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Sample Preparation Into Ultra-thin Sections.
Contents
Introduction
Tissue selection
Fixation
Glutaraldehyde
Osmium tetroxide OsO4
Potassium permanganate - KMnO 4
Acroline
Embedding
Dehydration
Resin infiltration
Polymerization
Ultramicrotomy
Trimming the capsule
Making glass knives
Thin sectioning
Staining sections
Miscellaneous procedures
Thick sections - light microscopy
Whole mounts on coated grids
Arts and Formulae
Photography
Developing film
Developing prints
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Introduction.
The Electron Microscope was developed from the coalescence of
several scattered ideas and hypotheses. The electron itself was discovered
at about 1897 and was shown to have wave properties in 1924. Much of
the early work with electrons centered on how to generate electrons and
how to deflect or aim them. Various "lens" were contrived and by 1931,
two German scientists were demonstrating the "first" EM which was largely
a modified oscillograph. No specimens could be viewed but it was
predicted that an electron microscope would have much better resolution
than the light microscope. By the late 1930's and early 1940, commercial
EMs were available with moderate resolution. These were used mainly to
study electron optics and to make improvements. Procedures for specimen
preparation came much later.
For the biologist, the EM is one of the most powerful tools available
for cell and tissue studies. It should be realized that electron microscopy is
not a science on its own, but merely a technique or tool. A thorough
understanding of the EM and of cells and their ultrastructures are an asset
to understanding most other areas of biology.
There are now many types of electron microscopes, but the two most
common types are the transmission electron microscope (TEM or just EM)
and the scanning electron microscope (SEM). Simply put, the SEM scans
the surface of coated specimens with an electron beam and by detecting
electrons scattered (reflected) by the object, forms an image on a TV like
monitor. This image is usually aesthetically pleasing and has a resolution
of 50 µ and up. The TEM transmits a beam of electrons through a
specimen and forms an image based on the removal of electrons from the
beam by the specimen (basically a high resolution shadow). Resolution can
attain <1 angstrom on research grade TEMs and the variety of specimen
preparations allows much greater versatility over the SEM. In essence,
you can not only view the surface of specimens (bacteria, viruses,
molecules, etc.) but you can also look inside the specimens using thin
sectioning techniques. This coupled with cytochemical, isotope, and
immunochemical techniques permits a process oriented study of biological
systems.
The TEM usually generates electrons by saturating a tungsten
filament with current such that electrons cascade from it (much like the
electron gun of a black and white television). The filament and the rest of
the microscope column are under a high vacuum so as to prevent oxidation
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of the tungsten filament (what happens if you crack the globe of an
incandescent light bulb?). This vacuum is usually in the range of 10-6 to
10 -7 µ of mercury and is achieved using a "diffusion" pump. For all
practical purposes, living tissues or cells cannot be viewed under the TEM
since the specimens are subjected to such high vacuum, heat and intense
radiation from the electron beam. This would suffice to kill the cells either
by volatilization of water (and other low melting point substances),
denaturation from heat, or ionizing radiation. In addition, the radiation
undoubtedly causes many chemical changes to occur. Polymers often
become insoluble and sublimation may occur.
The most popular and useful procedure used with EM is the thin
sectioning technique. This technique can be broken down into the
following sections and each will be dealt with separately.
1.
Tissue isolation
2.
Fixation with glutaraldehyde, OsO 4 and occasionally KMnO 4
3.
Embedding in a plastic resin
4.
Ultramicrotomy
5.
Post-staining of thin sections
TISSUE SAMPLES:
Tissues must be killed and fixed in a way to stabilize their structures
in the EM environment and to accurately reflect the true structure of the
tissue. The time between isolation of the tissue (e.g., dissection) and
addition of fixative should be minimized so as to avoid post-mortem
changes. The tissue size should be kept small. Blocks should be cut to less
than 1mm 3 in order to ensure thorough and quick penetration of fixatives
and embedding solutions. Such size concerns are not pertinent to cell
suspensions. It is usual practice to suspend the cells in the preparative
solutions and to pellet the cells by centrifugation between steps. However,
if a tissue block is too large, fixatives and embedding resins will usually
not penetrate to the middle. Some tissues of low density (e.g., lung, root
tips) are the exception. In the dense tissues, one can often note a "halo"
effect.
3
Cut in half
HALO EFFECT
Dark with
Osmium
Pale or White
No Osmium
TISSUE BLOCK
Animal tissues pose a few unique problems for optimum preparation.
Lacking rigid cell walls, the tissue is often limp and difficult to cut into
small blocks (e.g., liver). This can be overcome by allowing larger pieces to
be fixed for a short period of time in the primary fixative (glutaraldehyde,
10-15 minutes). The tissue will then be somewhat more firm and easily
cut into smaller pieces.
Some tissues are hard and mineralized (e.g., bone) and must be
demineralized with chelating or acidic solutions. This is fairly rare
however, and tissues such as hair and nails can be prepared without
special treatment. Some animal tissues are fairly dense and longer
treatment times may be necessary. This determination is made by
experience and/or trial and error.
Plant tissues also present some unique problems during preparation.
