Histopathology notes

Histopathologic Techniques

By: Harley Rose B. Bautista, RMT

FRESH TISSUE EXAMINATION

⮚ Methods of tissue examination may vary according to: the structural and chemical components of the cells
to be examined or studied, the nature and amount of the tissue to be evaluated, and the need for an
immediate examination of a tissue structure.

METHODS OF FRESH TISSUE EXAMINATION

1. Teasing or dissociation – process whereby selected tissue specimen is immersed in a watch glass
containing isotonic salt solution, carefully dissected or separated, and examined under the microscope.
Either unstained by phase contrast microscope or bright field microscope or stained with differential
dyes.
2. Squash preparation (crushing) – is a process whereby small pieces of tissue not more than one mm. in
diameter are placed in a slide and forcibly compressed with another slide or with a cover glass.
3. Smear preparation – process of examining sections or sediments, whereby cellular materials are spread
lightly over a slide means of a wire loop. This technique is especially useful in cytological
examinations, particularly for a cancer diagnosis.
3.1 streaking – with an applicator stick, the material is rapidly and gently applied in a direct or zigzag
line throughout the slide.
3.2 Spreading – has the advantage of maintaining cellular inter relationships of the material to be
examined. It is especially recommended for smear preparations of fresh sputum and bronchial
aspirtates and also for thick mucous secretions.
3.3 Pull apart – useful for preparing smears of thick secretions such as serous fluids, concentrated
sputum and blood smears.
3.4 Touch preparation (impression smear) – has the added advantage in that cells may be examined
without destroying their actual intracellular relationship, and without separating them from their
normal surroundings.
4. Frozen sections – for rapid diagnosis, especially recommended when lipids and nervous tissue
elements are to be demonstrated.
COMMONBLY USED METHODS OF FREEZING INCLUDE:
4.1 LIQUID NITROGEN
4.2 ISOPENTANE COOLED NITROGEN
4.3 CARBON DIOXIDE GAS
4.4 AEROSOL SPRAYS

STEPS IN PROCESSING OF TISSUES

1. Fixation
2. Dehydration
3. Clearing
4. Infiltration / impregnation
5. Embedding
6. Trimming
7. Section – cutting
8. Staining
9. Mounting
10. Labelling

FIXATION

-first and most critical step in histotechnology involves preserving fresh tissue examination.

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-primary aim: to preserve the morphological and chemical integrity of the cell in as life-like manner as
possible.

- secondary aim: to harden and protect the tissue from the trauma of further handling so that it's easier to
cut during gross examination.

- the most important reactions in maintaining morphology in the fixation of tissues for routine
histopathology are those that stabilize proteins.

TWO BASIC MECHANISMS INVOLVED IN FIXATION

1. ADDITIVE FIXATION

❖ Whereby the chemical constituent of the fixative is taken in and becomes part of the tissue by forming
cross links or molecular complexes and giving stability to the protein.
❖ Example: formalin, mercury and osmium tetroxide

2. NON ADDITIVE FIXATIVES

❖ Whereby the fixing agent is not incorporated into the tissue but alters the tissue composition and stabilizes
the tissue by removing bound water attached to H-bonds of certain groups within the protein molecule.

MAIN FUCTIONS INVOLVED IN FIXATIONS

1. Hydrogen Ion concentration

2. temperature
3. thickness of section
4. osmolality
5. concentration
6. duration of fixation

CONSIDERATIONSOF FIXATION

1. Speed
2. Penetration
3. Volume
4. Duration of fixation

EFFECTS OF FIXATIVES IN GENERAL

1. They harden soft and friable tissues and make the handling and cutting of sections easier.
2. They make the cells resistant to damage and distortion caused by hypotonic and hypertonic solutions used
during tissue processing.
3. They inhibit bacterial decomposition.
4. They act as mordants or accentuators to promote and hasten staining, or they may inhibit certain dyes in
favor of another.
5. They reduce the risk of infections during the handling and actual processing of tissues

CHARACTERISTICS OF GOOD FIXATIVES

1. Must be cheap
2. Must be stable
3. Must be safe to handle
4. Must kill the cell quickly thereby producing minimum distortion of cell constituents
5. Must be inhibit bacterial decomposition
6. Must produce minimum shrinkage
7. Must permit rapid and even penetration of tissues
8. It must harden tissues thereby making the cutting of sections easier

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TYPES OF FIXATIVES ACCORDING TO:

I. COMPOSITION

a) SIMPLE FIXATIVES – made up of only one component substance.

b) COMPOUND FIXATIVES – those that are made up of two or more fixatives.

II. ACTION

A. MICROANATOMICAL FIXATIVES – are those that permit the general microscopic study of
tissue structures without altering the structural pattern of the tissues.
B. CYTOLOGICAL FIXATIVES – preserves specific parts and particular microscopic elements of
the cell itself.

