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Файл:Ординатура / Хирургия / Библиотека им академика М.И. Перельмана / Книга_5535_Библиотеки_им_академика_М_И_Перельмана.pdf
1176
Add concentrated ammonia drop-by-drop with frequent mixing until the formed
precipitate just redissolves. Add 5 mL of 3.1% aqueous sodium hydroxide and mix.
A precipitate will form that gradually dissolves upon the addition of ammonia, dropby-drop as before. Stop when there are only a few precipitate granules remaining. Make
up the nal volume to 50 mL with distilled water.
1. Take the sections to water (dip through decreasing strength of alcohol).
2. Treat the section with acidied potassium permanganate solution for 5 min.
3. Wash o the slides (sections) in water for about 2 min.
4. Bleach the slides (sections) with oxalic acid solution for approximately 1 min or until
colourless.
5. Wash the slides (sections) completely in water and rinse in distilled water.
6. Treat the sections with iron alum solution for 5 min.
7. Wash the sections well in several changes of distilled water.
8. Treat the section with ammoniacal silver solution for 4–5 s with agitation of the slide.
9. Wash the section well in several changes of distilled water.
10. Reduce in 10% formalin in tap water for about 30–60 s with agitation of slide (section).
11. Wash the slides in tap water and then rinse with distilled water.
12. Treat the sections with 5% sodium thiosulphate solution for 5 min.
13. Wash the slides (sections) in distilled water.
14. Counterstain the sections, if desired, in 1% aqueous neutral red for 5 min.
15. Dehydrate, clear and mount the sections as desired.
• Reticulum bres—black (some pigments like melanin are also impregnated)
• Collagen—yellow-brown
• Background—clear if not counterstained and red if counterstained
Medical Laboratory Technology: Volume 3
Elastic fibres
Elastic bres are branching bres of varying size and diameter that consist of the protein
and glycoprotein microbrils. They are easily seen in tissue sites such as dermis of
skin, lung, heart and blood vessel walls. Elastic bres may be increased because of a stromal
reaction to the presence of tumour cells, as in or secretion of tumour cells, as in
the rare elastoma dorsi.
is required to identify changes in the
that may be increased due to hypertension or lost because of a degenerative process.
The ner elastic bres are not easily delineated in H&E preparation unless special stains are
used. Celloidinization of sections prior to staining, particularly of skin and blood vessels, is
desirable as they may detach from the slide.
Wiegert’s Resorcin-fuchsin stain There are several variations of the technique described
here. The rationale of these variations is obscure. Most of these methods, however, show a
remarkable selectivity for elastic bres and give the best demonstration of ne bres. The
techniques tend to be slow and the solutions are time-consuming to prepare. These solutions
stain well at room temperature and may be used repeatedly. Some variations in staining
avidity will be found with dierent dye batches so a variation in staining times will occur.
Staining at room temperature for longer periods gives beer results than a shorter time at
56°C. Pre-treatment with an acidied permanganate-oxalic acid solution sequence gives a
clearer background. Fixation is not critical. Any well-xed tissue can be used.
• Oxalic acid solution (5%, aqueous, i.e., 5 g oxalic acid in 100 mL water)
Laboratory Techniques in Histology
• Acidied potassium permanganate solution:
■ 0.3% aqueous potassium permanganate (3 g potassium permanganate in 100 mL water)
■ 3% sulphuric acid (3 mL concentrated sulphuric acid mixed with 97 mL distilled water)
Mix equal parts immediately prior to use. These can be conveniently kept as stock
solutions.
• Acid alcohol—Dierentiator
Conc. hydrochloric acid 1 mL
70% alcohol 99 mL
• Ammonia water (2%, aqueous, i.e., 2 mL of concentrated ammonia in 98 mL of water.
Watch the fumes. Work inside the hood with fan on.)
• Neutral red (1%, w/v, aqueous, i.e., 1 g in 100 mL water)—counter stain
• Weigert’s resorcinol fuchsin solution
Basic fuchsin 2 g
Resorcinol 4 g
Distilled water 200 mL
Ferric chloride, 30% (w/v, aqueous) 25 mL
95% alcohol 200 mL
Conc. hydrochloric acid 4 mL
Dissolve the basic fuchsin and resorcinol in the distilled water, bringing to the boil in
an evaporating dish. While boiling, slowly add the aqueous anhydrous ferric chloride
solution, stirring continuously. Continue for approximately 5 min. Cool and lter into
a conical ask taking care that all the precipitate is collected. Discard the ltrate, dry
the ask, and add the dried lter paper containing the precipitate to the ask. Add the
alcohol and heat gently on a hot plate until the precipitate is dissolved. Remove the lter
paper, add the concentrated hydrochloric acid, cool and lter. Make up the nal volume
to 200 mL by pouring fresh 95% alcohol through the used lter paper. Filter before use.
