Histotechniques
Dr Mulazim Hussain Bukhari
MBBS, DCP, MPhil, FCPS, PhD
Associate Prof Pathology
King Edward Medical University, Lahore
25th October 2008,Saturday
CME,DEpartment of Pathology,King Edward
Medical University,Lahore
1
Tissue Processing
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Specimen Accessioning
Gross Examination
Fixation
Tissue Processing
Sectioning
Frozen Sections
Staining
H and E staining
Cover slipping
Decalcification
Artifacts in Histologic Sections
Problems in Tissue Processing
Safety in the Lab
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The lab should be well illuminated and well-ventilated.
Rules and Regulations governing
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formalin and
hydrocarbonds
 such as xylene
 and toluene.
Limits set by the Occupational Safety and Health Administration
(OSHA) that should not be exceeded.
These limits should be revised and revived to reduced any mishap
Cont.
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Every chemical compound used in the laboratory should have a
materials safety data sheet on file
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that specifies the nature,
toxicity,
and safety precautions to be taken when handling the compound.
The laboratory must have a method for disposal of hazardous
wastes.
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Health care facilities processing tissues often contract this to a waste
management company.
Tissues that are collected should be stored in formalin
and may be disposed by incineration
or by putting them through a "tissue grinder" attached to a large sink (similar
to a large garbage disposal unit).
Cont.
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Check the sharpness of scalpel, scissors and quality of
other ones like ruler, probes weighing machines
Every instrument used in the laboratory should meet
electrical safety specifications and have written
instructions regarding its use.
Flammable materials may only be stored in approved
rooms and only in storage cabinets that are designed
for this purpose.
Cont.
 Fire safety procedures are to be
posted.
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Safety equipment including fire
extinguishers,
fire blankets,
and fire alarms should be within easy
access.
A shower and eyewash should be readily
available.
No smoking, eating or movements in
the labs
Use disposable gloves
Cont.
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Laboratory accidents must be documented and
investigated with incident reports and industrial
accident reports.
Specific hazards that you should know about
include:
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Bouin's solution is made with picric acid. This acid is
only sold in the aqueous state. When it dries out, it
becomes explosive.
Sodium azide
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Many reagent kits have sodium azide as a
preservative.
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You are supposed to flush solutions containing
sodium azide down the drain
 with lots of water, or
 there is a tendency for the azide to form metal
azides in the plumbing.
These are also explosive.
Drainage
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Benzidine, benzene, anthracene, and napthol
containing compounds are carcinogens and should not
be used.
Mercury-containing solutions (Zenker's or B-5) should
always be discarded into proper containers.
Mercury, if poured down a drain, will form amalgams
with the metal that build up and cannot be removed.
Hazards of usually used formalin
Objective
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Tissues from the body taken for diagnosis of disease processes must
be processed in the histology laboratory to produce microscopic
slides that are viewed under the microscope by pathologists.
The techniques for processing the tissues, whether biopsies, larger
specimens removed at surgery, or tissues from autopsy
The persons who do the tissue processing and make the glass
microscopic slides are histotechnologists
Specimen Accessioning
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Tissue specimens received in the surgical
pathology laboratory have a request form that
lists the patient information and history along with
a description of the site of origin.
The specimens are accessioned by giving them a
number that will identify each specimen for each
patient
Grossing
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Describing the specimen
Placing all or parts of it into a small plastic
cassette
When a malignancy is suspected
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Inking a gross specimen for margins
Fixation
Types of fixatives (AMAPO)
 Aldehydes
 Mercurials
 Alcohols
 Picrates
 Oxidizing agents
Fixation - factors affecting fixation
There are a number of factors that will affect the
fixation process:
 Buffering
 Penetration
 Volume
 Temperature
 Concentration
 Time interval
 Position of tissue
Buffering
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Fixation is best carried out close to neutral pH, in the range of
6-8.
Hypoxia of tissues lowers the pH, so there must be buffering
capacity in the fixative to prevent excessive acidity.
Acidity favors formation of formalin-heme pigment that
appears as black, polarizable deposits in tissue.
Common buffers include phosphate, bicarbonate, cacodylate,
and veronal.
Commercial formalin is buffered with phosphate at a pH of 7.
Penetration
 Penetration of tissues depends upon the
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diffusability of each individual fixative,
which is a constant.
Formalin and alcohol penetrate the best,
and glutaraldehyde the worst.
Mercurials and others are somewhere in
between.
One way to get around this problem is
sectioning the tissues thinly (2 to 3 mm).
Penetration into a thin section will occur
more rapidly than for a thick section
Volume  The volume of fixative is important.
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There should be a 10:1 ratio of fixative to
tissue.
Obviously, we often get away with less
than this, but may not get ideal fixation.
One way to partially solve the problem is
to change the fixative at intervals to
avoid exhaustion of the fixative.
Agitation of the specimen in the fixative
will also enhance fixation.
Temperature
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Increasing the temperature, as with all
chemical reactions, will increase the speed
of fixation, as long as you don't cook the
tissue.
Hot formalin will fix tissues faster, and this
is often the first step on an automated
tissue processor.
Concentration
of
fixative
 Concentration of fixative should be
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adjusted down to the lowest level
possible, because you will expend less
money for the fixative.
Formalin is best at 10%;
glutaraldehyde is generally made up at
0.25% to 4%.
Too high a concentration may
adversely affect the tissues and
produce artefact similar to excessive
heat.
Time interval
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Also very important is time interval from of
removal of tissues to fixation.
The faster you can get the tissue and fix it, the
better.
Artefact will be introduced by drying, so if tissue
is left out, please keep it moist with saline.
The longer you wait, the more cellular
organelles will be lost and the more nuclear
shrinkage and artefactual clumping will occur
Kaiserling formula for preservation
for surgical specimens for museum
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Formalin pure
Distilled water
Potassium acetate(CH3COOK )
Chloral hydrate
5 liter
22.5 liter
250gm
50 gm