I. ALDEHYDE FIXATIVES

1. FORMALDEHYDE

❖ One of the most widely used fixatives is 10% formalin, which must be diluted 1:10 or 1:20.
❖ Cheap readily available, easy to prepare.
❖ Stock solution: 40%
❖ Fumes are irritating to the nose and eyes and may cause sinusitis or excessive lacrimation.

2. 10% FORMOL SALINE

❖ Recommended for fixation of central nervous tissues and general post mortem tissues for
histochemical examination.

3. 10% NEUTRAL BUFFERED FORMALIN

❖ Recommended for preservation and storage of surgical, post mortem and research specimens.
❖ Best fixative for tissues containing iron pigments and for elastic fibers which do not stain well
after Susa, Zenker or chromate fixation.

4. FORMOL CORROSIVE (FORMOL SUBLIMATE)

❖ Excellent for many staining procedures including silver reticulum methods.

❖ There’s no need for washing out.

5. ALCOHOLIC FORMALIN ( GENDRE’S FIXATIVE)

❖ Fixation is faster.
❖ It can be used for rapid diagnosis because it fixes and dehydrates at the same time.
❖ Used to fix sputum since it coagulates mucus.

6. GLUTARALDEHYDE

❖ Made up of two formaldehyde residues , linked by three carbon chains.

❖ For small tissue fragments: 2.5% solution used fix for 2-4 hrs
❖ For larger tissues less than 4 mm thick: fixed in 6-8 hrs up to 24 hrs.
❖ More expensive

II. METALLIC FIXATIVES

1. MERCURIC CHLORIDE

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❖ Most common metallic fixative, frequently used in saturated aqueous solutions 5-7%.
❖ Contains black precipitates of mercury; remedy: treating the section with 0.5% iodine
solution in 70% ethanol for 5-10 mins. Sections are rinsed in water, decolorized for 5 mins.
In 5% sodium thiosulfate and washed in running water.
❖ Routine fixative of choice for preservation of cell detail in tissue photography.
❖ Recommended for renal disease , fibrin , connective tissues and muscles.

1.1 ZENKER’S FLUID

❖ Made up of mercuric chloride stock solution to which glacial acetic acid has been
added just before its use to prevent turbidity and formation of a dark precipitate.
❖ Recommended for small pieces of liver, spleen, connective tissue fibers and nuclei.

1.2 ZENKER FORMOL (HELLYS SOLUTION)

❖ Excellent micro anatomic fixative for pituitary gland, bone marrow and blood
containing organs such as spleen and liver.
❖ Preserves cytoplasmic granules well
❖ Brown pigments are produced if tissues are allowed to stay in fixative for more
than 24 hrs due to RBC lysis.
❖ REMEDY: immersing the tissue in saturated alcoholic picric acid or sodium
hydroxide.

1.3 HEIDENHAIN’S SUSA SOL

❖ Recommended mainly for tumor biopsies especially of the skin; it is an

excellent cytologic fixative.

1.4 B-5 FIXATIVE

❖ Commonly used for bone marrow biopsies.

2. CHROMATE FIXATIVES

2.1 CHROMIC ACID

❖ Used in 1-2% aqueous sol. Usually as a constituent of compound fixative.

2.2 POTASSIUM DICHROMATE

❖ Used in a 3% aqueous solution.

❖ Preserves lipids and mitochondria.

2.3 REGARD’S (MULLER’S) FLUID

❖ Recommended for demonstration of chromatin, mitochondria, mitotic figures, golgi bodies

and colloid-containing tissues.
❖ Does not preserve fats.
❖ Prolonged fixation blackens tissue pigments ; REMEDY: removed by washing the tissues
in running tap water prior to dehydration.

2.4 ORTH’S FLUID

❖ Recommended for acid mucopolysaccharides.

3. PICRIC ACID

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❖ Excellent fixative for glycogen demonstration.

❖ May produce yellow color; REMEDY: tissue may be placed in 70% ethyl acohol followed by
5% sodium thiosulfate and then washed in running water.

3.1 BOUINS SOLUTION

❖ Recommended for fixation of embryos and pituitary biopsies.

❖ Excellent fixative for preserving soft and delicate structures (eg: endometrial cutterings)
❖ Not suitable for fixing kidney structures, lipids and mucus.

3.2 BRASIL’S ALCOHOLIC PICROFORMOL FIXTAIVE

❖ Better and lessy messy than bouin’s solution

❖ Excellent fixative for glycogen.

4. GLACIAL ACETIC ACID

❖ Solidifies at 17 degcel.

5. ALCOHOL FIXATIVES

5.1 METHYL ALCOHOL 100%

❖ Excellent fixing dry and wet smears, blood smear and bone marrow biopsies.
❖ Fixes and dehydrates at the same time

5.2 ISOPROPYL ALCOHOL

❖ Used for fixing touch preparations.

