stains, and acid stains are protoplasmic. Neutral stains are
usually diffuse stains; when formed by the combination of acid and
basic dyes they usually act as selective stains for some especial
tissue-element or cell-constituent. =Metachromasia=, in its
narrowest sense, is the term applied to that staining-phenomenon,
in which a single-chemical entity gives different colors to
different tissue-elements. In this sense iodine, in its action
upon glycogen and amyloid, is a true metachromatic substance.
The majority of the so-called metachromatic stains, however,
do not possess true metachromasia, since their metachromatic
powers are dependent upon a mixture of two or more dye-stuffs in
the one compound. The most important stains of this class are
gentian violet, methyl violet, crystal violet, dahlia, thionin,
toluidin-blue, polychrome methylene blue, methylene azure, and
kresyl-echt-violett. The chief _chromotropic_ substances are
amyloid, mucin, mast-cell granules and cartilage. Metachromatic
reactions are at their best usually when examined in water; they
are affected by alcohol and usually destroyed by carbol-xylol.
Sections stained by metachromatic stains should be quickly
dehydrated by absolute alcohol and blotting, and cleared in xylol,
when mounted in balsam.
I. NUCLEAR STAINS.
1. =Haematoxylin= (C_{16}H_{14}O_{6}) is an ether extract of the
wood of _Hæmatoxylon campechianum_, a tree found in the West Indies
and Central America. In itself not a dye, it becomes one of the
most valuable when oxidized to hæmatein (C_{16}H_{12}O_{6}), and
combined with alum, iron or other mordants to form a lake. It then
stains nuclei a deep violet-blue or black color that is practically
permanent. Mucin, lime-salts, bacteria, and colonies of actinomyces
are also stained varying shades of blue. If the staining process
is prolonged the entire tissue, as well as celloidin, becomes more
or less heavily stained blue. A pure nuclear stain is obtained by
interrupting the stain at the right time (examine in water); or if
the sections are over-stained they may be differentiated in acid
alcohol (1 per cent hydrochloric acid in 70 per cent alcohol).
Hæmatoxylin stains well after all fixing-solutions except osmic
acid; some of its staining-formulæ stain slowly after fixation in
Zenker’s fluid. On the whole, it is by far the best general nuclear
stain for laboratory and diagnostic work. It is employed in numerous
staining formulæ, the most useful of which are given here. These
formulæ differ chiefly in the time of staining, “ripening” of the
stain (oxidation of hæmatein), intensity of stain, necessity of
differentiation, etc.
a. =Böhmer’s Alum-haematoxylin.=
Dissolve 5 grms. of hæmatoxylin crystals in 50 cc. of absolute
alcohol; add this drop by drop, while stirring, to 1,000 cc. of a 1
per cent solution of potassium alum. Expose in open vessel to air
and light for 1-2 weeks. Filter before using.
b. =Hansen’s Haematoxylin.=
To 200 cc. of alum-hæmatoxylin solution brought to the
boiling-point add 2 cc. of a concentrated solution of potassium
permanganate. Cool quickly; filter when cold. Can be used at once
without further ripening. It tends to stain diffusely.
c. =Delafield’s Haematoxylin.=
To 400 cc. of a saturated solution of ammonia alum add a solution
of 4 grms. of hæmatoxylin in 25 cc. of absolute alcohol. Expose
mixture to air and light for 3-4 days; filter; then add 100 cc. of
glycerin and 100 cc. of 95 per cent alcohol, and filter. Expose to
light until solution is dark enough, then keep in tightly-stoppered
bottle. It is a strong stain, and may be diluted with distilled
water when desired. The solution keeps well.
d. =Ehrlich’s Acid-haematoxylin.=
Dissolve 2 grms. of hæmatoxylin in 100 cc. of absolute alcohol.
Add this to a saturated solution of potassium alum in water 100
cc., glycerin 100 cc., and glacial acetic acid 10 cc. Allow mixture
to stand for a week exposed to air and light; then filter. Keep
in well-stoppered brown bottles. The solution stains best when it
is six months old, and may be kept for several years. It does not
overstain, and on the whole is more useful than Böhmer’s, Hansen’s
or Delafield’s.
e. =Mayer’s Haemalum.=
Dissolve 1 grm. of hæmatein in 50 cc. of 90 per cent alcohol and
warm. Add this solution to a solution of 50 grms. of potassium alum
in 1,000 cc. of distilled water dissolved by heating. Mix warm,
cool, and filter. With hæmatein no ripening is required, and the
solution can be used at once. Hæmalum is a more precise nuclear
stain, but stains more slowly than the formulæ given above.