27 liter
Potassium acetate is used in mixtures applied for tissue
preservation, fixation, and mummification. Most museums today
use the formaldehyde-based method recommended by Kaiserling in
1897 and containing potassium acetate.
For example, Lenin's mummy was soaked in a bath containing
potassium acetate
Gough Sections:
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Whole organs may be sectioned on paper
by the methods of Gough and Wentworth.
These sections provide valuable
information on whole organ structure and
serve as links between mounted museum
specimens and histologic sections.
Colour restoration
 Small amount of sodium
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hydrosulphite to preserve the
colour.
If the container is properly sealed,
the colour restoration is then
permanent.
For photography, the procedure is
to first wash and clean the
specimen.
It is then soaked in an excess of
60% ethanol until the colour has
been restored satisfactorily
Characteristics of Fixatives
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Chemical Fixatives
Freeze Substitution
Microwave Fixation
Ideal Fixative
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Penetrate cells or tissue rapidly
Preserve cellular structure before cell
can react to produce structural artifacts
Not cause autofluorescence, and act as
an antifade reagent
Chemical Fixation
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Coagulating Fixatives
Crosslinking Fixatives
Coagulating Fixatives
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Ethanol
Methanol
Acetone
Coagulating Fixatives
Advantages
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Fix specimens by rapidly changing hydration state
of cellular components
Proteins are either coagulated or extracted
Preserve antigen recognition often
Disadvantages
• Cause significant shrinkage of specimens
• Difficult to do accurate 3D confocal images
• Can shrink cells to 50% size (height)
• Commercial preparations of formaldehyde
contain methanol as a stabilizing agent
Crosslinking Fixatives
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Glutaraldehyde
Formaldehyde
Ethelene glycol-bis-succinimidyl succinate
(EGS)
Cross-linking Fixatives
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Form covalent crosslinks that are determined by
the active groups of each compound
Principles ofOnce
Fixation
tissues are removed from the body,
they undergo a process of self-destruction
or autolysis
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which is initiated soon after cell death by the
action of intracellular enzymes causing the
breakdown of protein and eventual
liquefaction of the cell.
Principles of Fixation
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Autolysis is independent of any bacterial action,
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retarded by cold,
greatly accelerated at temperatures of about 30°C
and
almost inhibited by heating to 50°C
Cont.
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Autolysis is more severe in tissues which are rich in
enzymes,
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such as the liver,
brain and kidney,
and is less rapid in tissues such as elastic fibre and collagen.
Cont.
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By light microscopy, autolysed tissue presents a
`washed-out' appearance with swelling of
cytoplasm,
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eventually converting to a granular, homogeneous
mass which fails to take up stains.
How Autolysed tissue looks like
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The nuclei of autolytic cells may show
some of the changes of necrosis
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including condensation (pyknosis),
fragmentation (karyorrhexis) and
lysis (karyolysis)
D/D
these are not accompanied by an
inflammatory or cellular response.
How Autolysed tissue looks like
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There may be diffusion of intracellular
substances of diagnostic significance, such as
glycogen which is lost from the cells in the
absence of prompt and suitable fixation.
Autolysis also causes desquamation of
epithelium which separates from its basement
membranes.
Bacterial Action on dead tissue
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Bacterial decomposition can also produce
changes in tissues that mimic those of autolysis
and is brought about by bacterial proliferation
in the dead tissue.
Bacterial Action on dead tissue
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Such bacteria may normally be present in the
body during life such as the non-pathogenetic
organisms present in the bowel, or may be
present in diseased tissues at the time of death
such as in septicaemia.
The objective of fixation
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is to preserve cells and tissue constituents in as
close a life-like state as possible and to allow them
to undergo further preparative procedures without
change.
Fixation arrests autolysis and bacterial
decomposition and stabilizes the cellular and tissue
constituents so that they withstand the subsequent
stages of tissue processing.
Aside from these requirements for the production of
tissue sections, increasing interest in cell
constituents and the extensive use of
immunohistochemistry to augment histological
diagnosis has imposed additional requirements.
Cont.
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Fixation should also provide for the
preservation of tissue substances and proteins.
Fixation is, therefore, the first step and the
foundation in a sequence of events that
culminates in the final examination of a tissue
section.
Common pitfalls of fixation
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It is relevant to point out that fixation in itself constitutes a major
artifact.
The living cell is fluid or in a semi-fluid state, Whereas fixation
produces coagulation of tissue proteins and constituents, a
necessary event to prevent their loss or diffusion during tissue
processing; the passage through hypertonic and hypotonic solutions
during tissue processing would otherwise disrupt the cells.
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For example, if fresh unfixed tissues were washed for prolonged
periods in running water, severe and irreparable damage and cell lysis
would result.
In contrast, if the tissues were first fixed in formalin, subsequent
immersion in water is generally harmless.
Summary of objective
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Fixation:
Confers chemical stability on the
tissue
Hardens the tissue (helps further
handling)
Halts enzyme autolysis
Halts bacterial putrefaction
May enhance later staining techniques
Introduces a 'consistent artifact'
Aldehydes
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include formaldehyde (formalin) and
glutaraldehyde.
Tissue is fixed by cross-linkages formed in the
proteins, particularly between lysine residues.
This cross-linkage does not harm the structure
of proteins greatly, so that antigenicity is not
lost.
Cont.
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Therefore, formaldehyde is good for
immunoperoxidase techniques. Formalin
penetrates tissue well, but is relatively slow.
The standard solution is 10% neutral buffered
formalin.
A buffer prevents acidity that would promote
autolysis and cause precipitation of formolheme pigment in the tissues.
Formaldehyde
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Formaldehyde, as 4% buffered
formaldehyde (10% buffered formalin), is
the most widely employed universal
fixative particularly for routine paraffin
embedded sections.
It is a gas with a very pungent odor,
