleic acids are
trapped in a matrix of insolubilized and cross-linked protein molecules but are not chemically changed by
formaldehyde unless fixation is prolonged for several weeks.
Fig. 2. Reactions involved in
fixation by formaldehyde. (A)
Addition of a formaldehyd
molecule to a protein. (B)
Reaction
of
bound
formaldehyde with another
protein molecule to form a
methylene cross-link. (C) A
more detailed depiction of the
cross-linking of a lysine sidechain to a peptide nitrogen
atom.
Practical considerations relating to formaldehyde
This is the most important bit. Formaldehyde penetrates tissues quickly (small molecules), but its reactions with
protein, especially cross-linking, occur slowly. Adequate fixation takes days, especially if the specimen must
withstand the osmotic and other stresses of dehydration and infiltration with paraffin. Brief fixation in
formaldehyde (ideally delivered by perfusion) can stop or greatly reduce autolysis and confer slight hardening
and some resistance (but not much) to liquids that are not iso-osmotic with the tissue. This can greatly improve
the structural integrity of cryostat and other frozen sections, especially if followed by infiltration with a
cryoprotectant such as sucrose (ideally 60 % but more usually 15-30 %).
When a specimen is dehydrated after only a few hours in formaldehyde, the largely unfixed cytoplasmic proteins
are coarsely coagulated. Nuclear chromatin, which contains DNA and strongly basic proteins, is also coagulated
by the solvent, forming a pattern of threads, lumps and granules. This is not unlike the appearance induced by
fixatives that contain acetic acid, but it is less satisfactory for identifying cell-types on the basis of nuclear
morphology. (After adequate formaldehyde fixation, chromatin displays a remarkably even texture, also of little
diagnostic value but possibly closer to the structure of the living nucleus.)
Glutaraldehyde
Glutaraldehyde has fairly small molecules, each with two aldehyde groups, separated by a flexible chain of 3
methylene bridges. It is HCO-(CH2)3-CHO. The potential for cross-linking is obviously much greater than with
formaldehyde because it can occur through both the -CHO groups and over variable distances. In aqueous
solutions, glutaraldehyde is present largely as polymers of variable size. There is a free aldehyde group sticking
out of the side of each unit of the polymer molecule (Fig. 3), as well as one at each end. All these -CHO groups
will combine with any protein nitrogens with which they come into contact, so there is enormous potential for
cross-linking, and that is just what happens (Fig. 4). There are also many left-over aldehyde groups (not bound to
anything) that cannot be washed out of the tissue.
Fig. 3. (A) Three representations of a molecule of monomeric glutaraldehyde. (B) Polymerization reaction of
glutaraldehyde, showing an aldehyde side-chain on each unit of the polymer.
Fig. 4. Reaction of
poly(glutaraldehyde) with amino
groups of proteins.
Practical aspects of glutaraldehyde fixation
Five important points must be remembered when using glutaraldehyde as a fixative for light or electron
microscopy.
1. If it's to be any use as a fixative, especially for electron microscopy, the glutaraldehyde solution must contain
the monomer and low polymers (oligomers) with molecules small enough to penetrate the tissue fairly quickly.
This means you must buy an "EM grade" glutaraldehyde (25 % or 50 % solution), not a cheaper "technical"
grade. The cheaper stuff, which is for tanning leather, consists largely of polymer molecules too large to fit
between the macromolecules of cells and other tissue components.
2. The chemical reaction of glutaraldehyde with protein is fast (minutes to hours), but the larger molecules,
especially the oligomers, penetrate tissue slowly. A rat's brain left overnight in a buffered glutaraldehyde
solution and sliced the next day shows a colour change and harder consistency to a depth of 2-3 mm. Objects
fixed for a few hours in glutaraldehyde are no longer osmotically responsive.
3. The free aldehyde groups introduced by glutaraldehyde fixation cause various problems. These include nonspecific binding of proteinaceous reagents, notably antibodies, and a direct-positive reaction with Schiff's
reagent). The free aldehydes must be removed or blocked by appropriate histochemical procedures, as described
in textbooks (Culling et al., 1985; Kiernan, 1999, Ruzin, 1999), before attempting immunohistochemistry, lectin
histochemistry, the Feulgen reaction of periodic acid-Schiff staining on glutaraldehyde-fixed material.