It may be prepared directly from hæmatoxylin crystals by dissolving
1 grm. of hæmatoxylin in boiling water; add water up to 1 litre,
and cool. Add 0.2 grm. sodium iodate and 50 grms. potassium alum,
dissolving at room temperature; 50 grms. chloral hydrate and 1 grm.
citric acid may be added to make solution keep better.
f. =Mayer’s Acid-haemalum.=
Add 2 cc. glacial acetic acid to 100 cc. of hæmalum solution. It
stains the nuclei more precisely than hæmalum.
g. =Weigert’s Iron-haematoxylin.=
Dissolve 1 grm. of hæmatoxylin in 100 cc. of 96 per cent alcohol.
Allow to ripen several days, but this solution should not be kept
longer than six months. Make a second solution of 4 cc. of liq.
ferri. sesquichlor. (German Pharmak. IV, sp. gr. 1,124), 1 cc. of
concentrated hydrochloric acid and 100 cc. of water. Mix equal
parts of each solution just before staining. The mixture stains
well for 5-8 days, so that a quantity of stain sufficient for this
time only should be made up. The two stock solutions are easily
made and keep well. The nuclei stain quickly and deeply, and
differentiation and long washing are unnecessary when hydrochloric
acid is included in the second solution, as given above.
After-staining with eosin, picric acid or the Van Gieson’s mixture
gives better results with Wiegert’s iron-hæmatoxylin than with any
other hæmatoxylin.
Method of Staining with Haematoxylin.
1. Stain 1-15 minutes, controlling progress of stain by examination
of section in water, on slide, using low-power.
2. If sections are over-stained, differentiate in ½-1 per cent
potassium-alum or in acid alcohol.
3. Wash thoroughly in tap water, until a good blue is obtained.
Exposure to ammonia vapor or washing for a few seconds in
lithium-carbonate solution will hasten the development of the blue
color. If these reagents are used the section should afterwards be
thoroughly washed in water.
(Stain with a plasma stain, if contrast is desired.)
4. Dehydrate in 80 and 95 per cent alcohols.
5. Clear in carbol-xylol.
6. Mount in balsam.
Over-ripened hæmatoxylins may stain reddish or even brownish,
and too diffusely. In such cases the celloidin will be deeply
stained. The addition of alum-water to the stain may counteract the
fault. Alum-hæmatoxylins must always be filtered before using, as
precipitates are constantly formed as the result of oxidation.
2. =Carmine.= Carmine is the coloring matter of cochineal, the
dried bodies of the female _coccus cacti_, and is obtained
chiefly from Honduras. The coloring principle is carminic acid
(C_{2}H_{22}O_{12}). When combined with alum, borax, lithium,
etc., carmine gives a good, permanent nuclear stain, varying from
reddish violet to deep scarlet. It is used chiefly in pathology to
give a contrasting nuclear stain to the various pigments, and when
specific blue stains have been used for fibrin, mucin, bacteria,
elastic tissue, etc., or when a blue injection-mass has been used.
Alum-carmine is the most precise nuclear stain. Differentiation
with acid-alcohol is necessary after staining with borax- or
lithium-carmine. Lithium-carmine is on the whole the best of the
three for use as a contrast-color to the various pigments.
a. =Alum Carmine.=
Carmine ½-1 grm., 1-5 per cent alum solution 100 cc.; boil 20
minutes; cool; filter. Add crystal of thymol as preservative.
1. Stain 15 minutes to several hours.
2. Wash thoroughly in distilled water.
3. Dehydrate in 80 and 95 per cent alcohols; clear in carbol-xylol:
mount in balsam. Nuclei are a light reddish violet; the plasma is
slightly stained (muscle) or not at all.
b. =Lithium Carmine.=
2-5 grms. of carmine to 100 cc. of a cold saturated water solution
of lithium carbonate; filter.
1. Stain 1-3 minutes; transfer directly to acid alcohol (1 cc.
of hydrochloric acid to 100 cc. of 70 per cent alcohol) without
putting section into water; differentiate ¼-6 hours, until nuclei
alone retain the color. Control differentiation by examination on
the slide in acid alcohol.
2. Wash thoroughly in water.
(Use plasma stain, as picric acid, if desired.)