soluble in water to a maximum extent of
40% by weight and is sold as such under
the name of formaldehyde (40%) or
formalin (a colorless liquid).
Formaldehyde
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Formaldehyde is also obtainable in a stable
solid form composed of high molecular weight
polymers known as paraformaldehyde.
Cont.
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Heated paraformaldehyde generates pure gaseous formaldehyde
which, when dissolved in water, reverts mostly to the monomeric
form.
Aqueous formaldehyde exists principally in the form of its
monohydrate, methylene glycol, CH2(OH)2, and as low molecular
weight polymeric hydrates or polyoxymethylene glycols.
It has been suggested that the hydrated form, methylene glycol, is
the reactive component of formaldehyde but this has been disputed.
Preparation of 10%
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Four per cent formaldehyde or 10% buffered
formalin is commonly prepared by adding 100
ml of 40% formaldehyde to 900 ml distilled
water with 4 g sodium phosphatase, monobasic
and 6.5 g sodium phosphate, dibasic
(anhydrous).
Formaldehyde solutions
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10% neutral buffer formalin (4% formaldehyde)
REAGENTS REQUIRED
1 40% formaldehyde 100 ml
2 Distilled water 900 ml
3 Sodium dihydrogen orthophosphate 4 g
4 Disodium hydrogen orthophosphate
(anhydrous) 6.5 g
(sodium hydrosulphite)
METHOD
Prepare, using quantities indicated. Fixation time:
24-72 hours
Buffered formaldehyde-glutaraldehyde
200 mOsm38
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REAGENTS REQUIRED
1 -Sodium dihydrogen orthophosphate 1.6 g
2-Sodium hydroxide 0.27 g
3-Distilled water 88 ml
4-40% formaldehyde 10 ml
5-50% glutaraldehyde 2 ml
METHOD
Prepare, using quantities indicated. Fixation
time: 16-24 hours.
Formol saline
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REAGENTS REQUIRED
1- 40% formaldehyde 100 ml
2 -Sodium chloride 9 g
3 -Tap water 900 ml
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METHOD
Prepare, using quantities indicated
Some other forms of Fixatives
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Baker's formol-calcium (modified)
REAGENTS REQUIRED
1 40% formaldehyde 100 ml
2 Distilled water 900 ml
3 10% calcium chloride 100 ml
4 7 g of cadmium chloride is sometimes added to the mixture
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METHOD
Prepare, using quantities indicated. Fixation time: 16-24 hours.
Formol saline
REAGENTS REQUIRED
1 40% formaldehyde 100 ml
2 Sodium chloride 9 g
3 Tap water 900 ml
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METHOD
Prepare, using quantities indicated.
Alcoholic formaldehyde
REAGENTS REQUIRED
1 40% formaldehyde 100 ml
2 95% alcohol 900 ml
3 0.5 g calcium acetate may be added to this mixture to ensure neutrality
METHOD
Prepare, using quantities indicated. Fixation time: 16-24 hours.
Fixatives - general usage
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Formalin is used for all routine surgical
pathology and autopsy tissues when an H
and E slide is to be produced.
Formalin is the most forgiving of all fixatives
when conditions are not ideal, and there is
no tissue that it will harm significantly.
Zenker's fixatives
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Zenker's fixatives are recommended for
reticuloendothelial tissues including
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lymph nodes,
spleen,
thymus, and
bone marrow.
Zenker's fixes nuclei very well and gives good
detail.
However, the mercury deposits must be removed
(dezenkerized) before staining or black deposits
will result in the sections
Bouin's solution
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Bouin's solution is sometimes
recommended for fixation of
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testis,
GI tract, and
endocrine tissue.
It does not do a bad job on hematopoietic
tissues either, and doesn't require
dezenkerizing before staining
Glutaraldehyde
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Glutaraldehyde is recommended for
fixation of tissues for electron
microscopy.
The glutaraldehyde must be cold and
buffered and not more than 3 months
old.
The tissue must be as fresh as
possible
Preferably sectioned within the
glutaraldehyde at a thickness no more
than 1 mm to enhance fixation
Alcohols
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Alcohols, specifically ethanol, are used primarily
for cytologic smears.
Ethanol (95%) is fast and cheap.
Since smears are only a cell or so thick, there is
no great problem from shrinkage, and since
smears are not sectioned, there is no problem
from induced brittleness.
Note: For fixing frozen sections, you can use just
about anything--though methanol and ethanol are
the best
Glutaraldehyde
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Glutaraldehyde causes deformation of alpha-helix
structure in proteins so is not good for
immunoperoxidase staining.
However, it fixes very quickly so is good for electron
microscopy.
It penetrates very poorly, but gives best overall
cytoplasmic and nuclear detail.
The standard solution is a 2% buffered glutaraldehyde
Glutaraldehyde
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First used in 1962 by Sabatini et al*
Shown to preserve properties of
subcellular structures by EM
Renders tissue autofluorescent so less
useful for fluorescence microscopy,
but fluorescence can be attenuated by
NaBH4.
Forms a Schiff’s base with amino
groups on proteins and polymerizes
via Schiff’s base catalyzed reactions
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Forms extensive crosslinks - reacts with the -amino
group of lysine, -amino group of amino acids - reacts
with tyrosine, tryptophan, histidine, phenylalanine and
cysteine
Fixes proteins rapidly, but has slow penetration rate
Can cause cells to form membrane blebs
Glutaraldehyde
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Supplied commercially as either 25% or 8% solution
Must be careful of the impurities - can change
fixation properties - best product from Polysciences
(Worthington, PA)
As solution ages, it polymerizes and turns yellow.
Store at -20 °C and thaw for daily use. Discard.
Summary of Formaldehyde
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Crosslinks proteins by forming
methelene bridges between reactive
groups
The ratelimiting step is the deprotonation of amino groups, thus the
pH dependence of the crosslinking
Functional groups that are reactive are
amido, guanidino, thiol, phenol,
imidazole and indolyl groups
Can crosslink nucleic acids
Cont.
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Therefore the preferred fixative for in situ
hybridization
Does not crosslink lipids but can produce
extensive vesiculation of the plasma
membrane which can be averted by
addition of CaCl2
Not good preservative for microtubules at
physiologic pH
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Protein crosslinking is slower than for
glutaraldehyde, but formaldehyde penetrates 10
times faster.
It is possible to mix the two and there may be
some advantage for preservation of the 3D
nature of some structures.
Mercurials
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fix tissue by an unknown mechanism.
They contain mercuric chloride and include such
well-known fixatives as B-5 and Zenker's.
These fixatives penetrate relatively poorly and
cause some tissue hardness, but are fast and
give excellent nuclear detail.
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Their best application is for fixation of
hematopoietic and reticuloendothelial
tissues.
Since they contain mercury, they must be
disposed of carefully
Alcohols including methyl alcohol (methanol)