4. The thorough cross-linking of a glutaraldehyde-fixed specimen impedes the penetration of fairly large paraffin
wax molecules. This makes for difficult cutting and peculiar differential shrinkage artifacts within the specimen.
You can stain mitochondria nicely in cells surrounded by obviously abnormal spaces. This is an exaggeration of
the inadequacy of formaldehyde and osmium tetroxide as fixatives to precede paraffin, and it also highlights the
shortcomings of predominantly coagulant fixatives (AFA, Davidson's, Bouin etc), which preserve the microanatomy well but destroy or displace little things like organelles. Fortunately, plastic monomers penetrate
glutaraldehyde-fixed tissue adequately. It has been shown that they do not enter every crevice, but there is
enough support to allow the cutting of ultrathin sections for electron microscopy.
5. Immunohistochemistry, which requires as many intact amino acid side-chains as possible, is severely
impaired by glutaraldehyde fixation. Nevertheless, clever people have generated antibodies to individual amino
acids that are glutaraldehyde-bound to protein. These allow the detection of soluble amino acid neurotransmitters
such as glutamate, GABA and even glycine in presynaptic axon terminals in glutaraldehyde-perfused central
nervous tissue. Extensive cross-linking also results in the loss or severe reduction of most histochemically
demonstrable enzymatic activities, though several are retained after brief fixation.
Mixtures containing formaldehyde and glutaraldehyde
The combination of formaldehyde with glutaraldehyde as a fixative for electron microscopy takes advantage of
the rapid penetration of small HCHO molecules, which initiate the structural stabilization of the tissue. Rapid
and thorough cross-linking is brought about by the more slowly penetrating glutaraldehyde oligomers.
DEHYDRATION
Dehydration is the chemical removal of water from the specimen.
Common dehydrating fluids are ethanol and acetone. The potential problems of dehydration
are shrinkage of the specimen, plasmolysis, and removal of soluble components from the
specimen. Dehydration must be conducted relatively rapidly in order to prevent excessive
extraction of alcohol and acetone-soluble compounds, but slow enough to prevent
plasmolysis.
Extraction of specimen components is difficult to control. Low molecular weight
carbohydrates are particularly susceptible, since carbohydrates are usually poorly cross-linked
if at all following fixation. Proteins tend to be cross-linked by glutaraldehyde during primary
fixation and the lipids by osmium tetroxide during secondary fixation. The carbohydrates are
essentially unfixed. Linked to the problem of extraction is that of shrinkage. Both problems
are most serious at low concentrations in the dehydration series. In general, rapid dehydration
is best for these reasons.
By 70 % alcohol, the tissue no longer shrinks as much, but does begin to harden. In fact,
extended periods of dehydration in the higher concentrations of alcohol may make the tissue
quite brittle. If a stopping point is needed, most histologists choose 70 % to 100 % alcohol as
a good place to stop for the evening.
If there is evidence of plasmolysis, perhaps additional dehydration steps (and/or longer
changes) may be required. Cell membranes sometimes retain some osmotic activity after
short periods of fixation. Longer periods of fixation in glutaraldehyde can reduce osmotic
sensitivity as well. (Membranes are essentially insensitive to osmotic changes after 48 hours
of fixation in glutaraldehyde.) Poor fixation will aggrevate problems with dehydration.
Dehydration at refrigerator temperatures slows the process down a bit and tends to lend some
rigidity to the tissue. It may also reduce plasmolysis slightly. Plants are the most sensitive to
poor dehydration, and therefore, refrigerated dehydration is preferred for these tissues.
When changing solutions, make sure that the specimen does not dry out.
INFILTRATION
Infiltration is the replacement of the dehydrating fluid or transition solvent with plastic resin.
The goal of infiltration is simply the complete penetration of resin into the specimen.
EMBEDDING
Embedding involves the final positioning of the tissue specimen in the liquid plastic
(embedding medium, resin) and its subsequent polymerization.