5.3 ETHYL ALCOHOL

5.4 CARNOY’S FLUID

❖ Recommended for fixing chromosomes, lymph glands and urgent biopsies

❖ Used to fix brain tissues.
❖ Most rapid fixative
5.5 NEWCOMER’S FLUID
❖ Recommended for fixing mucopolysaccharides and nuclear proteins
❖ Acts as both as a nuclear and histochemical fixative

6. OSMIUM TETROXIDE (OSMIC ACID)

6.1 FLEMMINGS SOLUTION

❖ Most common chrome-osmium acetic acid fixative use.

❖ Excellent fixative for nuclear structures
❖ Permanently fixes fat

6.2 FLEMMING’S SOL WITHOUT ACETIC ACID

❖ Recommended for cytoplasmic structure paticulary6 the mitochondria

7. TRICHLOROACETIC ACID

❖ May be used as a weak decalcifying agent

8. ACETONE

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❖ Recommended for the study of water diffusible enzymes esp. lipase and phosphatase
❖ Used in fixing brain
❖ Dissolves fats

SECONDARY FIXATION

❖ Is the process of placing an already fixed tissue in a second fixative in order:

✔ To facilitate and improve the demonstration of particular substances
✔ To make special staining technique possible (with secondary fixative acting as mordant)
✔ To ensure further and complete hardening and preservation of tissues

POST CHROMATIZATION

❖ A form of secondary fixation whereby primarily fixed tissue is placed in aqueous sol of 2.5-3% potassium
dichromate for 24 hrs to act as mordant for better staining effects and to aid in cytologic preservation of
tissues.

WASHING OUT

❖ Process of removing excess fixative from the tissue after fixation.

❖ Several solution may be used:
✔ Tap water – used to remove
● Excess chromates from tissues fixed in kellys , zenkers and flemming solution
● Excess formalin
● Excess osmic acid
✔ 50-70% alcohol – used to wash out excess amount of picric acid (bouins solution)
✔ Alcoholic iodine – used to remove excess mercuric fixatives

FACTORS THAT AFFECT FIXATION

A. RETARDED BY
✔ SIZE AND THICKNESS OF THE SPECIMEN
✔ PRESENCE OF MUCUS
✔ PRESENCE OF FAT
✔ PRESENCE OF BLOOD
✔ COLD TEMPERATURE

B. ENGHANCEDBY

✔ SIZE AND THICKNESS OF THE SPECIMEN

✔ AGITATION

DECALCIFICATION

❖ Procedure whereby calcium or lime salts are removed from tissues following fixations.
❖ Recommended ratio of fluid to tissue volume for decalcification is20:1

METHODS OF DECALCIFICATION

1. ACID
I. NITRIC ACID
❖ Most common and the fastest decalcifying agent used so far. However it inhibits nuclear stain
and destroys tissues especially in concentrated solutions.

A. Aqueous nitric add solution 10%

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✔ Recommended for urgent biopsies and for needle and small biopsy specimens to
permit rapid diagnosis within 24 hrs. or less.
B. Formol – nitric acid
✔ Rapid acting; recommended for urgent biopsies
✔ Produce yellow color; remedy: neutralizing tissue in 5% sodium sulfate and
washing in running tap water for atleast 12 hrs. addition of 0.1% urea to pure
concentrated nitric acid will also make discoloration disappear without affecting
the efficiency of the decalcifying agent.
C. Perenyi’s fluid
✔ Recommended for routine purposes
✔ Nuclear and cytoplasmic staining is good.
✔ Maceration is avoided due to the presence of chromic acid and alcohol
✔ Not recommended for urgent work.
✔ Composed of chromic acid, ethyl alcohol and nitiric acid
D. Phloroglucinnitiric acid
✔ Most rapid decalcifying agent.
II. HYDROCHLORIC ACID
A. Von Ebner’s fluid
✔ Moderately rapid decalcifying agent.
✔ Does not require washing out before dehydration
✔ Recommended for teeth and small pieces of bone.
III. FORMIC ACID
✔ May be used as both fixative and decalcifying agent.
✔ Recommended for routine decalcification of postmortem research tissues.
B. FORMIC ACID – SODIUM CITRATE SOLUTION
✔ Recommended for autopsy materials, bone marrow, cartilage.
IV. TRICHLOROACETIC ACID
✔ Very slow acting; weak decalcifying agent
✔ Suitable for small spicules of bone.
V. SULFUROUS ACID
✔ Is a very weak decalcifying solution suitable only for minute pieces of bone.
VI. CHROMIC ACID (FLEMMING’S FLUID)
✔ May be use as both fixative and decalcifying agent
✔ Suggest for minute bone specimens.
✔ Consider as carcinogenic
VII. CITRIC ACID – CITRATE BUFFER SOL.
✔ pH 4.5
✔ excellent nuclear and cytoplasmic staining.
✔ Too slow
✔ Contains chloroform as preservatives.
2. CHELATING AGENTS
❖ Substances which combine with calcium ions and other salts like iron and magnesium.
I. EDTA (VERSENE)
✔ Most common chelating agent
✔ Recommended for detailed microscopic studies
✔ Very slow decalcifying agent
✔ For small specimens: 1-3 weeks
✔ 6-8 weeks or longer totally decalcifies dense cortical bone.
✔ Excellent for immunohistochem or enzyme staining and electron microscopy.