There is usually a higher water content with most mature plant cells
having a large central vacuole. This necessitates attention to the
dehydration step of embedding. Another problem often encountered is the
presence of wax on certain plant surfaces which may retard the
penetration of fixatives and embedding solution. It may also cause the
separation of the tissue from the surrounding plastic upon trimming and
sectioning. The waxes (cutin and suberin) may be partially removed with
organic solvents (acetone, ether) prior to processing. It is not necessary to
remove all wax in that the solvents tend to make the waxy surface less of
a barrier and more tenacious to the plastic (similar to cleaning and
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abrading a surface before painting or gluing).
problems since they are hydrophilic gels.
Cell walls pose no special
FIXATION
As mentioned earlier, living tissues cannot be viewed with the
Electron Microscope. The goal of specimen preparation is to preserve the
tissue in a form which hopefully represents its' natural, in vivo form.
Fixation serves to "kill" the cells, and to stabilize and preserve them and
their structures during subsequent preparation steps. Throughout the last
three decades, several chemical fixatives have been studies for use in
specimen preparation. Out of this, two fixatives have emerged as virtually
"universal" and will be described here.
Glutaraldehyde - a five carbon structure
with an active aldehyde on each end.
GLUTARALDEHYDE
Glutaraldehyde is referred to as a bifunctional fixative due to the two terminal
aldehydes and is usually used as the first of two fixatives. It is fairly
stable in concentrated form and at cold temperatures (-20°C). At room
temperature and especially when diluted to working strength (1-3%), it is
unstable and impurities and polymers accumulate. Oxidation to glutaric
acid is the usual consequence but this reaction is inhibited by low pH
which is produced by the oxidation. Thus, it is somewhat of a self-limiting
process, but also explains why buffered (pH 7) solutions are so unstable.
The stock solution of glutaraldehyde (even when purchased new) should
be checked often for impurities. This is easily done spectrophotometrically
with a 0.5-1% solution. The aldehyde has an absorption maximum of about
280 nm while impurities absorb maximally at 235 nm. The impurity peak
should be half the height of the aldehyde peak. The pH of the stock
solution should be above 3.5 as well. If impurities have accumulated to
unacceptable levels, the glutaraldehyde can be easily redistilled in a
vented fume hood with the distillate collected at 100O C in small fractions
until the pH of the distillate is less than 4.0. It is then convenient to freeze
several aliquots at -20°C at which it is chemically stable for many months.
Glutaraldehyde is relatively safe to use. Avoid skin and eye contact
and never pipette by mouth. Use in a well ventilated area since strong
fumes can irritate and mildly fix the epithelial lining of the
nasopharyngeal tissues.
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Fixation and stabilization are due to the cross linking of structures
that are reactive with aldehydes. Structures that are composed of proteins
(enzymes, etc.), glycoproteins, nucleoproteins, lipoproteins, glycogen, and
starch (occasionally) will react with the aldehydes. Most structures within
the cell have these components. Membranes loose their fluidity and
usually become very permeable. Structures that do not react with
glutaraldehyde will then tend to diffuse out of the cell. It is important to
follow this fixation step (often called a pre-fixation) with a second step
(often called post-fixation) using another fixative (see next).
Glutaraldehyde fixation does not cause significant shrinkage and can be
carried out at room temperature. Cold temperatures cause much of the
cytoskeleton of the cell to "dissolve" or disassemble, thus altering the
ultrastructural representation. A 1-3% buffered solution (pH 7) is
recommended and fixation should not exceed 60 minutes unless the tissue
is naturally dense or impermeable (e.g., some insects and plant tissues).
The choice of buffer is important. Veronal buffers (containing barbitals)
should not be used with aldehyde fixatives and phosphate buffering may
form a precipitate in the presence of calcium and uranyl ions. If the
specimen is known to contain these ions, use a different buffer (e.g., Tris,
Hepes, cacodylate). Also, the use of phosphate buffers with the
glutaraldehyde fixative occasionally causes a precipitin to form during the
second fixation step with osmium tetroxide. To prevent this, wash the
specimen with saline or water after the first fixation so as to remove all
traces of the phosphate. This problem rarely arises however and its cause
is not understood.
Osmium tetroxide - OsO4
Note: Osmium is extremely dangerous, the crystals, liquid
and vapors are all hazardous. The vapors can fix the cornea and lens
of the eye and both vapors and liquid are absorbed rapidly and act as a
nerve gas (it was in fact used as this in the World Wars) and attacks the
CNS. Use only with proper ventilation (e.g., fume hood). It
usually is purchased in crystalline form in preweighed sealed glass
ampoules. Always prepare the fixative solution in a fume hood. Report
any and all accidents immediately.
Usually a 1-2% buffered solution (see appendix) is used as the second
fixative and the fixation should be complete in 60-90 minutes since the
osmium molecule is small and penetrates rapidly. Most workers agree that
osmium works by saturating double (or triple) bonds since it is such a
strong oxidizing agent. For this reason, it is deposited at lipoidal sites quite
6
heavily thus causing them to look dark. Thus, membranes and lipid
droplets usually "stain" darkly with the osmium. It should be noted also
that since osmium is used as an aqueous solution, it has a tendency to not
reach the middle hydrophobic region of some membranes.