This solution keeps well for several months. The puried basic fuchsin commonly used
for Scru’s reagent gives inferior results.
1177
1. Take the sections to water by passing through decreasing strengths of alcohol.
2. Dip the slides in acidied potassium permanganate solution for 5 min.
3. Wash in water and bleach with 5% oxalic acid for approximately 1 min.
4. Wash well with water to remove the bleach and nally rinse in alcohol.
5. Stain in Weigert’s resorcinol fuchsin solution for 1–3 h at room temperature. The time will
vary per batch and type of solution. Check with microscope and stain until elastic bres
are black. Save solution. If over stained, rinse in ammonia water for 3 min. This will
remove excess stain.
6. Dierentiate the stain in acid–alcohol until the background is clear of stain.
7. Wash well in tap water.
8. Counterstain as required with neutral red stain (or eosin or van Gieson). Save solution.
If over stained, water will remove excess stain.
9. Dehydrate, clear and mount as desired.
• Elastic bres—blue-black
• Background—red (or according to the colour of the counterstain used)
Verhoe’sstain This is a rapid method for staining elastic bres a strong black colour. The
disadvantage is that some expertise is necessary to stain the ner elastic bres and obtain a
well-dierentiated background. The principle of the technique is rather obscure. Both iodine
1178
and ferric chloride, with which the haematoxylin is combined, are oxidizing agents and it is
this property that probably accounts for the production of a black dye with cationic properties
(nuclei are also stained), rather than a simple dye-mordant tissue mechanism. Any well xed
tissue can be used.
• Verhoe’s solution:
For best results, make up solutions the same day they are to be used. Freshly prepared
solutions give stronger staining results.
Solution A:
Haematoxylin 5 g
Absolute alcohol 100 mL
Dissolve with the aid of heat. Cool and lter.
Solution B:
Ferric chloride 10 g
Distilled water 100 mL
Solution C (Lugol’s iodine solution):
Iodine 1 g
Potassium iodide 2 g
Distilled water 100 mL
Add 8 mL of solution B into 20 mL of solution A and to that add 8 mL of solution C.
Note Follow the sequence as given.
• Ferric chloride solution (2%)
Medical Laboratory Technology: Volume 3
1. Take sections (dehydrate)—Dip through increasing concentrations of alcohol.
2. Stain with freshly made Verhoe's solution until the sections are black (15–45 min).
3. Wash in distilled water.
4. Dierentiate the stain in 2% ferric chloride with agitation, only for a few minutes. Check
dierentiation by rinsing in distilled water and examining under the low power of the
microscope. Discontinue when elastic bres and nuclei are black and other tissues are
gray or weakly stained. Should the section be over dierentiated, it may be returned to
Verhoe's solution for further staining.
5. Wash in water and then in alcohol for approximately 5 min to remove the iodine
colouration of the background.
6. Wash in water and counterstain in van Gieson’s stain for 1–2 min.
7. Dehydrate, clear and mount as desired.
• Elastic bres—black
• Nuclei—gray to black
• Background—according to counterstain
StainS fOr particuLar SubStanceS
Staining of carbohydrates, amyloids, pigments and minerals (iron and calcium) and
microorganisms is occasionally done in the histopathology laboratory for the diagnosis of
specic pathologic conditions. Staining of fat is done with frozen sections. This is discussed
separately under the frozen section technique.
Laboratory Techniques in Histology
1179
Carbohydrates
A simple carbohydrate molecule is a monosaccharide such as glucose that plays a central
role in nutrition but is dicult to demonstrate within tissues. Glycogen, on the other hand,
is a polysaccharide, with long chain sugar molecules. It is stored in liver and in muscle
and available to the body in times of its need. Shorter and more complex chains of sugar
molecules may be covalently linked to protein and lipid molecules forming glycoproteins
and glycolipids, respectively. The carbohydrate component modies the function of the
main protein or lipid molecules. Mucin, present in the connective tissue is an amino sugar.