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and ethyl alcohol (ethanol), are protein
denaturants and are not used routinely
for tissues because they cause too
much brittleness and hardness.
However, they are very good for
cytologic smears because they act
quickly and give good nuclear detail.
Spray cans of alcohol fixatives are
marketed to physicians doing PAP
smears, but cheap hairsprays do just
as well
Oxidizing agents
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include permanganate fixatives
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dichromate fixatives
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Potassium permanganate,
Potassium dichromate,
Osmium tetroxide.
They cross-link proteins, but cause
extensive denaturation.
Some of them have specialized
applications, but are used very infrequently.
Bouin’s Solution (Picrates)
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include fixatives with picric acid.
Foremost among these is Bouin's
solution.
It has an unknown mechanism of
action.
It does almost as well as mercurials
with nuclear detail but does not
cause as much hardness.
Picric acid is an explosion hazard in
dry form.
As a solution, it stains everything it
touches yellow, including skin.
On the left with H&E staining black mercuric
chloride precipitate is seen in this lymphoma fixed in
B-5 and not properly dezenkerized. This precipitate
is seen on the right under polarized light
microscopy.
Removal of Pig
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Formalin pigment
1. Dewax the sections, rinse in 100% alcohol,
rinse in 70% alcohol, rinse in distilled water.
2. Treat in saturated alcoholic picric acid for 30
minutes to 2 hours.
3. Wash well in running tap water.
4. If yellow staining of the section persists rinse in
dilute lithium carbonate.
5. Rinse in tap water.
6. Continue with method.
Mercury pigment