Ideal qualities of embedding medium:
-solubility in dehydrating agents
-low viscosity as monomer for penetration
-uniform polymerization
-little volume change during polymerization
-good preservation of fine structure
-good sectioning quality that includes homogeneity, hardness, plasticity and elasticity
-resistance to heat generated by sectioning
-adequate specimen stainability
-stability in electron beam
-electron lucent
-easily available
-uniformity from one batch to another
The choice of a resin is determined by the type of the specimen and the objective of the study.
The materials most commonly used as embedments are 1. epoxy resins (e.g. Epon, Araldit)
and 2. acrylate-based resins (e.g. LR White, Lowicryls).
1. Epoxy resins, water immiscible embedding media, are ideal for structural studies or even
for high resolution electron microscopy (esp. Araldite). These resins are stable with respect to
light, heat and oxygen. Hardening is accomplished without significant shrinkage or uneven
polymerization. Sections of these resins are relatively stable under electron bombardment.
The remarkable stability is the primary reason for the excellent clarity of the picture obtained.
A minor disadvantage is a reduction in the contrast between sample and background.
2. The hydrophilic properties of acrylic resins provide two distinct advantages. During
dehydration and infiltration the specimens may be kept in partially hydrated state.
Polymerization of these media is started either by heat or when the need to preserve the
sample's antigenicity is paramount, then also by a low temperature UV cure. Lowicryls are
particularly useful for immunolabeling of sections because of improved preservation of
antigenicity and a significantly lower background labeling.
HM20 or HM23 (Lowicryl types) can be used to produce high contrast images of completely
unstained thin sections in the scanning transmission electron microscope by Z-contrast. HM20
and HM23 are particularly suitable for dark-field observation because of their low density
compared to conventional embedding media.
Progressive Lowering of Temperature Method of Dehydration & Embedding in
Lowicryl Resins (PLT)
The results obtained with Lowicryls clearly show the advantages of this approach to obtain
good structural preservation of cellular contents and ultrastructure. Furthermore, the PLT
method employs low temperature to reduce protein denaturation and to maintain a degree of
hydration, which may be important in preserving protein structural conformation. Specimens
suffer most during dehydration by organic solvents, mainly ethanol, whereas final infiltration
by resin monomers and polymerization seems to be less critical.
In order to minimize molecular thermal vibration, which can have adverse effects on
specimens weakly fixed with paraformaldehyde, one can dehydrate samples partially or
totally at low temperature. PLT technique that combines increasing solvent concentration with
decreasing temperature, after which infiltration and polymerization are carried out.
II. Physical (cryo) fixation:
Cryopreservation
One alternative to standard chemical fixation is the use of low-temperature methods otherwise
known as cryopreservation. In cryopreservation samples are rapidly frozen and then further
processed using a variety of techniques.
Essentially the same goals of standard fixation apply here namely to arrest cellular processes
rapidly and preserve the cell in as near to the living state as possible. We have a lot of
confidence that this is the case with cryopreservation as it has been shown that rapidly frozen
cells can remain viable following warming. Cryopreservation offers a number of advantages
over conventional fixation among these are:
1) Rapid arrest of cellular processes. One is not dependent on the speed of penetration of the
fixative. (milliseconds vs. seconds)
2) Avoidance of artifacts induced by changes in osmolarity, pH, or chemical imbalance.
3) Because cellular constituents are not subjected to biochemical alterations they remain in
more of their natural configuration. Labile components are retained and antigenicity is usually
improved.
4) Cells can be examined without introduction of other possible artifacts caused by
dehydration or embedding.
5) One can examine cellular domains that might otherwise be inaccessible (e.g. IMPs) or from
a view that is usually not possible (e.g. 3-D view via deep etch).
There are however a number of disadvantages as well and among these are:
1) The need for specialized freezing and processing equipment (-80oC freezer,
cryoultramicrotome, freeze fracture device, etc.)
2) Freeze damage due to poor freezing rates.
3) Limited view of specimen and or difficulty in manipulating the frozen material.