3. Deyhdrate in 80 and 95 per cent alcohols; clear in carbol-xylol;
mount in balsam.
c. =Borax Carmine.=
Dissolve by boiling 0.5-0.75 grm. carmine and 1-2 grms. of borax in
100 cc. of water. To the hot solution add 5 cc. of a O.5 per cent
acetic acid until solution is deep red. After 24 hours filter and
add crystal of thymol.
Stain and differentiate as with lithium-carmine, but leave the
sections somewhat longer in the stain (15 minutes).
3. =Basic Aniline Stains.= The basic aniline stains are used as
general stains for bacteria in sections, and at the same time stain
the nuclei. Methylene blue and fuchsin are employed especially for
this purpose. The metachromatic dyes (thionin, kresyl-echt-violett,
etc.) are also used as nuclear stains in combination with their
metachromatic reactions with mucin, amyloid, mast-cells, etc.
Methylene blue is used in the study of the blood-forming organs,
cell-inclusions, parasites in the tissues, etc. Safranin and fuchsin
are used for staining mitotic figures. (See Staining of Mitoses.)
Bismarck brown is employed sometimes in preparing sections for
microphotography. Methyl green is an intense chromatin stain and
is used in various combinations. Since these dyes are not very
permanent, and are easily washed out in dehydrating and clearing
fluids, as well as “running” in balsam mounts, they are rarely
employed as pure nuclear stains in pathologic work.
When the basic-aniline stains are used a saturated water or
concentrated alcoholic solution (1½-2 per cent in 40 per cent
alcohol) may be employed as stock-solution and diluted 1:5 or
1:10, as desired. The sections are stained 3-5 minutes, then
differentiated in absolute alcohol, cleared in xylol, and mounted
in balsam. Methylene-blue is used by some workers as a nuclear
stain contrasted with eosin for tissues fixed in Zenker’s
solution, giving better effects with this fixation than the
hæmatoxylins. =Unna’s alkaline methylene-blue formula= is employed
(methylene-blue 1 grm., carbonate of potassium 1 grm., water 100
cc.). Dilute 1:10 or 1:5 for staining. Stain 10-15 minutes, wash
quickly in water, differentiate in 95 per cent alcohol; dehydrate
in absolute, clear in xylol, mount in balsam. When celloidin
sections are used, 95 per cent alcohol may be used for dehydrating,
blotting on the slide with several changes of xylol.
The =nucleolus= has an affinity for the acid stains. With the
modifications of the Romanowsky methylene-blue-eosin methods the
nucleolus stains red, the nucleus blue.
II. DIFFUSE OR PLASMA STAINS.
The most commonly used diffuse stains are eosin, erythrosin, acid
fuchsin, orange G, and picric acid. They are practically never
used alone, but are employed as contrast-stains to the nuclear
stains. Eosin, orange G, acid fuchsin and picric acid may be used
as counterstains for hæmatoxylin; picric acid is used as the best
contrast to the carmines, while eosin and orange G are employed as
counterstains for methylene blue. The combination of hæmatoxylin and
eosin is by far the best general staining method for laboratory and
diagnostic work, except for tissues fixed in Zenker’s solution; for
these the combination of methylene blue and eosin is preferable.
Ammonia carmine is sometimes used as a diffuse stain for bone and
the central nervous system. The majority of the diffuse stains wash
out easily in water, alcohol and xylol, hence sections thus stained
should not be allowed to remain too long in these fluids.
a. =Eosin.= Two forms of eosin are obtainable, one soluble in
water, the other in alcohol. Saturated solutions of both kinds
should be kept as stock solutions and diluted as occasion demands.
For use after hæmatoxylin a ½ per cent solution is advisable; with
Zenker’s fixation a more dilute solution may be used, as tissues
so fixed stain intensely in eosin. If used as a contrast-stain to
methylene-blue, eosin is used first in a 5 or 10 per cent solution,
as the basic nuclear stain takes out some of it. Some workers
express a preference for the aqueous solution of eosin, others for
the alcoholic; the alcoholic solution stains more uniformly and
with less differentiation than the other. Eosin is particularly
good as a contrast-stain for tissues containing red blood cells
when fixed in formol, mercuric chloride or Zenker’s.
b. =Orange G.= Used in a 1 per cent water solution. Requires longer
time for staining than eosin.
c. =Acid Fuchsin.= A saturated water solution is kept in stock and
diluted as needed. Must be used in weaker solutions than eosin, as
it more quickly overstains, and cannot be washed out so well.