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1. Dewax the sections, rinse in 100%
alcohol, rinse in 70% alcohol, rinse in
distilled water.
2. Treat in Lugol's iodine for 2 minutes.
3. Decolourise in 5% sodium thiosulphate
for 5 minutes.
4. Wash well in running tap water.
5. Continue with method.
Dichromate pigment

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1. Dewax the sections, rinse in 100%
alcohol, rinse in 70% alcohol, rinse in
distilled water.
2. Treat in 2% HCl in 70% alcohol 16-24
hours.
3. Rinse in tap water.
4. Continue with method.
Tissue Processing
 Once the tissue has been fixed, it must
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be processed into a form in which it can
be made into thin microscopic sections.
The usual way this is done is with
paraffin.
Tissues embedded in paraffin, which is
similar in density to tissue, can be
sectioned at anywhere from 3 to 10
microns, usually 6-8 routinely.
The technique of getting fixed tissue
into paraffin is called tissue processing


Dehydration
Clearing
Dehydration
Wet fixed tissues (in aqueous solutions) cannot
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be directly infiltrated with paraffin.
First, the water from the tissues must be
removed by dehydration.
This is usually done with a series of alcohols,
say 70% to 95% to 100%.
Sometimes the first step is a mixture of
formalin and alcohol.
Other dehydrants can be used, but have major
disadvantages.