Rapid Freezing:
The major obstacle to good cryopreservation is the introduction of artifacts due to formation
of ice crystals that disrupt the cellular structure. The goal of rapid freezing is to prevent the
formation of ice crystals and preserve the aqueous component of the cell in near to the
vitreous state. Vitreous refers to glass or glass like, and just as glass is really a supercooled
liquid and not a solid, water can also exist in this quasi-solid state. In general this is very
difficult to accomplish with biological samples and usually we simply strive to keep ice
crystal formation to a minimum which is often defined as whether or not the crystals are
visible in the electron microscope. This cannot be accomplished by simply putting the sample
in the freezer.
Perhaps the most important aspect of rapid freezing is the choice of cryogen or freezing
medium. A good cryogen should have several properties.
1) Low freezing point - need to have a good thermal gradient between the sample and the
cryogen.
2) High boiling point - must minimize the formation of a vapor barrier near specimen due to
latent heat of sample. The formation of an insulating vapor barrier around the sample is
known as the leidenfrost or "bad frost" phenomenon and prevents the cryogen from making
direct contact with the surface of the sample. This tends to slow the freezing rate and produce
ice crystals.
3) It should have a high heat capacity and thermal conductivity (latent heat). In plain terms it
should be able to absorb heat without increasing its own temperature. Because of this low
molecular weight liquids such as N2 and He tend not to very good cryogens.
Cryogen melting pt. boiling pt.
Freon 22
-160
-40.8
Freon 13
-181
-81.1
Freon 12
-155
-29.8
isopentane
-160
27.85
propane
-189
-42
nitrogen
-209
-196
ethane
-183
-88.6
helium
-272 (1o K)
-268.9
An alternative to liquid cryogens is the use of a nitrogen slurry or slush. By lowering the
pressure of liquid nitrogen it can be induced to freeze and become a solid. When brought back
to room pressure the liquid and solid nitrogen exist side by side. Just as a glass of ice and
water remains at 4 degrees longer than does a glass of pure 4 degree water, the nitrogen slush
has a higher latent heat and can thus absorb more heat from the sample before boiling. This
reduces the leidenfrost effect and improves freezing rates.
The rate at which a specimen freezes is usually the determining factor in the amount of ice
crystal formation and subsequent damage there is. Slow freezing rates such as 1 C/min results
in significant damage. The extracellular water freezes first and pulls out the water from the
cell as the concentration gradient changes. In general cells do not contain large amounts of
unbound water so the formation of very large ice crystals usually does not happen but the
specimen can become shrunken and distorted.
Rapid freezing is usually defined as a change in temperature in excess of 10,000 C/sec. (vs. 1
C/min). One of the major problems associated with rapid freezing is the total amount of heat
that must removed from the specimen. If internal heat from the specimen continues to warm
those portions that are cooling it will prevent the water from undergoing a rapid phase change
and large ice crystals can form. For this reason the size of the specimen should be kept to a
minimum regardless of the freezing method used and the specimen carrying device should be
made of a small amount of material that has excellent thermal conductivity. Thin pieces of
copper or gold are usually used.
Rapid Freezing Methods
There are seven main rapid freezing methods presently available. They are
1. immersion freezing - the specimen is plunged into the cryogen.
2. slam (or metal mirror) freezing - the specimen is impacted onto a polished metal
surface cooled with liquid nitrogen or helium.
3. cold block freezing - two cold, polished metal blocks attached to the jaws of a pair of
pliers squeeze-freeze the specimen.
4. spray freezing - a fine spray of sample in liquid suspension is shot into the cryogen
(usually liquid propane).
5. jet freezing - a jet of liquid cryogen is sprayed onto the specimen.
6. high pressure freezing - freezing the specimen at high pressure to subcool the water.
7. excision freezing - a cold needle is plunged into the specimen, simultaneously freezing
and dissecting the sample.
Mostly used rapid freezing methods depending on sample size:

PLUNGE FREEZING - Only suspensions (< 1 μm) or thin tissues containing antifreeze. In this practical, the method will be shortly presented.

SLAM FREEZING - Suspensions and thin tissues (few μm, only front well frozen ca.
1 μm)

JET FREEZING (JFD) - Adequate freezing of suspensions not thicker than 15 μm.
Thicker specimen require anti-freeze

HIGH PRESSURE FREEZING (HPF) - Freezing under high pressure (2000 bar).