Mucopolysaccharides are group of compounds composed of protein and complex sugars
oen referred as The staining and identication of these polysaccharides
and mucopolysaccharides have contributed greatly to our understanding of tissue structures
such as liver (glycogen), heart, striated muscle, gastrointestinal tract glands, respiratory lining
cells and others. The periodic acid-Schi stain is generally presumed to be for a demonstration
of carbohydrate but the stain is rather non-specic.
Periodic Acid-Schiff (PAS) stain
Periodic acid is a strong oxidizing agent and under the controlled conditions of the
staining reaction, it reacts with the aldehyde group of the carbohydrates without allowing
over-oxidation. Schi reagent then reacts with the product. A red or purple-red colour
indicates a positive PAS reaction. PAS positive substances include a host of organic compounds
(amyloid, cerebrosides, glycogen, etc.), microorganisms (amoebae, mycobacterium, fungi,
etc.), body tissues and cells.
The use of post-Schi sulphite rinses to reduce background colouration is not necessary,
provided washing in running water is thorough and that the alkalinity of the tap water is not
too high.
• Periodic acid solution (1%, w/v, aqueous):
Periodic acid crystals 1 g
Distilled water 100 mL
• Schi’s reagent:
Basic fuchsin 1 g
Distilled water 200 mL
Potassium or sodium metabisulphite 2 g
Cone, hydrochloric acid (Analar grade) 2 mL
Decolourizing charcoal 2 g
• Harris’s haematoxylin solution: Preparation of this reagent is described earlier (see routine).
• Light green counterstain
Light green 100 mg
Acetic acid (0.1%) 100 mL
1. Sections to distilled water—take the slides through decreasing strength of alcohol.
2. Treat with solution for 5 min (longer time, 10 min, for basement
membranes).
3. Rinse well in distilled water.
4. Treat with reagent for 15 min.
5. Wash in running tap water for 5–10 min. This intensies the colour reaction.
6. Stain nuclei with as counter stain or light green.
7. Dierentiate and blue.
8. Dehydrate (take through decreasing strength of alcohol).
9. Clear and mount as desired.
1180
• PAS-positive materials (sections that picked up the stain): These sections bear magenta
which is rose to purple red. Tissues picking up this colour include glycogen, mucin,
reticulin, brin of thrombi, hyaline of arteriosclerosis, most basement membranes,
colloid pituitary stalk and thyroid, amyloid inltration and others.
If light green is used as the counterstain, the section will have a pale background of the
counterstain.
• Nuclei—blue or blue-black
PAS staining is mainly used for staining structures containing a high proportion of
carbohydrates. This includes glycogen, glycoproteins, proteoglycans typically found in
connective tissues, mucus and basement membranes. It is helpful to study the biopsies of
liver and kidney. It is able to recognize certain glycogen storage diseases in striated muscles
and also in suspected fungal infections.
PrincipleofAction:Some products of carbohydrate origin like glycogen, neutral mucins and
glycoproteins, that has adjacent hydroxyl group, react with periodic acid-Schi. This reaction
produces a bright magenta coloration in areas of tissue containing these carbohydrates.
Periodic Acid 0.5% 500 mL
Schi Reagent 500 mL
Gill II Hematoxylin 500 mL
StorageandStability:Follow the instruction provided by the manufacturer. Do not use the
kit aer the expiration date.
Procedure:Follow manufacturer’s instruction.
Medical Laboratory Technology: Volume 3
Amyloids
Amyloid is a starch-like material, principally a glycoprotein, formed by the combination
of carbohydrate with protein. In various pathologic conditions abnormal quantities of
glycoprotein may occur in organs and tissues, leading to amyloidosis with pathologic lesions.
Some of the commonly used dyes to stain amyloid are Congo red, toluidine blue and crystal
violet. These stains show metachromasia. In other words, the colour that appears in the
section following these stains is dierent from the dominant colour of these stains. Hence, to
avoid confusion, these stains must be used without a counterstain. In a section stained with
toluidine blue, the metachromatic areas will be red-purple and the remaining tissue various
shades of blue. These stains tend to fade and therefore the sections should be examined soon
aer they are stained. Stained sections should be examined immediately as many mounting
media reverse the metachromasia. Another disadvantage of these stains is that they are not
truly specic towards amyloid.
Congo red stain
Congo red, an anionic dye, is commonly used for the staining and demonstration of amyloid
or glycoprotein in microscopic tissue sections. Bennhold’s Congo red stain is simple and
reliable and the stained sections can be kept indenitely, since it is dehydrated and mounted
in a permanent mounting medium.