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✔ Inactivates ALP; to restore, add magnesium chloride.

3. ION EXCHANGE RESIN
❖ Not recommended for fluids containing nitric acid or hydrochloric acid.
❖ The decalcifying agent is then added, usually 20-30 times of the volume of the tissue.
❖ The tissue will stay in solution 1-14 days.
4. ELECTROPHORESIS
❖ Most rapid method.

FACTORS INFLUENCE RATE OF DECALCIFICATION

1. Concentration and volume

❖ More concentrated, more rapidly; more harmful
2. Structure and temperature
❖ Ratio: 20:1
❖ Higher concentrations and greater amounts of fluid will increase the speed of the process.
3. Heat
4. Mechanical agitation
❖ Accelerates the rate of diffusion and speeds up the decalcification process.

MEASURING EXTENT OF DECALCIFICATION

1. Physical or mechanical test

❖ Touching or bending the tissue with the fingers.
❖ Inaccurate
2. X – ray or radiological method
❖ Very expensive; most ideal; most sensitive and most reliable method.
❖ Detects even smallest amount of calcium which appears opaque in axray plate.
3. Chemical method (calcium oxalate test)
❖ Simple, reliable and convenient method
❖ Cloudiness will signify incomplete decalcification.

POST DECALCIFICATION

❖ Involves lithium carbonate to wash the tissue after the decalcification is complete.
❖ Also decalcified tissue can rinse in running tap water to remove acids.

TISSUE SOFTENERS

1. Perenyi’s fluid
✔ May act as both decalcifying and tissue softeners
2. 4% aqueous sol
3. Molliflex
✔ Tissues immersed in molliflex appear swollen and soapy.

DEHYDRATION

❖ Starts by placing the fixed specimen in 70% to 95% to 100% ethyl alcohol.
❖ For delicate tissues, particularly embryonic tissues starting 30% ethyl alcohol
❖ The amount of dehydrating agent should not be less than 10 times the volume of the tissue to ensure
complete penetration of the tissue by the dehydrating agent.
1. ALCOHOL
❖ Anhydrous copper sulfate will accelerate dehydration by removing water from dehydrating fluid.
A blue discoloration will indicate a complete dehydration.

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1.1 ETHYL ALCOHOL

✔ Recommended for routine dehydration of tissues.
✔ Best dehydrating agent.
1.2 METHYL ALCOHOL
✔ Toxic dehydrating agent
✔ Used for blood and and tissue film for smear preparations
1.3 BUTYL ALCOHOL
✔ For plant and animal microtechniques
✔ Slow dehydrating agent
2. ACETONE
❖ Cheap, rapid acting dehydrating agent
❖ For urgent biopsies which dehydrates 30 mins to 2 hrs
❖ Limited for only small specimen
❖ Volatility and inflammable
❖ Most lipids are removed from tissues with this dehydrating agent.
3. DIOXANE (DIETHYLENE DIOXIDE)
❖ Excellent dehydrating agent and clearing agent
❖ Readily miscible with water and paraffin
❖ Tends tissue ribbon poorly
❖ Highly toxic in man and expensive
4. CELLOSOLVE (EHTYLENE GLYCOL MONOETHYL ETHER)
❖ Cellosolve dehydrates rapidly. The tissue may be transferred from water or normal saline directly
to cellosolve and stored in it for months without producing hardening or distortion.
5. TRIETHYL PHOSPHATE
❖ Used to dehydrate sections and smears following certain stains and produces minimum shrinkage.
6. TETRAHYDROFURAN (THF)
❖ Both dehydrates and clears tissue since it is miscible in both water and paraffin.

CLEARING

❖ De-alcoholization; process whereby alcohol or a dehydrating agent is removed from the tissue
and replaced with a substance that will dissolve the wax with which the tissue to be impregnated.
❖ Must be miscible with water, paraffin and mounting medium.