Adequate freezing of samples up to 200 μm thickness without anti-freeze. In this
practical, the HPF method will be shortly presented.
In details:
Specimens are rapidly placed or "plunged" into the cryogen and held there for 20 - 30 seconds. It is important
that the specimen be as small as possible as good freezing will only occur on the outer surface and one wants to
reduce the heat load placed on the cryogen. Plunge freezing is best used on very small specimens or cell
suspensions.
One problem associated with plunge freezing is the fact that as the cryogen removes heat from the specimen it
begins to warm up. This is a localized effect but results in either a decrease in the thermal gradient between the
cryogen and specimen or even worse in the formation of leidenfrost. To avoid this it is desirable to have a fresh
supply of cryogen constantly moving over the sample and taking away any excess heat with it. This can be done
by either moving the sample rapidly through the cryogen (projectile freezing) or moving the cryogen past a
stationary specimen. This is the theory behind jet freezing. The most commonly used cryogen for jet freezing is
liquid propane and the device is known as a propane jet freezer. Basically the unit operates by putting the
specimen on a very thin support foil or holder and then placing it between two thin pipe with opposing ports.
Liquid propane (which was liquefied by a bath of liquid nitrogen) is stored in a bomb underneath the output ports
and is then forced out from the ports under great pressure by introducing dry nitrogen to the the propane bomb.
Two opposing streams of liquid propane the hit the specimen from both sides and carry away the excess heat.
Cooling rates of 30,000 C/sec have been claimed for propane jet freezing and heat exchange is 2 - 30 times faster
than with plunge freezing alone. These are dangerous to use and we are experimenting now with a device I
helped to design which uses six ports (3 above, 3 below) that uses a stream of liquid nitrogen.
A second alternative to rapid freezing samples with liquids is to bring them in rapid contact with a very cold
surface. Although this will result in severe ice damage in the sample that is not immediately in contact with the
surface, it can produce excellent results in the region immediately adjacent to the surface. Contact freezing is
accomplished by pre-cooling a large metal block (usually polished copper, brass, or gold) and then rapidly
bringing the sample in contact with the block. Because latent heat and leidenfrost is not a concern in this method
one simply wants to create the largest thermal gradient possible. For this reason liquid nitrogen or even better
liquid helium is used. The primary reason that most researchers choose to use liquid nitrogen is that it costs
approximately 45 cents per liter whereas liquid helium costs $200 per liter.
One problem with bringing the sample in contact with the block is the possibility that it will bounce and thus
damage the specimen. For this reason a special freeze slamming device is used that has a glycerol hydraulic
damping system to drop the specimen onto the block but prevent it from bouncing. A modification of this
procedure involves grabbing the specimen between two precooled metal surfaces. These cryopliers are widely
used in cryopreservation of specimens such as muscle fibers.
A modification of surface freezing is known as spray freezing. In spray freezing the sample in the form of a
suspension is spray or atomized onto a precooled metal block or into a cryogen. This avoids the problems of
bouncing and keeps specimen size to a minimum (1 ul or less volume). It has the disadvantage that the specimen
must be one that can be sprayed and is often difficult to handle afterwards as it must be collected without
rewarming the sample.
The latest in freezing devices is known as a high pressure freezer. At extreme pressures of 2100 bar (Bar = 1
ATM = 760 mm Hg) the nucleation of ice is significantly reduced. A second thing that happens is that the
melting point of water is lowered to - 22C (vs. 0 C at 1 ATM). This is one reason that cold water on the ocean
bottom does not freeze. At these pressures the critical cooling rate is raised to 100 C/sec (vs. 10,000 C/sec at 1
ATM). The device works by initially pressurizing the chamber with isopropanol followed by liquid nitrogen.
Because the cells are pressurized for only a few milliseconds before the LN2 is introduced they are generally not
harmed too much. LN2 can be used because at these pressures it will not boil so no leidenfrost is formed.
Cryoprotectants
Regardless of the freezing method used many specimens are treated with a cryoprotectant to
reduce the possibility of ice damage. Cryoprotectants function by both increasing the number
of ice nuclei and retarding the growth of ice crystals. By either binding to water molecules or
substituting for water molecules cryoprotectants reduce the number of water molecules
available for binding to growing ice nuclei and thus greatly slow the growth of these crystals.