An important feature of Congo red staining is the red to green birefringence seen when
using polarized light, although this is not very specic. Hence a weak birefringence may
require other conrmatory techniques.
Carnoy’s uid or absolute alcohol is used as the xative. Ten percent formalin or Zenkerformal may be used.
Laboratory Techniques in Histology
Alkaline Congo red technique This is a progressive method requiring no dierentiation
step. Salts act as ionic competitors for the dye, and background (polar) staining is eliminated;
only the non-polar binding of Congo red occurs. The main disadvantage of the method lies in
its complexity and the short bench-life of the solutions used.
• Harris’s alum haematoxylin solution: Described earlier.
• Sodium hydroxide solution (1%, w/v, aqueous): Keep in air-tight plastic bole. Discard the
solution when turbid.
• Stock alcoholic sodium chloride: Saturated sodium chloride in 80% alcohol. This is a stable
solution that keeps well. It is used as stock for the following reagent.
• Working solution: This solution is good for 15 min.
Stock solution 50 mL
1% aqueous sodium hydroxide 0.5 mL
Mix and lter, use within 15 min.
• Alkaline Congo red solution
A. Stock solution:
Congo red in 80% alcohol saturated with sodium chloride. Filter and keep in a tightly
stoppered container. The stock solution will be stable for several months.
B. Working solution:
Stock solution (Congo red, Solution A) 50 mL
1% aqueous sodium hydroxide (1 g sodium hydroxide in 100 mL water) 0.5 mL
1181
1. Section to water (take through decreasing concentrations of alcohol).
2. Stain the nuclei with Harris’s alum haematoxylin solution.
3. Dierentiate and blue.
4. Treat with the alcoholic sodium chloride-hydroxide solution for 20 min; drain.
5. Stain with the Congo red solutions for 20 min.
6. Rinse with alcohol.
7. Dehydrate (bath through series of increasing concentrations of alcohol).
8. Clear and mount as desired.
Amyloid—orange-red
Nuclei—blue
Background—clear
Toluidine blue stain
Toluidine blue is a basic dye that stains many tissue components—including amyloids—an
orthochromatic blue colour. Under polarized light, amyloid is distinguished by its striking
dark red birefringence. Occasional amyloid deposits, especially those of endocrine origin, are
negative with this method and minimal deposits are sometimes dicult to visualize.
• Isopropanol (50%, v/v, aqueous)
• Toluidine blue solution: 1% solution in 50% isopropanol
1182
1. Well-paranized sections to water, removing xation pigment, where necessary.
2. Stain in toluidine blue solution for 30 min at 37°C (in oven).
3. Blot section carefully then place in absolute isopropanol for 1 min.
4. Clear and mount as desired.
Amyloid and many other tissue components stain an orthochromatic blue colour, but when
examined under polarized light amyloid gives a dark red birefringence.
Medical Laboratory Technology: Volume 3
Lipids
Lipids can be dened as substances that are insoluble in water and may be extracted from
the tissues by organic solvents. Lipids are normal constituents of tissues, found in adipose
tissue as stored lipid for energy production, or as specialist lipid structure such as myelin.
Lipids are rarely found in a pure state in tissue sections; they are usually in combination
with carbohydrates in glycolipids or with proteins in lipoproteins. The demonstration of
lipid in the routine laboratory is infrequently called for and, unless dealing with a suspected
lipid storage disorder, is unlikely to require precise identication of the type present. The
accumulation of fat in some tissues may be diagnostic of certain pathologic conditions. Fat
embolism (obstruction of a blood vessel) is not uncommon in bone injuries and fractures.
Extraneous examples of conditions requiring lipid staining include aortic atheroma and lipid
pneumonia of the lungs (due to aspiration of fay material from the oral cavity). Certain
tumours, too, contain signicant quantities of fat and this can sometimes be used to advantage
in their recognition.
Fat is soluble in alcohol. Thus, it is not possible to use sections cut from ordinary paranembedded blocks to demonstrate fat, as the fat will have been dissolved by the alcohols used
in the embedding procedures.
In addition, there is no really good xative for lipids.