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Acetone is very fast, but a fire
hazard, so is safe only for small,
hand-processed sets of tissues.
Dioxane can be used without
clearing, but has toxic fumes
Clearing

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Removal of the dehydrant with a
substance that will be miscible with
the embedding medium (paraffin).
The commonest clearing agent is
xylene.
Toluene works well, and is more
tolerant of small amounts of water left
in the tissues, but is 3 times more
expensive than xylene.
Chloroform used to be used, but is a
health hazard, and is slow.
Methyl salicylate is rarely used
because it is expensive, but it smells
nice (it is oil of wintergreen).
Embedding
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Finally, the tissue is infiltrated with the
embedding agent, almost always paraffin.
Paraffins can be purchased that differ in
melting point, for various hardnesses,
depending upon the way the histotechnologist
likes them and upon the climate (warm vs.
cold).
Wax hardness (viscosity) depends upon the
molecular weight of the components and the
ambient temperature.
High molecular weight mixtures melt at higher
temperatures than waxes comprised of lower
molecular weight fractions.
Paraffin wax is traditionally marketed by its
melting points which range from 39°C to 68°C.
Other agents

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A product called paraplast contains added
plasticizers that make the paraffin blocks easier
for some technicians to cut.
A vacuum can be applied inside the tissue
processor to assist penetration of the embedding
agent.
methyl methacrylate,
glycol methacrylate,
araldite, and epon.
Methyl methacrylate
General Embedding Procedure

METHOD
1 Open the tissue cassette, check against
worksheet entry to ensure the correct number of
tissue pieces are present.

2 Select the mould, there should be sufficient room
for the tissue with allowance for at least a 2 mm
surrounding margin of wax.

3 Fill the mould with paraffin wax.

4 Using warm forceps select the tissue, taking care
that it does not cool in the air; at the same time.

5 Chill the mould on the cold plate, orienting the
tissue and firming it into the wax with warmed
forceps. This ensures that the correct
orientation is maintained and the tissue surface
to be sectioned is kept flat.

6 Insert the identifying label or place the
labelled embedding ring or cassette base onto
the mould.

7 Cool the block on the cold plate, or carefully
submerge it under water when a thin skin has
formed over the wax surface.

8 Remove the block from the mould.

9 Cross check block, label and worksheet
Sectioning tissues






Turn on the water bath and check that the
temp is 35-37ºC.
Use fresh deionized water (DEPC treated
water must be used if in situ hybridization will
be performed on the sections).
Blocks to be sectioned are placed face down
on an ice block or heat sink for 10 minutes.
Place a fresh blade on the microtome.
Insert the block into the microtome chuck so
the wax block faces the blade and is aligned
in the vertical plane. Set the dial to cut 4-10
µM sections.
The blade should angled 4-6º.







Face the block by cutting it down to the desired
tissue plane and discard the paraffin ribbon.
If the block is ribboning well then cut another four
sections and pick them up with forceps or a fine
paint brush and float them on the surface of the
37ºC water bath.
Float the sections onto the surface of clean glass
slides.
If the block is not ribboning well then place it back
on the ice block to cool off firm up the wax.
If the specimens fragment when placed on the
water bath then it may be too hot.
Place the slides with paraffin sections in a 65°C
oven for 20 minutes (so the wax just starts to
melt) to bond the tissue to the glass.
Slides can be stored overnight at room
temperature
H and E staining




Hematoxylin is the oxidized product
of the logwood tree known as
hematein.
Since this tree is very rare nowadays,
most hematein is of the synthetic
variety.
In order to use it as a stain it must be
"ripened" or oxidized.
This can be done naturally by putting
the hematein solution on the shelf
and waiting several months, or by
buying commercially ripened
hematoxylin or by putting ripening
agents in the hematein solution.
Cont.