Generally cryoprotectants are viscous and in this way they also slow down the rate of
diffusion of water from the specimen as the exterior water freezes. This helps to reduce the
shrinkage effects of slow freezing. Some commonly used cryoprotectants are glycerol
(penetrating type) or sucrose (non- penetrating type) and are generally used in concentrations
of 10-30%. One of the disadvantages of cryoprotectants is that it has been shown that
extensive exposure to cryoprotection can alter the internal structure by applying osmotic
pressure to the cytoplasm. Usually marine organisms have a number of dissolved salts in the
medium which act as cryoprotectants and often these can be frozen without further
cryoprotection.
What to do with rapidly frozen samples
1. One of the things that can be done with rapidly frozen samples is to replace the
aqueous component of the specimen with an organic solvent without allowing the to
change from its frozen arrested state. During the freeze subsitution process a rapidly
frozen sample is held for one to two days in a vial of organic solvent at -80 C. Over
this time period the frozen water molecules are replaced or "substituted" by molecules
of the organic solvent. This happens despite the fact that the water is never allowed to
return to the liquid state. Acetone is usually the solvent of choice although ethanol and
methanol have been used as well. The organic solvents have some fixitive properties
of their own which can be enhanced by the addition of standard fixitives such as
osmium tetroxide. Recently anhydrous glutaraldehyde has become available for use in
organic solvents during freeze substitution. Thus the cells are chemically cross linked
and fixed before their components have an opportunity to change from their frozen
positions. The samples are then gradually brought to room temperature (done slowly
to prevent renucleation of ice crystals), the fixitive, if any, rinsed out with pure organic
solvent, and infiltrated and embedded as usual. Thus in freeze substitution the fixation
and dehydration steps are combined into a single step.
High pressure freezing is mostly followed by FREEZE SUBSTITUTION and embedding
20C
SUBSTITUCE
FIXACE
IMPREGNACE
POLYMERIZACE
-50C
-90C
Aceton+0,1%UA
Ac
Ac/HM20
HM20
One great advantage of rapid freezing and freeze substitution as oppposed to standard
chemical fixation is that many of the artifacts associated with chemical fixation can be
eliminated or greatly reduced. A prime example of this is in the study of membranes
and membrane bound organelles. The length of time a fixitive takes to penetrate a cell
and the changes it induces in terms of periability often results in shrinkage or
wrinkling of membranes and membrane bound organelles. If one compares these to
chemically prepared cells the smoothness and roundness of freeze substituted material
is quite surprising. Also rapid cellular processes such as the fusion of membrane
bound vesicles can be captured because although the fusion process itself is very rapid,
the freezing rate is even faster.
A second great advantage of freeze substitution is seen when one uses the fixation properties
of the organic solvent alone to preserve the cell. This has the great advantage of hlding all
cellular components in place while at the same time not cross linking the cell so completely
that not cytochemistry can be done. In fact cells preserved in this way have better
ultrastructural preservation and greater ability to react in cytochemical treatments than any
other method. A variety of methacrylate resins have been developed which facilitate
immunocytochemical processing of cells including Lowicryl which remains a liquid down to 40 C and can be polymerized at that temperature using U.V. light. Thus cells are freeze
substituted, infiltrated, and polymerized without ever regaining the unfrozen state. Cell
structures and biochemicals can therefore be preserved in nearly their native state.
2. Slam, jet and plunge freezing are mostly followed immediately by cryoelectron microscopy.
High-resolution observation of small isolated proteins (~20-100 nm in diameter) or viruses in
close-to-native state requires specialized approach named Single Particle Analysis. Such
proteins have to be observed only in electron microscope whose electron beam has brutally
damaging impact on the delicate protein structure. Therefore the dose of electrons must be
kept small andaveraging of many identical particles present in the specimen has to be
introduced. The result is a three-dimensional structure of the studied molecule (protein, virus,
enzyme...).