This gives a
tri-layered appearance (dark-light-dark) to these membranes and is easily
seen at higher magnifications. The tissue specimen will begin to turn black
almost immediately upon the addition of the osmium. The more dense the
tissue, the darker it will appear. (That's how you will know if you get it on
you, your skin, clothes, etc. will turn black. If you do spill some on
yourself, don't panic, wash it off with lots of water and immediately notify
your instructor or health office). The tissue often becomes brittle when
over fixed in osmium. It is advisable to use this fixation at refrigeration
temperatures so as to decrease the volatility of the solution. The unused
solution should be stored at 4°C in a scrupulously clean, foil wrapped 50ml
volumetric flask that is tightly corked (do not use ground glass or rubber
stoppers - they leak). The long narrow neck of the flask retards
evaporation of the fixative and offers a "handle" to the user. Use a long
tipped Pasteur pipette, taking care not to draw the osmium up into the
pipetting bulb (bippy). The flask should be stored in a refrigerator, and be
sure it is well stoppered. If it leaks, the interior of the refrigerator will
gradually turn black and there is a possibility that the osmium vapors can
accumulate in the confined space to dangerous levels. The osmium
solution will appear purple or violet in color when degraded or
"exhausted". It should be carefully pipetted into a flask containing 95%
ethanol. This will degrade the osmium for later disposal. Keep this waste
flask in the fume hood at all times.
Potassium permanganate - KMnO 4
Occasionally, an investigation may center on the study of
membranous structures and the cytoplasmic matrix is not of interest. In
this circumstance, a 1-2% buffered solution of KMnO4 can be used as the
sole fixative or in tandem with OsO4 (wash in between the two - they react
together). This is a rapid process and KMnO4 may also be used as a post
stain to enhance contrast from the Glutaraldehyde/OsO 4 preparation. This
is usually not necessary however.
Most non-membranous structures are washed away in subsequent
embedding steps. The time required for the fixation is usually 30 minutes
and the KMnO 4 kills the cells quickly, being a strong oxidizing agent. It
will permanently stain skin and clothing but it is not as hazardous as the
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other fixatives. In
skin. That doesn't
pipetted by mouth.
the same structures
fact, it is often used to treat fungal infections of the
mean that you can drink it however, so it should not be
It is chemically stable when kept at 4°C. It binds to
as does OsO 4 but is not as electron dense.
Acroline
A very toxic, flammable and volatile substance, it penetrates very
rapidly and is thus good for fixing large and dense tissues. It is not
thought of as a "common" fixative since it is potentially hazardous to use.
In addition, it does not fix lipids (in fact it dissolves them), denatures most
enzymes to inactive form and does not preserve the cytoskeletal network
very well. If it is deemed absolutely necessary to use this aldehyde, best
results are obtained by combining it with other aldehydes and following
with osmium post-fixation. Partially degraded and polymerized solutions
appear to be as effective as fresh or redistilled acroline. Use extreme care
when handling acroline. It is included here as a precaution to those who
choose to use it and to those who read reference to it.
EMBEDDING
The fixatives as well as most cellular components are aqueous. The
plastic resins that are used for embedding tissues are not miscible with
water. Thus, an intermediate solvent that is miscible with both water
and plastic resin is needed. Although there are many to choose from,
acetone is probably the best. Some microscopists prefer ethanol, often out
of habit from light microscopic procedures. Ethanol reacts with unbound
O s O 4 to form a fine dense precipitate thus extensive washing after osmium
fixation is needed. Acetone, which does not react, requires only minimal
washing to remove the osmium and buffer salts. Methanol is less reactive
than ethanol but has no advantage over acetone. In essence, water is
replaced by solvent and solvent will be replaced with plastic.
DEHYDRATION
Use glass vials (or centrifuge tubes) since acetone can dissolve many
plastics. Although a graded series of acetone solutions is commonly used it is unnecessary. A three step dehydration process is adequate, -50%,
95%, and two changes of 100% acetone (re-distilled - stored with molecular
drying sieves, see appendix). It is essential that all water diffuses out of
the tissue, otherwise holes will be created in the sections when viewed
with the EM since residual water will vaporize under the extreme vacuum.
8
Two changes of 100% acetone are a precautionary measure. As water
diffuses into the first 100% acetone, it is no longer absolute. A second
change dilutes out the water molecules even more. Use a Pasteur pipette
to add and remove the acetone solutions to and from your specimen vial
(as opposed to transferring the tissue block to a new vial with the next
solution).
Dehydration may cause some changes in the secondary and tertiary
structures of macromolecules and usually causes some shrinkage of the
tissue. The shrinkage is usually proportional to the water content of the
specimen. Fixation lessens this effect. Do not let the tissue dry in air.
Make transfers rapidly but neatly. During dehydration as during fixation
and embedding steps, keep the vials capped. 100% acetone is hydroscopic
and will absorb water from the air. Acetone will often dissolve unfixed or
poorly fixed components of cells such as saturated lipids (which do not
react with osmium) and chlorophyll and other lipoidal membrane
components. Starch is difficult to fix but is often so highly polymerized
and cross-linked in vivo that it is often "naturally fixed". However, it will
occasionally be leached out during dehydration.