Formalin xes only a minority of lipids. It is inert towards simple lipids and allows appreciable
diusion and loss of phospholipids. Cryostat sections of fresh unxed tissue involve the
least loss of lipids. Lipids (broadly includes both simple fats and conjugated fats) may be
demonstrated histochemically by a variety of techniques. Of all the methods, the use of oil
soluble colorants is most popular. This includes oil red O and Sudan black stain. These will
be described here. Counterstaining presents problems as most dyes diuse or are bleached by
aqueous mountants. For the red lipid dyes haematoxylin is used, but it tends to fade in time.
Oil red O stain
The deep red staining of lipids by this method makes it one of the most popular techniques
for the identication of fat in the tissue sections. Always use a covered container and ample
amounts of solution when staining and use care in washing. Of the various alternative staining
solutions, the isopropanol variant is more commonly chosen, which is described here.
• Isopropyl alcohol (absolute and 60%, v/v, aqueous or 60 mL isopropyl alcohol and 40 mL
water)
• Acetic acid(l%, v/v, aqueous)
• Ammonia water (2%, v/v, aqueous)
• Oil red O stain:
Dissolve 0.5 g of oil red O dye in 200 mL of isopropyl alcohol (absolute). Warm
the solution in a long-necked container (2-L volumetric ask) in a 56°C water bath for
1 h. Cool.
Laboratory Techniques in Histology
is prepared prior to use by adding 4 parts of distilled water to 6 parts
of stock solution. Mix and stand for 10 min. Filter through a ne lter paper (Whatman
No. 42). This working dilution should be used within 2–4 h. Such saturated staining
solutions must be kept in air-tight staining vessels to avoid precipitation of the dye
because of the evaporation of the solvent.
• Counterstain: Harris’s haematoxylin or light green
• Glycerine jelly (mounting medium):
Gelatine 10 g
Distilled water 60 mL
Heat until gelatine is dissolved then add:
Glycerine 70 mL
Phenol 1 mL
1. Rinse frozen sections in water.
2. Rinse in 60% isopropyl alcohol.
3. Stain in oil red O for 10 min.
4. Wash briey in 60% isopropyl alcohol.
5. Wash gently in tap water.
6. Counterstain in Harris’s haematoxylin for 1–2 min.
7. Wash in water.
8. Blue in ammonia water: If sections are too dark when removed from the haematoxylin,
they may be dierentiated in 1% acetic water for a few seconds, and then blend into blue
in ammonia water.
9. Wash in water and mount in an aqueous mountant (glycerine jelly)
• Fat—orange to bright red
• Phospholipids—pink
• Nuclei—blue
• Other tissues—pale brown
1183
Sudan black stain
Sudan black is the most sensitive lipid stain known. In fact, it is too sensitive as a general fat
stain. Valid objections, however, exist to the use of acetone and ethanol as vehicles for these
dyes as these solvents remove a signicant portion of the lipids. Thus, small fat droplets are
likely to be dissolved out and escape detection. Isopropyl alcohol, propylene or ethylene
glycol is less objectionable.
• Sudan black solution: Saturated solution of Sudan black in 70% alcohol. It is made by
adding the dye into 70% alcohol in increasing amount until the dye cannot dissolve
(saturated).
• 2% carmalum solution:
Carminic acid 2 g
5% aqueous ammonium alum 100 mL
Salicylic acid or thymol 0.2 g
Add the carminic acid to the ammonium alum solution and dissolve by boiling for 1 h.
Cool and restore to original volume with distilled water. Add fungicidal agent (salicylic
acid or thymol) and mix thoroughly. Filter and use. The solution keeps quite well if
stored at 4°C.
• Glycerine jelly (aqueous mountant): Described earlier.
1184
1. Mount the sections on to slides and allow to dry.
2. Rinse in 70% ethanol.
3. Stain in a saturated solution of Sudan black (in 70% ethanol) for 15 min. Filter before
use.
4. Remove excess stain in 70% ethanol.
5. Stain nuclei with the carm-alum solution for 5–30 min.
6. Wash well in water and
7. Mount in an aqueous mountant (glycerine jelly).
• Fat—blue-black
• Nuclei—pale red
Medical Laboratory Technology: Volume 3
Pigments and Minerals
The pigments seen in the sections are broadly classied as artefact, endogenous, and
exogenous. Artefacts originate from xation materials such as formalin, mercury and
chromate. Endogenous pigments are those produced within the tissue while exogenous
pigments are those which are gaining access to the tissues—mainly minerals. Fixation
pigments (artefacts) are not clinically important but they should be recognized in order to
avoid improper diagnosis. Many histochemical pathologists have paid considerable aention
to endogenous tissue pigments. The chemistry of these substances seems less important
and less known. Nevertheless, the presence of haemosiderin and melanin in tissues are of
diagnostic signicance. The deposition of haemosiderin in excessive amounts inside the
tissue may indicate red blood cell related disorders,
or a haemorrhagic condition or iron overload. Melanin is a black pigment normally present
in certain tissues but some malignant tumours produce this pigment and
their diagnosis is oen based on the histochemical identication of melanin in tissues.