3. The most difficult of all cryo combinations is CEMOVIS. CEMOVIS is Cryo-Electron
Microscopy of Vitrified Sections. After high pressure freezing (mostly) the sample is sectioned
in the frozen, hydrated state and the sections are observed at - 180°C.
STAINING OF SECTIONS
Double possitive staining with uranyl acetate and lead citrate
The heavy metal salts used as stains in the electron microscope consist of ions of a high
atomic number with a large number of protons and electrons that scatter the beam electrons.
The two most commonly used positive stains are uranyl acetate (MW 422) and lead citrate
(MW 1054). It is known that uranyl ions react strongly with phosphate and amino groups so
that nucleic acids and certain proteins are highly stained. With lead stains, it is thought that
lead ions bind to negatively charged components such as hydroxyl groups and osmiumreacted areas. Uranyl and lead stains are termed general or nonspecific stains when used in the
routine manner since they will stain many different cellular components.
The simplest method used for staining the grids with uranyl acetate and lead citrate is as
follows.
Several individual drops of a saturated solution of uranyl acetate are placed on clean parafilm
or sheet of dental wax in a Petri dish. The grid with mounted sections is floated (section side
down) on a drop of the uranyl acetate for 5-15 min. The grid is rinsed twice in boiled distilled
water and kept in a covered Petri dish for subsequent staining with lead. It is recomended that
the grid not be allowed to dry before staining with lead.
Because CO2 in the air is the primary source of lead precipitation, a CO2 free chamber is
prepared by using several NaOH pellets. The NaOH rapidly absorbs CO2 in the covered dish,
providing a CO2-free staining chamber. The preparation of this setup should be done before
the grids are stained with uranyl acetate in a second dish, so that by the time the first staining
is completed. The atmosphere in the chamber will be free of CO2 and ready foe lead staining.
The gris readily stained with uranyl acetate and wetted with boiled distilled water is
immediately placed (section side down) on the drop of lead citrate and after 1 minute rinsed
with boiled distilled water.
Precautions to minimize section contamination
1 Maintain a CO2 free environment during lead staining. Avoid breathing on sections during
staining.
2 Eliminate the air-stain interface during lead staining.
3 If the white precipitate (PbCO3) is visible in the bottle containing the lead solution, discard
the solution instead of filtering it. However, this precipitate can be sedimented by
centrifugation
4 Use clean boiled distilled water for preparing lead and rinsing solutions
5 Avoid staining longer than necessary
6 Do not allow the grids dry between various staining steps
Negative staining
Negative staining is the method of choice, when single particles (such as, protein complexes,
viruses, nucleoprotein complexes etc) are to be analyzed by transmission electron microscopy.
The preparation of the sample is simple and fast. Several techniques exist. The most often
used is the deposition of the solution containing the sample on the EM grid covered with
carbon or formvar/carbon film, previously made hydrophilic by glow discharge. The particles
adsorb on the film. The grid is than placed on the staining solution (2% uranyle acetate in
water) for several seconds and carefully dried. The deposition of the stain on the structure of
the studied object will produce the contrast necessary to see them in TEM. The method will
be demonstrated using TMV (tobacco mosaic virus).
Tokuyasu method
CHEMICAL FIXATION followed by CRYOPROTECTION AND FREEZING
1. Transfer cubes of paraformaldehyd fixed tissue or cell pellets to 1 ml of 2.3M sucrose in an
Eppendorf tube, seal the cap and leave overnight on a rotating wheel at 4°C. The sucrose
solution will infiltrate the specimen.
2. Transfer each infiltrated specimen onto a clean specimen stub, remove excess sucrose with
a damp filter paper and immerse the specimen stub and specimen in liquid nitrogen until
freezing is complete.
SPECIMEN TRANSFER TO PRE-COOLED CRYOCHAMBER
1. Pre-cool the cryochamber of the ultramicrotome to –80°C, TRIMMING
2. CRYOSECTIONING, -120°C, dry knife
IMMUNOLABELLING ON ICE
CONTRASTING AND DRYING IMMUNOLABELLED CRYOSECTIONS
Advatage: Avoiding of dehydration and resin.