COMMON CLEARING AGENTS

1. Xylene (Xylol)
❖ Colorless clearing agent, commonly used in histology laboratories.
❖ Miscible with absolute alcohol and paraffin
❖ It is cheap; used for celloidin sections
❖ Highly inflammable
❖ Not suitable for nervous tissues and lymph nodes
❖ Xylene turns milky when an incompletely dehydrated tossue is immersed in it.
2. Toluene
❖ May be used as a substitute for xylene and benzene.
❖ More expensive
❖ Miscible with absolute alcohol and paraffin
3. Benzene
❖ Preffered by some as a clearing agent in the embedding process of tissues because it penetrates
and clears tissue rapidly.

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❖ Carcinogenic, may damage the bone marrow resulting aplastic anemia

❖ Miscible with absolute alcohol
4. Chloroform
❖ does not make tissue translucent
❖ recommended for tough tissues (skin, fibroid, decalcified tissues)
❖ toxic to liver
❖ tissues tend to float in chloroform to avoid this wrapped the tissues with absorbent cotton gauze to
facilitate sinking of the section in solution.
❖ Miscible with absolute alcohol
5. Cedarwood oil
❖ Recommended for central nervous system tissues and cytological studies. (particularly smooth
muscles)
❖ Requires two changes in clearing solution
❖ It makes tissue transparent
❖ Extremely slow clearing agent
6. Aniline oil
❖ Recommended for clearing embryos, insects and very delicate specimens.
7. Clove oil
❖ Unsuitable for routine clearing process
8. Carbon tetrachloride
❖ Properties are very similar to chloroform
9. Methyl benzoate and methyl salicylate
❖ Double embedding technique is needed.

IMPREGNATION (INFILTRATION)

❖ Process whereby the clearing agent is completely removed from the tissue and replaced by a medium that
will completely fill the cavities, thereby giving a firm consistency to the specimen and allowing easier
handling and cutting suitably thin sections without any damage to the tissue and its cellular components.
1. PARAFFIN WAX IMPREGNATION
✔ Simplest, most common and best embedding medium, used for routine processing.
✔ Very rapid
✔ Prolong impregnation will cause excessive shrinkage and hardening making cutting of sections
difficult.
✔ Not recommended for fatty tissues.
✔ For routine work melting point is : 56-58 degcel
✔ For the lab temp between 15-18 degcel melting point is: 50-54 degcel
✔ For the lab temp between 20-24 degcel melting point is 54-58 degcel

METHODS OF PARAFFIN WAX IMPREGNATION

A. MANUAL PROCESSING

✔ Four changes of wax are required at 15 minutes intervals.

B. AUTOMATIC PROCESSING

✔ Use auto-technicon which fixes, dehydrates, clears and infiltrates tissues.

✔ 2 -3 changes of wax are required.

C. VACUUM EMBEDDING

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✔ Involves the wax impregnation under negative atmospheric pressure inside an embedding oven to hasten
removal of air bubbles and clearing agent.
✔ Most rapid
✔ Recommended in urgent biopsies

SUBSTITUTES FOR PARAFFIN WAX

1. PARAPLAST

✔ Mixture of highly purified paraffin and synthetic plastic polymers

✔ Melting point: 56-57 deg cel
✔ More elastic and resilient
✔ For bones and brains

2. EMBEDDOL

✔ Synthetic wax substitute similar to paraplast

✔ Melting point is : 56-58 deg cel

3. TISSUE MAT

✔ Contains rubber

4. ESTER WAX

✔ Has a lowering melting point 46-48 degcel

✔ Harder than paraffin

5. WATER SOLUBLE WAX (CARBOWAX)

✔ Melting point: 38-42 degcel or 45-46 degcel

2. CELLOIDIN IMPREGNATION
⮚ Purified form of nitrocellulose soluble in many solvents
⮚ Suitable for specimens with large hollow cavities which tend to collapse for hard and dense
tissues such as bones teeth and for large tissue sections such as whole embryos.
⮚ Recommended for processing neurological tissue
⮚ Very slow

TWO METHODS FOR CELLOIDIN IMPREGNATION

1. WET CELLOIDIN METHOD

✔ Recommended for bones teeth large brain and whole organs

2. DRY CELLOIDIN METHOD

✔ Recommended for whole eye sections

✔ Does not use alcohol due to the presence of cedarwood oil in the block.
3. GELATIN IMPREGNATION
✔ Rarely used except for histochem and enzyme studies
✔ Used for delicate specimens and frozen tissue
✔ Water soluble
✔ Tissue should not be more than 2-3mm thick; add 1% phenol to prevent molds
✔ Excess gelatin may be removed by floating the sections to paper and trimming them with
scissors.

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EMBEDDING

✔ After impregnation, the tissue is placed into a mold containing the embedding medium and this medium
allows it to solidify.
✔ In this process orientation is very important.
✔ Temperature of melted paraffin is 5-10 deg cel above the melting point
✔ Immersed in cold or ref temp to solidify

1. Leukhart’s embedding mold

✔ Contains two L- shaped strips of heavy brass.