RESIN INFILTRATION
There are many types of plastic resins available for embedding
tissue, each having attributes. The three principle types of resins used are
the epoxy resins, polyester resins and methacrylate resins. The most
commonly used resins are the epoxides Epon (Epon is no longer made but
other similar resins are available with similar names e.g., Epox) and
Araldite. They have adequate viscosity, are fairly stable under the intense
electron beam, and are of very fine grain. The purpose of the embedding
medium is to provide a stable, hard matrix throughout a tissue or cell in
order that very thin sections may be cut, usually on the order of 400-800
A.
Wax such as the light microscopist paraffin is not firm enough for such
thinness and it will melt under the electron beam. Epon and Araldite are
both epoxide resins and when polymerized are virtually indestructible and
insoluble (as are tissues embedded in them). Remember that water is
replaced with acetone and acetone is replaced by plastic.
Note - Epoxys can be irritating to skin and eyes; use with caution. Many
epoxides are known to be carcinogenic. The polymerized capsule however
is not carcinogenic. Use acetone on a cloth or wipe to remove any resins
from your skin. For eye contact - flush with warm (not hot) water.
9
Do not pipette resins. They are viscous enough that pipetting is
inaccurate. Simply pour the components into a 50ml disposable beaker
according to the formula given. The plastic mixture to be used contains
both epon and araldite along with DDSA (Dodecenyl succinic anhydride)
and NMA (nadic methyl anhydride) which are curing agents, and increased
amount of DDSA will result in softer plastic when polymerized. A
polymerizing catalyst or accelerator, DMP-30 (dimethyl aminomethyl
phenol) is used to speed the polymerization process.
A four (4) step series of plastic concentrations is used for the
embedding process:
1.
2.
3.
4.
3 part acetone - 1 part plastic mixture (w/o DMP-30)
1 part acetone - 1 part plastic mixture (w/o DMP-30)
1 part acetone - 3 parts plastic mixture (add 4 drops DMP-30
for every 10ml used
Pure plastic mixture with DMP-30 (two changes)
The plastic mixture is:
Total =
16 ml of a mixture of 5 parts Epon and 3 parts Araldite (506)
9 ml NMA
10 ml DDSA
35 ml
- stir exhaustively
For pure plastic mixture steps of embedding (step 4) add 15-17
drops of DMP-30 to the above mixture using a disposable Pasteur pipette.
DMP-30 is kept refrigerated. Allow it to warm to room temperature prior
to use to avoid condensation of water into the bottle. Epoxy resins,
especially araldite are also somewhat hydroscopic. Be sure to stir the
mixture well. It will quickly turn amber in color but will lighten to nearly
colorless during polymerization if done slowly. Two changes of the pure
plastic mixture with DMP-30 is recommended in order to ensure that all of
the acetone is removed (i.e., replaced) in the tissue. Since the pure plastic
mixture is viscous it is easier to remove the tissue block with a hooked
needle (dissecting) and place it in a fresh vial containing the pure plastic.
This will minimize the carryover of any acetone that had diffused into the
first pure plastic step. Keep the vials capped during embedding.
The times for each step will vary depending upon the density and
permeability of the tissue. For most tissues a 30 minute period for each
step is adequate. It is better to have each step longer than necessary than
shorter than necessary - residual acetone can result in poor sections.
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For the final step, use the 00 size polyethylene capsule molds
available. Fill them to about 2mm from the top and place your tissue block
to the bottom of the mold. Place a small paper label (written in pencil)
around the top perimeter of the mold. For cell suspensions, pellet the cells
with centrifugation in the pure plastic mixture after the allotted time.
Using a Pasteur pipette, draw off the pellet as a cell slurry and place 2-3
drops of the slurry in the capsule molds. Layer pure plastic mixture over
this to within 2mm of the top and centrifuge the capsule mold in a tabletop clinical centrifuge on a setting of 5 or 6 until the cells are concentrated
at the tip of the mold. Then place a label around the top rim of the
capsule.
POLYMERIZATION:
Allow the tray(s) of capsules to stand (wrapped in aluminum foil)
overnight at room temperature and then place the capsules in an oven at
60°C for 2-3 days. If the correct amount of DMP-30 has been used, the
capsules should be adequately polymerized within 3 days and the plastic
will have lost most of the amber color. A good test for correct
polymerization is to try and dent one of the side ridges of the tip of the
capsule with a fingernail (after removing the capsule from the mold of
course). If there is an indentation from the fingernail the polymerization
at 60°C should continue until the capsule is hard enough to show no
indentations. To remove the capsule from the mold, carefully cut the mold
lengthwise with a razor blade and peel the cut edges from the top (not the
tip) of the capsule. The capsule can then be easily removed. If the side
facets near the tip show cracks and/or bulging, it usually indicates too
rapid polymerization.
OTHER RESINS:
Relatively recently, a new monomeric resin called LR-White has been
introduced from England. It is a single solution that is stable at 4°C and is
used with 4 to 6 changes after dehydration which must be carried out with
absolute ethanol. Acetone can not be used since it generates free radicals
which interfere with the polymerization reaction. Using ethanol
necessitates that excess osmium be thoroughly removed by washing. The
capsules may be polymerized by heating to 50°C overnight but
polyethylene capsule molds should not be used since they are permeable
to oxygen which also interferes with polymerization. The result will be
"tacky" capsules and this may be avoided by using gelatin capsules as a
11
mold. Their only drawback is that the ends are rounded and are more
difficult to trim for sectioning.