Haemosiderin contains active ferric iron which combines with potassium ferrocyanide in
acid solution to form ferric ferrocyanide or Prussian blue. Only a few exogenous minerals are
important for the histochemical diagnosis. These include iron and calcium. Iron is recognized
by the Prussian blue reaction (same as for haemosiderin) while calcium is recognized by the
silver impregnation method.
Haemosiderin and iron stain
is the product of the breakdown of haemoglobin and therefore occurs at the
site of previous haemorrhage. It has a golden brown granular pigment. The request for iron
staining is to check for the presence of haemosiderin in the section. Haemosiderin, however,
should be dierentiated from melanin, bile, formalin pigment and malaria pigment which
do not contain iron. The iron-containing pigment haemosiderin reacts with potassium
ferrocyanide in acid medium and yields a Prussian blue colour. This is called Perl reaction
and is used to demonstrate ferric iron (and ferritin). It is one of the classic histochemical
methods that were introduced by Perl in 1867.
Prussian blue stain
The principle of this method is that the reactive ferric iron, exemplied by haemosiderin,
produces an intense and insoluble precipitate of Prussian blue that indicates the presence of
iron. In this method a common artefact is the presence of blue granules, either on or around
the section, following treatment with the hydrochloric-ferrocyanide mixture. This could be
due to old ferrocyanide mixture or iron-contaminated water (rust). The simplest remedy is to
have a suitably sized beaker containing distilled water to one side of the bath. The distilled
Laboratory Techniques in Histology
water is maintained at the correct temperature and it is simple maer to oat out sections to
be stained by the Perl’s reaction.
• Aqueous potassium ferrocyanide (2%, w/v, aqueous; mix 2 g ferrocyanide and 100 mL
water)
Caution It is a
• Hydrochloric acid (2%, v/v, aqueous; 2 mL concentrated hydrochloric acid and 98 mL
water).
• Neutral red (1%, w/v, aqueous)
1. Take the test sections and a control section to distilled water (give bath through
decreasing concentrations of alcohol).
2. Mix equal parts of the hydrochloric acid and potassium ferrocyanide solutions and
lter.
3. Place the reagent on the sections placed on slide.
4. Leave for 30 min at room temperature, changing to a fresh solution aer 15 min.
5. Aer 15 min wash for several minutes in water.
6. Counterstain with neutral red solution for 5 min.
7. Wash in water.
8. Dehydrate (pass through increasing concentration of alcohol). The alcohol will
dierentiate.
9. Clear and mount in a DPX-type mountant.
1185
• Iron pigment (haemosiderin)—blue
• Nuclei—red
• Background—pale red
von Kossa silver nitrate procedure for calcium
Insoluble inorganic calcium salts are normally found in bone and teeth. Calcium circulates
in the blood in the free ionic form, which is not demonstrable histochemically. Abnormally,
calcium is formed in tissue in hyperparathyroidism, necrosis (e.g., tuberculous caseation) and
in association with some tumours such as myeloma.
for the demonstration of the salts in tissue
sections is the oldest and widely used. Here the calcium is substituted by silver giving a black
metallic silver colour by light photographic developer. Although not specic (melanin also
tends to blacken), it remains the method of choice. Use buered neutral formalin for xation.
The method described here has omied the post-silver hypo treatment as partial bleaching of
the blackened calcium salts may occur.
• Silver nitrate (2%, w/v, aqueous): Store the solution in a brown bole.
• Counterstain: van Gieson’s stain or 1% aqueous neutral red.
All glassware should be rinsed with distilled water prior to use.
1. Sections to distilled water, two or three changes.
2. Transfer sections to a clear glass container (Coplin staining jar) containing the silver
nitrate, or place on a slide rack and cover the section with the solution.
3. Expose the section to bright sunlight or high intensity light (a desk lamp with 60-wa
bulb source) for 60 min. If exposed to sunlight the time may be less; check microscopically.