2. Compound Embedding unit

✔ Made up of a series of interlocking plates resting on a flat metal base.

3. Plastic embedding rings and base molds

✔ Consist of special stainless steel base mold fitted with a plastic embedding ring, which later serves as the
block holder during cutting.

4. Tissue-tek

✔ Equipped with a warm plate to manage the impregnated specimen.

5. Disposable embedding molds

● Peel-away
● Plastic ice trays
● Paper boats

*Celloidin or nitrocellulose method – recommended for embedding hard tissues.

*double-embedding method-process in which tissue first infiltrates by celloidin and embedded in paraffin.

MICROTOMY

❖ Process by which processed tissue, most commonly a paraffin embedded tissue, is trimmed and cut into
uniformly thin slices or sections to facilitate studies under microscope.

ESSENTIAL PARTS OF MICROTOME

1. BLOCK HOLDER

✔ Tissue held in position.

2. KNIFE CARRIER AND KNIFE

✔ For actual cutting of tissue sections.

3. PAWL, RATCHET FEED WHEEL AND ADJUSTMENT SCREWS

✔ Line up the proper position with the knife.

5 KINDS OF MICROTOME

1. ROCKING (CAMBRIDGE) MICROTOME

✔ For cutting large blocks of paraffin embedded tissues.

✔ Invented by paldwell trefall in 1881

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✔ Simplest among the different types of microtome

✔ Cut the tissue passes to the knife edge in slightly curved plane in 10-12 u thickness

2. ROTARY (MINOT) MICROTOME

✔ For cutting paraffin embedded sections.

✔ Invented by Minot in 1885-1886
✔ Most common type used for both routine and research laboratories.
✔ Cut the tissue in 4-6 u thickness.

3. SLIDING MICROTOME

✔ For cutting celloidin embedded sections

✔ Developed by Adams in 1789
✔ 3.1 base sledge microtome- consists of two movable pillars holding the adjustable knife clamps.
✔ 3.2 Standard sliding – the blocks remain stationary, the knife is moved backward and forward during the
process of sectioning. It is for cutting celloidin sections.
✔ Most dangerous type of microtome.

4. FREEZING MICROTOME

✔ For cutting unembedded frozen sections.

✔ Invented by Queckett in 1848
✔ Used carbon dioxide to freeze the block holder and tissue evenly.

5. Cryostat or cold microtome

✔ Used in fresh tissue microtome

✔ Consist of rotary microtome
✔ For rapid diagnosis
✔ Maintained a temperature between -5 to -30 degcel average is -20 degcel.

6. ULTRATHIN MICROTOME

✔ Cut tissue at 0.5 micra

✔ For electron microscopy

MICROTOME KNIVES

PLANE-CONCAVE KNIFE One side of the knife is flat while the 25 mm in length
other is concave. Less concave are
for cutting celloidin embedded
tissue blocks. More concave is for
paraffin sections
BICONCAVE KNIFE Both sides concave, recommended 120 mm in length
for cutting paraffin embedded
sections
PLANE-WEDGE KNIFE Both sides are straight, for frozen 100 mm in length.
sections.

*BEVEL ANGLE – formed between the cutting edges; normally about 27-32 degcel.

HONING

❖ Removal of gross nicks on the knife edge

❖ 10-20 strokes; 20-30 strokes; heel to toe movement or direction; edge first.

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OIL STONES:
1. BELGIUM YELLOW- for manual sharpening; usually gives the best result.
2. ARKANSAS – gives more polishing effects.
3. FINE CARBORUNDUM - used only for badly nicked knives.
LUBRICANTS
1. MINERAL 2. CLOVE OIL
3. XYLENE 3. LIQUID PARAFFIN
*use to remove scattered small particles of stones and metals

STROPPING

❖ Process whereby the burr formed during honing is removed and the cutting edge of the knife is polished
and sharpened.
❖ Toe to heel direction; 40-120 double strokes are required.
❖ Use paddle strop made of horse leather that is usually treated with vegetable oil.
❖ Mineral oil is not recommended and should never be touched or in contact in strope it will tend to blister
and destroy the leather.

TRIMMING – cutting of excess wax from the block.

CLEARANCE ANGLE – between the facet and tissue block; 0-15 degcel
CAMEL HAIR BRUSH- Use to pick up the complete ribbons.
*Sections are floated out to a water bath set at 45-50 degcel or 6-10 degcel lower than the melting point
of the wax used for embedding tissue.
*Sections should not be left in the water bath for a long time. 30 secs will be enough to avoid distortion.