The LR-White resin is a general purpose embedding medium that can
be used for light microscopy preparations. Under the electron microscope,
the tissues have a tendency to look washed out or leached and they don't
take up the post-strains (e.g., lead citrate) as well as the epoxy resins
described above. Except for the convenience of not having to mix together
the plastic resin components, there is no overriding advantage apparent
for choosing LR-White. The same number of steps are needed for
adequate infiltration. It is not as irritating or toxic as the epoxides,
however, and this concern may merit its use for general studies or
teaching.
ULTRAMICROTOMY
Now that the messy part is over with, it is time to master the skills of
electron microscopy that require precision and perfection. Although it is
most convenient to hire technicians to do the microtomy and microscopy,
you will not have an adequate appreciation for the results unless it is
learned first hand and it is truly one of the few procedures that are most
easily learned correctly by doing them and making mistakes. This is due
to the large number of variables that affect the quality of the result, i.e.,
the micrograph. The steps needed to master this section include: trimming
the capsule so as to expose the tissue for proper sectioning, making glass
knife edges fitted with a water boat (for sections to float on when cut from
the tissue (capsule) "face", the actual sectioning using the ultramicrotome
and placing the sections on "grids".
TRIMMING THE CAPSULE
This is easier to demonstrate than to explain in written form. Excess
plastic surrounding the tissue must be trimmed away in a fashion that will
yield a square or rectangular section. The capsule mold produced a 1mm2
face on the tip (see figure). This must be trimmed to a pyramid where the
pyramid tip and sides are exposed tissue. The angle of the pyramid sides
(called facets) should be about 45°. Too steep of an angle will not allow
enough lateral support when sectioning while too flat (or low) of an angle
will cause the "face" being sectioned to enlarge too quickly during
sectioning. The tip of the pyramid may be a point (giving square sections)
or a ridge (giving rectangular sections).
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TRIMMING
Top view
Side view
Gives rectangular
Sections
gives square
sections
Trim the capsule while viewing under the dissecting microscope using old
glass knives or knives not suitable for sectioning. Use smooth slicing (not
chiseling) strokes that cut through the plastic in one stroke. Take very
thin slices so as to leave a smooth side surface (important for good
sectioning). Your instructor will demonstrate.
MAKING GLASS KNIVES
Although most electron microscope laboratories have automatic knife
makers, it is good practice to learn the art of making knives by hand. Not
all types of glass are suitable for knives and despite occasional claims,
hardware store plate glass is rarely adequate. In theory, a semi-liquid
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knife edge is made by bringing two natural fractures to a 45° apex. The
quality (smoothness, sharpness and durability) of the edge depends upon
the density, temper and composition of the glass. For these reasons, most
labs purchase good quality glass from vendors of EM supplies. The glass
usually comes as one inch wide strips varying in thickness; usually 1/4,
5/16, or 3/8 inches. The strips must be scrupulously cleaned with acetone
or alcohol. The glass can then be scored using a diamond glass scribe
either free hand or by using a simple Plexiglas scoring guide. Two scores
are made, one across (perpendicular to the length) the glass strip to yield a
1" square piece and one diagonal score towards the first score. Both scores
are made at the same time and should be made with enough pressure so as
to just see and "hear" the score. Be sure to align the edge of the diamond
scribe flush with the guide edges to ensure a straight and precisely placed
score.
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Use the glaziers pliers to first make the cross (perpendicular) fracture and
then the diagonal fracture. Do not touch the knife edge or sides with your
fingers. The contaminants of the fingerprint will prevent adhesion of the
water boat to be mounted on the knife. Examine the knife edge for its
shape and horizontal angle. Note the size of the spur, fracture ridge (burr
line) and curve of the edge (see the diagram).
The boat can be made with short sections of black vinyl electrical tape cut
in half lengthwise. Wrap the tape around the knife (diagonal side) so that
the top edge of the boat is perpendicular with the vertical side of the knife.
Do not leave a gap at the back of the boat and use your fingernail to seal
the adhesive against the glass sides (air bubbles disappear). Seal the back
and sides of the boat with nail polish and allow to dry. The knife may then
be used or stored under cover. It is not a good practice to store knives for
long periods of time (no more than 2-3 days) since they have a tendency
to clutter up the microtome area and to get dull.
THIN SECTIONING
(using the Sorvall Porter-Bloom MT-1 ultramicrotome)
Ultramicrotomy is one of the most difficult techniques to master
since there are many variables contributing to the cutting process. Some
of these are:
plastic hardness
knife quality
knife angle
boat water level
trimmed edge smoothness
vibration
temperature
humidity
cutting speed
tongue in wrong position
15
A good deal of patience is necessary along with steady hands. First,
read the instruction manual for the ultramicrotome and memorize each
control and component. It is convenient to begin a microtomy session by
resetting the specimen holder arm to the rearmost position. Remove the
knife holder and secure the specimen capsule in the collet holder. Be sure
that the knurled ring securing the ball and socket pivot is tight. This
should be done with the specimen arm hook clamp in place so as to not
damage the lead alloy threaded advancing rod inside the microtome. A
new knife may be secured in the holder and placed in its locking
mechanism. You should not have to move or adjust the cool light source
which may be turned on at the start of the session. Unhook the specimen
arm and rotate the sectioning knob to bring the specimen to knife edge
height. Then manually advance the knife stage to within 1-2 mm of the
specimen. Both should now be in view through the dissecting microscope.