Hematoxylin will not directly stain
tissues, but needs a "mordant" or link
to the tissues. This is provided by a
metal cat ion such as iron, aluminum,
or tungsten.
The variety of hematoxylins available
for use is based partially on choice of
metal ion used.
They vary in intensity or hue.
Hematoxylin, being a basic dye, has
an affinity for the nucleic acids of the
cell nucleus.
Cont.





Hematoxylin stains are either "regressive" or
"progressive".
With a regressive stain, the slides are left in the
solution for a set period of time and then taken
back through a solution such as acid-alcohol that
removes part of the stain.
This method works best for large batches of slides
to be stained and is more predictable on a day to
day basis.
With a progressive stain the slide is dipped in the
hematoxylin until the desired intensity of staining
is achieved, such as with a frozen section.
This is simple for a single slide, but lends itself
poorly to batch processing.
Eosin




Eosin is an acidic dye with an affinity
for cytoplasmic components of the cell.
There are a variety of eosins that can
be synthesized for use, varying in their
hue, but they all work about the same.
Eosin is much more forgiving than
hematoxylin and is less of a problem in
the lab.
About the only problem you will see is
overstaining, especially with decalcified
tissues
Decalcification




Some tissues contain calcium deposits which
are extremely firm and which will not section
properly with paraffin embedding owing to the
difference in densities between calcium and
parffin.
Bone specimens are the most likely type here,
but other tissues may contain calcified areas
as well.
This calcium must be removed prior to
embedding to allow sectioning.
A variety of agents or techniques have been
used to decalcify tissue and none of them
work perfectly.




Mineral acids,
organic acids,
EDTA, and
electrolysis have all been used.
Strong mineral acids


nitric and
hydrochloric acids



rapid
damage cellular morphology,
so are not recommended for delicate
tissues such as bone marrow.
Organic acids





acetic and
formic acid are better suited to bone
marrow, since they are not as harsh.
However, they act more slowly on dense
cortical bone.
Formic acid in a 10% concentration is
the best all-around decalcifier.
Some commercial solutions are available
that combine formic acid with formalin to
fix and decalcify tissues at the same
time.
EDTA and Electrolysis

EDTA can remove calcium and is
not harsh (it is not an acid)




but it penetrates tissue poorly and
works slowly and is
expensive in large amounts.
Electrolysis has been tried in
experimental situations where
calcium had to be removed with
the least tissue damage.

It is slow and not suited for routine
daily use.
Procedure:


Specimens should be decalcified in hydrochloric
acid/formic acid working solution 20 times their
volume.
Change to fresh solution each day until
decalcification is complete.





It may take 24 hours up to days or months
depending on size of the specimens.
See below for the testing procedures
Once the decalcification is complete, rinse
specimens in water briefly and transfer to
ammonia solution to neutralize acids left in
specimens for 30 minutes.
Wash specimens in running tap water
thoroughly up to 24 hours.
Routine paraffin embedding.
End-Point of Decalcification



X-ray (the most accurate way)
Chemical testing (accurate)
Physical testing (less accurate and
potentially damage of specimen)
Chemical Test:










5% Ammonium Hydroxide Stock:
Ammonium hydroxide, 28% -------------------5 ml
Distilled water ---------------------------------- 95
ml
Mix well
5% Ammonium Oxalate Stock:
Ammonium oxalate ---------------------------- 5
ml
Distilled water --------------------------------- 95
ml
Mix well
Ammonium Hydroxide/Ammonium Oxalate
Working Solution:
Use equal parts of the 5% ammonium
hydroxide solution and the 5% ammonium
oxalate solution.
Procedure:




Insert a pipette into the decalcifying
solution containing the specimen.
Withdraw approximately 5 ml of the
hydrochloric acid/formic acid
decalcification solution from under the
specimen and place it in a test tube.
Add approximately 10 ml of the
ammonium hydroxice/ammonium
oxalate working solution, mix well and
let stand overnight.
Decalcification is complete when no
precipitate is observed on two
consecutive days of testing. Repeat
this test every two or three days.
Physical Tests:





The physical tests include bending the
specimen or inserting a pin, razor, or
scalpel directly into the tissue.
The disadvantage of inserting a pin,
razor, or scalpel is the introduction of
tears and pinhole artifacts.
Slightly bending the specimen is safer
and less disruptive but will not
conclusively determine if all calcium
salts have been removed.
After checking for rigidity, wash
thoroughly prior to processing.