DRYING TECHNIQUES

❖ PARAFFIN OVEN – maintained a temperature 2-5 deg cel above the melting
point of the paraffin used.
❖ Thermostatically controlled incubators may be used at 37 deg cel and 45-55 deg cel

ADHESIVES

1. MAYER’S RGG ALBUMIN

✔ Most commonly used ; it is very easy to make; convenient and inexpensive
✔ Equal amount of egg white and glycerin
2. DRIED ALBUMIN
✔ DRIED ALBUMIN+nacl= dried albumin
✔ Addition of thymol to prevent growth of molds
3. GELATIN 1%
✔ Gelatin+ distilled water+glycerol+phenol crystals
4. GELATIN FORMALDEHYDE MIXTURE
✔ 1% gelatin + 2% formaldehyde
5. STARCH PASTE
✔ Powdered starch+ distilled water+ hydrochloric acid+ thymol
6. PLASMA
✔ Readily available from outdated blood storeed in blood banks or hematology.
✔ Dispensed it into sterile tubes of 0.5 ml each.
7. POLY-L-LYSINE
✔ Must be used within a few days after they are prepared, since its effectiveness slowly
decreases in time.
8. APES (3-aminopropytriethylxysilane)

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✔ Better than poly-l-lysine, it can be stored for a long time without losing its adhesiveness.

STAINING

✔ Applying dyes on the section to see and study the architectural pattern of the tissue and physical
characteristics of the cells.

MAJOR GROUPS OF STAINING

1. HISTOLOGICAL STAINING – process whereby tissue constituents are demonstrated in sections by

direct interaction with a dye. Used to demonstrate the general relationship of tissues and cells with
differentiation of nucleus and cytoplasm.
2. HISTOCHEMICAL STAINING (HISTOCHEMISTRY) – process whereby various constituents of
tissues are studied thru chemical reactions.
*PERL’S PRUSSIAN BLUE – REACTION FOR HGB
*PAS – FOR CARBOHYDRATES

3. IMMUNOHISTOCHEMICAL STAINING - immunologic and histochemical techniques that allow

phenotypic markers to be detected using an enzyme labeled antibodies.

METHODS OF STAINING

1. DIRECT STAINING – giving color to the sections by using aqueous or alcoholic dye solutions.
Example: methylene blue; eosin
2. INDIRECT STAINING – whereby the action of the dye is intensified by adding another agent or
mordant.
*MORDANT- serves as a link or bridge between the tissue and dye.
Example: Potash alum/potassium alum in erlichshematoxylin.
*ACCENTUATOR – accelerates or hastens the speed of staining reaction.
Example: potassium hydroxide in loeffler’s methylene blue and phenol in carbolthionine and
carbolfuchsin.
3. PROGRESSIVE STAINING – tissue elements are stained in definite sequence and staining. No
decolorization required.
4. REGRESSIVE STAINING – tissue is first overstained and the excess stain is removed or decolorized
from unwanted parts of the tissue until the desired color is obtained.
5. DIFFERENTIATION/DECOLORIZATION – removal of excess stain from the tissue during
regressive staining.
6. METACHROMATIC STAINING – use of specific dyes which differentiate particular substances by
staining them with a color that is different from that of the stain itself.
7. COUNTERSTAINING – application of a different color stain to provide contrast and ack ground to the
staining of the structural components to be demonstrated.
*MOST COMMON COUNTER STAINS
o CYTOPLASMIC STAINS
RED YELLOW GREEN
Eosin Y Picric acid Light green SF
Eosin B Orange G Lissamine green
Phloxine B Rose Bemgal

● NUCLEAR STAINS
RED BLUE
Neutral red Methylene blue
Safranin O Toluidine blue
Carmine Celestine blue
Hematoxylin

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8. METALLIC IMPREGNATION – process where specific tissue elements are demonstrated not by
stains but by colorless solutions of metallic salts which are thereby reduced by the tissue, producing an
opaque usually black deposit on the surface of the tissue or bacteria.

9. VITAL STAINING – selective staining of living cell constituents, demonstrating cytoplasmic

structures by phagocytosis of the dye particle; nucleus is resistant to vital stain therefore it is not
demonstrated.

*demonstration of nucleus in vital staining indicates death cells.

Intravital staining – staining of living cells is done by injecting the dye top any part of the animal body.
Common dyes used are lithium, carmine and india ink

Supravital staining – used to stain living cells immediately after removal from the living body.
Common dyes used are:

*neutral red- best vital dye

*janus green- recommended for mitochondria

ROUTINE H AND E STAINING (REGRESSIVE STAINING)

1. First xylene bath for 3 mins; for decolorization removal of excess paraffin wax

2. Transfer to second xylene bath for 2-3 mins

3. Immerse in the first bath of absolute ethyl alcohol for 2 minutes

4. Transfer to a bath of 95% ethyl alcohol for 1 to 2 minutes for hydration descending grades of alcohol

5. Rinse in running water for 1 minute

6. Stain with harris alum hematoxylin for 5 mins. (primary dye)

7. Wash in running tap water to remove excess stain

8. Differentiate in 1% acid alcohol for 10-30 secs.

9. Rinse in tap water

10. Blue in ammonia water average of 5mins; to intensify the color of the nucleus

11. Wash in running tap water for 5 mins.

12. Counterstain with 5% aqueous eosin for 5 mins. If alcoholic stain is used time is reduced to 30 secs or 1
minute.

13. Dehydrate, clear and mount.

RESULT:

*nuclei: blue to black *karyosome- dark blue *cytoplasm- pale pink

*RBC, keratin- bright orange-red *cartilage- pink *calcium and calcified bones-
purplish blue *muscle fibers- deep pink

1. NATURAL DYES – obtained from plants and animals , previously utilized from dyeing wool and cotton.

1.1 HEMATOXYLIN- derived by extraction from the core or the heartwood of a Mexican tree known as
hematoxylin campechianum.

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A. Alum hematoxylin – recommended for progressive staining of tissues, stain nuclei but leave the background
tissue unstained. It also uses alum or potassium aluminum as mordant.

*Ehrlich’s hematoxylin – ripened by sodium iodate

* Harris hematoxylin – ripened by: Mercuric oxide

*Cole’s hematoxylin- ripened by an alcoholic solution

*Mayer’s hematoxylin- ripened by sodium iodate

B. Iron hematoxylin - it used for photomicrography

*Weigert's solution- use ferric ammonium chloride as mordants

*Heidenhain’s solution – use ferric ammonium sulfate (iron alum) as mordants

C. PHOSPHOTUNGSTIC ACID HEMATOXYLIN (PTAH) – demonstrates structures in paraffin as well as

celloidin and frozen sections. Staining is usually 12-24 hours. Use 1% aqueous phosphotungstic acid as
mordant.

1.2 COCHINEAL DYES – is an old dye extracted from the female cochineal bug (coccus cacti).

*Picrocarmine- used in neuropathological studies

*best’s carmine- used for demonstration of glycogen

1.3 ORCEIN – vegetable dye extracted from certain lichens. It is used mainly in staining of elastic fibers.

2. SYNTHETIC DYES – known as “coal tar dyes” derived from hydrocarbon benzene collectively known as
aniline dyes.

3. LYSOCHROMES – oil soluble dyes.

3.1 SUDAN BLACK – most sensitive of the oil soluble dyes. It has a much greater affinity for phospholipids
than other lysochrome.

3.2 SUDAN IV– recommended for staining neutral lipids or triglycerides.

3.3 SUDAN III – first sudan dye introduced in immunohistochemistry; good stain for central nervous system.

MOUNTING – To preserve and support a stained section for light microscopy, it is mounted on a clear glass
slide, and covered with a thin glass coverslip. The slide and coverslip must be free of optical distortions, to
avoid viewing artifacts.

CHARACTERISTICS OF GOOD MOUNTING MEDIUM

1. A refractive index should be as near as possible to that of the glass which is 1.518.
2. Should be freely miscible with xylene and toluene
3. Should not dry quickly
4. Should not crack or produce artifactual granularity upon drying.
5. Should not dissolve out or fade tissue sections.
6. Should not cause shrinkage and distortion of tissues
7. Should not leach out any stain or affect staining
8. Should not change in color or pH
A. AQUEOUS MOUNTING MEDIA
1. WATER – has a low refractive index , moderately transparent; good for temporary
mounting.
2. GLYCERIN – also be used as a preservative , has a high refractive index . This is a very
suitable for semi permanent mounting medium with a refractive index of 1.46.

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3. FARRANT’S MEDIUM – does not need to be heated before use; require ringing.
Refractive index of 1.43
4. APATHY’S MEDIUM – does not require ringing; recommended for methylene blue stained
nerve preparations and as a general purpose aqueous mountant. Refractive index of 1.52
5. BRUN’S FLUID – for mounting frozen sections.

B. RESINOUS MOUNTING MEDIA

1. CANADA BALSAM - natural resin extracted from the canabian tree, (ABUS BALSAMEA).
Recommended for whole mounts and for thick sections because it does not shrink much. It is
miscible with xylene. Refractive index of 1.524
2. DPX – recommended for small tissue sections but not for whole mounts because of
shrinkage produced on drying. Refractive index of 1.532
3. XAM – synthetic resin mixture , dries quickly without retraction and preserves stains well.
Refractive index of 1.52
4. CLARITE – soluble in xylene. Refractive index of 1.544

ANNE GRACIELLA F. ORIG, RMT MARCH 2025!

IF YOU REALLY WANT TO BE A RMT, WORK AND FIGHT FOR IT!

GODBLESS AND HAPPY ARAL! ☺

REFERENCES:

HISTOPATHOLOGIC TECHNIQUES SECOND EDITION

BY JOCELYN H. BRUCE-GREGORIOS, M.D

NOTES FROM SIR HANZ

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