Adjust the microscope to the highest magnification and focus. Add fresh,
clean distilled water to the boat so as to have a silver reflection from the
surface. This will occur with the water surface slightly concave and the
water should be adjacent to the knife edge.
Once the knife and specimen are roughly aligned manually the upper
half of the knife stage may be advanced manually using the course and
fine advance controls until cutting the first sections from the tip of the
specimen. With the first piece of plastic section (it will probably be fairly
thick) the knife may then be advanced using the fine advance adjustment
which is calibrated in microns. Advance the fine control one half micron at
a time to create a "face". If done correctly, each time the face comes into
view under the microscope it should appear mirror-like. At this point it is
advisable to back the knife edge away from the specimen using the fine or
coarse adjustment control and to move the knife edge laterally to a new
area. Secure the knife using the locking lever and slowly advance the
knife to the specimen face once more. If the light and water level are
adjusted correctly, as you advance the knife you should eventually see the
reflection of the knife edge in the mirror-like face. The knife edge and its
reflection can be thought of as two lines which should be parallel. If they
are parallel, it will ensure that the first section that is taken by the new
knife edge will not section only part of the face. Although it is difficult to
have the lines exactly parallel one can usually come close by adjusting the
specimen block using the ball socket pivot. It is extremely important that
the first section using the new area of the knife be as thin as possible.
Otherwise the knife edge will dull. Most problems arise with beginning
microtomy due to improper water level, too fast a cutting stroke, wrong
knife angle (this should be about 5°) and a dull knife edge. Unfortunately,
16
this technique is one which is learned most rapidly by doing it and making
mistakes. Your instructor will help you understand the problems as they
arise.
If you are successful at getting good sections they will appear gold or
silver. The thickness of the sections is determined by their refractive color
as calibrated on the thickness chart provided with the microtome. Gold
sections are 800-900 A thick, yield better contrast but slightly less
resolution. If maximum resolution is needed, gray sections on the order of
300-400 A can be attempted. This, however, is extremely difficult and
requires a good deal of experience in attaining.
The sections should float on the water and should adhere to one
another in the form of a ribbon. As the knife edge cuts through the plastic,
it causes the sections to become compressed. This may be alleviated by
exposing the sections to vapors of organic solvents such as chloroform or
ether. This is done by merely dipping a long handled cotton swab in the
solvent and holding it close to but not in contact with the sections near the
knife edge while viewing through the microscope. The sections will appear
to become smooth and large and this will be very noticeable to the eye.
The sections are then ready to be maneuvered to the center of the boat to
be picked up on the grids.
PLACING SECTIONS ON GRIDS
If a noticeable amount of water has evaporated from the boat
causing a more concave surface, it is advisable to add a small amount of
water to the boat so the surface is nearly level before moving the sections.
The sections may be maneuvered using a fine needle (a 0000 stainless
steel insect mounting pin slightly bent at the tip pressed onto a wooden
handle is convenient.) After centering the sections they may be adhered to
the proper grid by grasping the grid in fine tweezers (Pick up the grid only
by the very edge) and bending the grid against the bottom of the petri
dish so that it may be placed dull side down on the surface of the water
and sections. Be careful as you touch down on the surface of the sections
that they are oriented in the middle of the grid. Do not push hard enough
that you break the surface tension of the water. You need only to barely
touch the surface and the sections along with a small drop of water will
adhere to the grid. If the water droplet does not spread out evenly on the
grid it signifies that the grids are dirty. If this is the case you will usually
find that the sections have become wrinkled and do not span over the
holes. If this is the case the grids must be cleaned (see appendix). The
sections on the grids are now ready to be stained.
17
STAINING
Additional contrast of the EM image can be gained by staining the
tissue sections with heavy metals. The strains most commonly used are
uranyl acetate and lead citrate. Both metals apparently bind at sites of
osmium deposition and lead also binds with (i.e. stains) nucleic acids and
glycogen.
Lead Citrate
Lead citrate (Reynold's) is perhaps the best stain available since it
can be used at a high pH and stains a wide variety of cellular components
including nuclear components, ribosomes, membranes, microfilaments and
glycogen. The precise chemical nature of the binding is not well
understood. Care must be taken since lead citrate will react with
atmospheric CO 2 to form a fine precipitate of lead carbonate.
To Stain Sections:
Pour a generous amount of either sodium or potassium hydroxide
pellets around the perimeter of a plastic disposable petri dish. Place the
cover on the dish. Carefully place (don't drop) a 2-3 mm drop of lead
citrate (one for each grid to be stained) on the center surface using a
Pasteur pipet and lifting the plate cover just enough to give the pipet
clearance. The drops will not spread but will remain as droplets. Place the
grids, with section side down, on top of the droplets and cover the plate.
The grids will float. Do not breath onto the petri dish while placing grids
on the drops of stain. Stain for 15-20 minutes. Remove the grids and
immediately but gently dip and stir them in a weak NaOH solution (or KOH
- one or two pellets in 30-40 ml dHOH in a small beaker). Only a few
seconds are needed. Rinse the grids in distilled water by gently dipping
and stirring; blot dry by pressing the surface of the grid not having
sections on a piece of filter paper. This will "wick" most of the water away.