Electron tomography
In general, tomography is a method for acquisition of three-dimensional mass distribution of
the observed object. Each image acquired by a transmission electron microscope (TEM) is a
two-dimensional projection of the observed object, therefore (based on mathematical
approaches) it is possible to obtain the 3D structure of the object by its tilting in the TEM
followed by combining of the acquired projections. The resolution of the 3D structures in
biologic applications amounts to several nanometers.
In this practical, the whole process will be presented in detail. We will start by analogy of
electron tomography and computer tomography used in medicine and explain the principle of
projection formation of each type. Further the principle of reconstruction and the effect of
limited range of tilting will be illustrated and the first part of the practical will be closed by
examples of reconstructed volumes.
The second part will take place at the electron microscope. It is the FEI Tecnai Sphera 20,
with up to 200 kV acceleration voltage, equipped with LaB6 gun, specimen tilting stage and a
CCD camera. The operator will show you the specimen handling, the kinds of specimen
holders that we use and explain the principle and composition of the electron microscope. At
the end, a small series of tilted images will be acquired.
The last part will be dedicated to the practical process of 3D reconstruction. You will follow
the progress on the graphical interface of the reconstruction programme. Finally, the
interpretation of the reconstruction will be discussed.
Based on:
http://www.hei.org/research/aemi/emt.htm
http://www.liv.ac.uk/emunit/fixation.html
www.ou.edu/research/electron/bmz5364/embedding-media.html
http://www.springerprotocols.com/Abstract/doi/10.1385/1-59259-201-5:111
http://www.emsdiasum.com/microscopy/technical/datasheet/14330.aspx
Electron Microscopy Methods and Protocols
http://www.uga.edu/caur/temnote2.htm
http://www.protocol-online.org/cgi-bin/prot/view_cache.cgi?ID=2680
http://www.bristol.ac.uk/vetpath/cpl/emtechs.htm
E.M. PROCESSING SCHEDULE - ACRYLIC RESIN (for immunogold labelling)
Cells are grown on cover slips as a monolayer in Petri dish
1. Fix tissue the cells by adding of double strength fixative (8% formaldehyde/0,2%
glutaraldehyde) to the cell culture (1:1 ratio)
2. Wash
PBS buffer
3 x 5´
3. Dehydration
30% ethanol
1 x 1´
50% ethanol
2 x 5´
70% ethanol
2 x 5´
90% ethanol
2 x 5´
100% ethanol
2 x 10´
(transfer cover slips into glass P.dishes)
4. Impregnation
LR White resin
30´
LR White resin
overnight
4°C
5. Embedding
FLAT - cover slips (cells side-down) on the top of
gelatin / beem capsules filled with LR White resin
Cell pellets or tissues
embed in gelatine capsules with fresh resin
6. Polymerization
at 60oC
24 hours
________________________________________________________________________
IMMUNOGOLD LABELLING PROTOCOL
1. 5% NGS (Normal Goat Serum)
(Practical course)
20μl drop
5´ RT
wet chamber
Masking of non-specific binding sites by floating the grids, section-side down on small drops
of blocking agent on parafilm.
2. pAb in BSA/PBS/T20
10μl drop
1 – 2 hours RT
wet chamber
Dilute the primary antibody (pAb) in the blocking buffer (0.1 % BSA / 0.05% Tween 20 /
1xPBS). Antibody concentrations in the microgram range (1-5 μg/ml) are usual. Put small
drops of diluted antibody onto a clean parafilm surface in a wet chamber. Transfer the grids
and float on the antibody drops.
NGS
20μl
pAb
10μl
3. 1xPBS (wash)
3 x 5´
on parafilm
5´ RT
wet chamber
Wash with three changes of 1xPBS.
4. 5% NGS (Normal Goat Serum)
20μl drop
Transfer the grids, section-side down, to drops of NGS on parafilm, in a wet chamber.
5. sAb in BSA/PBS
10μl drop
25´ RT
wet chamber
Incubate with immunogold conjugated secondary antibody appropriately diluted blocking
buffer (0.1%BSA / 1xPBS).
6. 1xPBS (wash)
3 x 5´
on parafilm
2 x 3´
on parafilm
Wash with three changes of 1xPBS
7. dH2O (wash)
Final wash 2x 3min with distilled water.