Also blot (wick) away the water between the tweezer tips so the grid can
be placed in a holder without being wetted by a fountain pen-like action.
The grid will air dry quickly and is ready for viewing under the EM.
Throw the petri dish away after taping it shut.
18
Uranyl Acetate
Uranyl acetate may be used during the dehydration process by
making the 50% acetone up to 2% with the stain. Uranyl acetate is not
soluble in pure acetone. An aqueous solution of 2% concentration can be
used to float or dip sections mounted on grids. Epon and Araldite do not
take up aqueous stains well unless they are alkaline. Organic solvent
solutions will usually leave a fine precipitate on the sections and is not
often used. Rinse the solutions well with distilled water. Caution - uranyl
acetate is radioactive. Do not pipette by mouth or spill.
Phosphotungstic acid
May be used as a "negative" stain in that it does not bind particularly
well to anything but instead caused areas other than cellular (organic)
material to appear dark. It is especially useful for viewing molecules such
as proteins (e.g., antibody, DNA) and suspensions of subcellular structures
such as membranes (e.g., mitochondria - elementary particles, etc.) Usually
a 1-2% aqueous solution is used to stain a tissue block during dehydration
for about 30 minutes.
OTHER PROCEDURES
Thick Sections:
It is generally not advisable to begin a structure oriented study at
the electron microscope level. Some form of light microscopy is usually
performed to become familiar with the general structure, orientation and
on occasion, to locate a specific site within the specimen. Specimens
prepared for the electron microscope can be thick sectioned and
appropriately stained for light microscopic examination with ease.
The trimmed specimen capsule is mounted in the ultramicrotome the
same as for thin sectioning. A knife is also secured and may or may not
have a boat. The "face" is formed in the usual manner and thick sections
(2-3 µ ) are made using the fine advance adjustment control. The sections
often have a tendency to roll up like a scroll. This can be prevented by
using a new knife edge and a slow smooth cutting stroke. The sections are
then placed on a clean microscope slide with enough water present to
permit positioning of the sections in an orderly arrangement. The slide is
dried, lightly heat fixed and stained for 5 minutes with Toluidine blue.
Excess dye is rinsed off with distilled water and the sections are then
destained (excess stain in the sections) with ethanol (90-95%) for 2
19
minutes. The slide is dried and a cover slip is secured with a small drop of
the pure plastic mixture used for the embedding procedure. A vial of this
mixture (with DMP-30 added) may be kept for several months in the
freezer (-20°C). The section may be viewed to see if they are of
appropriate quality and can be labeled and stored after curing the plastic
mixture (used for mounting the cover slip) on a warming plate set at 60°C.
Place a weight on the cover slip to insure that it will press the sections flat
against the glass slide.
Whole mounts:
It is often necessary to view a whole or solid object such as a
bacteria, virus, or molecules. For this, it is routine to place the specimen on
a film-coated grid and to coat the object with a thin film of metal. The
grids are usually coated with a Formvar film of varying thickness. A 0.25%
solution in anhydrous ethylene dichloride is kept in a tightly sealed
volumetric. Water will cause holes in the film. A small drop of the
solution is gently laid onto the convex (mounded) surface of distilled water
in a petri dish. The Formvar will spread into a very thin film and the
solvent will evaporate almost immediately. Carefully place clean grids
onto the film surface dull side down using tweezers. The film is thicker at
the center of the dish. Do not allow vibrations while making the film. The
coated grids can be recovered from the plate surface by carefully dropping
a piece of filter paper onto (over) the surface allowing it to wet completely.
Then remove the paper, invert it (grids face up) and place it on absorbent
towels to blot excess water away. After the paper has dried, the grids may
be recovered by raising them straight up off of the paper using sharp
tweezers. The grids may then be used or stored.
Specimens are usually applied to the coated grids by aspiration or by
placing a small drop of water suspension on the surface and allowing it to
dry.
20
Arts and Formulae
PBS -phosphate buffered saline, use either potassium or sodium phosphate.
(9.0 g NaC1, 2.7 g KH2PO 4, d HOH to 1 L, adjust pH)
Formvar resin solution for coating EM grids
0.25% dissolved in ethylene dichloride (0.157 g in 50ml = 0.25%)
Glutaraldehyde
23 ml PBS
2 ml 25% Glut.
OsO4
25 ml PBS, 0.5 g OsO4crystals
CAUTION
Lead Citrate Post Stain:
1.33g Lead nitrate (Pb(NO3)2)
1.76g Sodium citrate (Na3 (C 6 H 5 O 7 )2H 2 O
30 ml dHOH in 50ml volumetric
Shake vigorously for 2 min. then at intervals for 1/2 hr.
8 ml of 1 N NaOH_ (0.4g/10ml dHOH-fresh)
Dilute to 50 ml w dHOH
clear, pH=12
KMnO 4 - 1 g in 50 ml PBS
Plastic Mixture:
16 ml of a 5:3 Epon:Araldite mixture
9 ml NMA
10 ml DDSA
35 ml total - stir exhaustively
15-17 drops DMP-30 - stir again, will turn dark amber.
Toluidine blue for thick sections
1 g borax (sodium carbonate)
100 ml HOH-dissolve
1 g Toluidine blue dye
Stir - filter if necessary
21
Cleaning copper grids
Immerse grids in a 2-4 N HC1 solution for 5 minutes (a drop of detergent
will help the grids sink)
Rinse with distilled water and then acetone in a filter funnel attached to a
vacuum flask. Use filter paper in the funnel and allow the grids to air dry.
Place grids in a covered glass (paper bottom) petri dish (plastic allows
static charge to occur).
Specimen Preparation with L. R. White
I.
Fixation
A.
Dissect tissue into 1mm 3 cubes shortly after procurement.
B.
Place cubes in Phosphate Buffered Saline (PBS) for a 10 minute
wash.
C.
Transfer to a solution of 2% glutaraldehyde in Phosphate
Buffered Saline. (60 min.)
D.
Wash in PBS for 10 minutes.
E.
Transfer blocks of tissue to an osmium tetroxide solution;
consisting of the following: (60-90 minutes)
1.
5 ml. of PBS
2.
5 ml. of 0.1 N HC1
3.
15 ml. of distilled HOH
4.
0.5g OsO4 crystals
II.
Dehydration
A.
Transfer
B.
Transfer
C.
Transfer
D.
Transfer
blocks
blocks
blocks
blocks
to
to
to
to
50% ethanol (15 min.)
75% ethanol (15 min.)
95% ethanol (15 min.)
100% ethanol (Twice) (15-20 min./switch)
III.
Infiltration
A.
Transfer blocks to L.R. White resin (45 min.) four (4) times in
oven at 60° C.
B.
Place blocks in gelatin capsules (1/cap.) and fill to rim with L.
R. White resin.
IV.
Polymerization
A.
Polymerize 20-24 hours at 60°C.
B.
Remove the capsule when fully polymerized.
22
Photography
Making negatives:
The films should be exposed according to the instructions in the EM
operating manual. Generally speaking, contrast is enhanced by longer
exposures with low illumination. However, exposures longer than 1 second
are more prone to blurring from vibration in the building, thus negating
the fine resolution of the EM. It is desirable to have as much contrast in
the recorded image as possible. It is often advantageous to slightly over
expose the film so as to have a "dense" negative. In this case, you would
adjust the illumination (with the condenser lens control) to indicate just
less than a 1 second exposure on the exposure meter and expose (timer
set) for 1 second.
Film development:
The 35mm film is developed as follows:
Dektol (D-72 straight) = 2 minutes
Water (tepid)= 2 changes
Hypo (Fixer)= 5 minutes (save hypo)
Water wash= 5 minutes
Distilled water rinse - dry
Plate film is developed in trays or tanks as follows:
D-19 (2:1 water:D-19)= 4 minutes
Water wash (stop bath)= 10 seconds
Hypo (Fixer - save)= 8 minutes
Water wash= 5 minutes
Printing - Black and White
Additional contrast can be gained during the printing process by
using high contrast paper or polycontrast paper and high contrast filters.
Also, a certain amount of contrast can be gained by again using a longer
exposure with low illumination.
The exposure and illumination settings can be determined by using
"test strips" or an exposure guide on a test print. This consumes a sizeable
quantity however, and it is best to acquire a "feel" for proper exposures by
practice. Most print papers have a generously wide exposure latitude but
the exposure should be such so as to allow the image to develop within 123
1.5 minutes in developer. Among the most often used print papers is
Kodak Kodabromide polycontrast RCII paper in E or F surface. RC stands
for "resin coated" which eliminates the need for a print dryer and it
shortens processing as well. The E and F surfaces differ in that the E
surface is a "matte" surface and the F surface is more "glossy".
Paper development is as follows:
Dektol (D-72, 2:1, water:D-72)= 1-1.5 min.
Stop bath (indicator)= 10-15 sec.
Hypo (can be used)= 3-5 min.
Water wash= 5 min.
Squeegee and air dry
The instructor will demonstrate the use of the photographic enlarger.
It is fairly straight-forward. Take care to remove dust from the negative
and lenses. Always leave the darkroom clean. Wipe up hypo spills since
dry hypo is a fine white powder and can easily contaminate surfaces of
negatives and lenses.
Rinse hands well before handling paper or touching the enlarger or
light switch. It is alright to transfer paper and hands from developer to
stop bath to hypo but it is not good practice to go the other way. Rinse
hands thoroughly of hypo before going back to the developer or stop bath.
Prints may be conveniently labeled using transfer letters and
numbers available in most office supply stores. These transfer markings
are virtually permanent but can be removed by gently scraping with a
scalpel or razor blade.
Magnification (recorded when the film was exposed) can be
multiplied by the magnification factor of the enlarger and represented by
a "micron" or other unit measure on the print as a line or bar.
Prints should be stored out of direct sunlight in a dry environment.
There may be a tendency for the prints to curl or roll up due to absorption
of moisture by the paper backing while the emulsion remains relatively
moisture free (since it is fairly thin). The prints may be mounted on
various types of poster board by glue adhesives or by heat sensitive "dry
mounting" sheets (essentially a wax that melts and adheres to the print
and mount). The print image is virtually permanent if stored or cared for
properly.
24