d. =Picric Acid.= Keep in stock either a saturated water or
saturated alcoholic solution, and dilute as needed. As it washes
out more readily than eosin, the staining solution should
be stronger than for the latter, and the sections somewhat
over-stained to allow for some loss of stain. Picric acid gives a
brownish tint to nuclei stained with hæmatoxylin or carmine, and
will take out the stains completely, if allowed to act too long.
e. =Ammonia Carmine.= One grm. of carmine is dissolved, without
heating, in 50 cc. of distilled water and 5 cc. of strong ammonia
water. The fluid is then exposed in an open dish until the odor of
ammonia is lost; it is then filtered. When ready for use dilute by
filtering 1-2 drops into 20 cc. of distilled water.
III. COMBINED NUCLEAR AND DIFFUSE STAINS.
The diffuse or plasma stains may be combined with the nuclear in one
staining solution, or used in succession. The latter method gives
better results. The nuclei are stained first, the diffuse stain being
used after the washing-out following the use of the nuclear stain,
except in the methylene-blue and eosin method in which to obtain the
best results it is necessary to stain with the eosin first. Nuclear
hæmatoxylin may also be followed by a combination of plasma stains,
as in the Van Gieson’s mixture of picric acid and acid fuchsin,
Delépine’s mixture of rubin and orange, White’s erythrosin and picric
acid mixture, etc. In these mixtures the different affinities of the
plasma-stains give rise to differential or selective staining effects.
a. =Haematoxylin and Eosin.=
1. Stain in any one of the hæmatoxylins.
2. Wash thoroughly.
3. Stain in dilute water or alcoholic eosin until section is
bright rose-red.
4. Differentiate eosin-staining, as desired, by rapid or slow
washing in water.
5. Dehydrate quickly in 80 and 95 per cent alcohols.
6. Clear quickly in carbol-xylol. (If carbol-xylol takes
out eosin too rapidly add some of the dry eosin stain to
it. Hæmatoxylin-stained sections can be placed in such
eosin-carbol-xylol and will take up the eosin beautifully.)
7. Mount in balsam.
Hæmatoxylin and eosin can also be combined in one stain, but the
results are not as good as those obtained by successive staining.
b. =Haematoxylin and Picric Acid.=
1. Stain with Weigert’s iron-hæmatoxylin, or overstain with any
other hæmatoxylin.
2. Wash thoroughly.
3. Stain in saturated water solution of picric acid, diluted
one-half, until sections are a bright yellow. If left too long
in the stain, the hæmatoxylin will become brown or may be wholly
lost.
4. Wash, dehydrate and clear quickly, as for eosin. (Dry picric
acid may be added to the carbol-xylol.)
5. Mount in balsam.
c. =Haematoxylin and Acid Fuchsin.=
Stain with hæmatoxylin, and after washing use a 1 per cent water
solution of acid fuchsin, until section is sufficiently red; wash;
dehydrate; clear; mount.
d. =Haematoxylin and Orange G.=
Stain with hæmatoxylin, and after washing use a 1 per cent water
solution of orange G, staining ¼-3 hours. Treat otherwise as for
eosin-staining.
e. =Carmine and Picric Acid.=
Stain with borax- or lithium-carmine; differentiate in acid alcohol
and wash thoroughly. Then counterstain with picric acid, as for
hæmatoxylin and picric acid. Carmine and picric acid may also be
combined in one stain, as picro-carmine, but this is rarely used at
the present.
f. =Eosin and Methylene-blue.=
(For tissues fixed in mercuric chloride or Zenker’s.)
1. Stain in a 5-10 per cent aqueous eosin for 20 minutes or
longer, until a deep eosin-stain is obtained.
2. Wash out excess of eosin in water.
3. Stain in Unna’s alkaline methylene-blue, diluted 1-4 or 5 with
water, 10-15 minutes.
4. Wash in water.
5. Differentiate in 95 per cent alcohol, keeping the section
in constant motion to obtain a uniform decolorization. Control
process under microscope. _Wolbach_ advises the use of a
0.75-1.5 per cent solution of colophonium in methyl alcohol as
a differentiating medium instead of 95 per cent alcohol. For
tissues fixed in formol or alcohol a 10 per cent solution should
be used.
6. When the background is pink, dehydrate quickly with absolute
alcohol, or in 95 per cent by blotting on slide with xylol, until
clear.
7. Clear in xylol.
8. Mount in balsam.
g. =Van Gieson’s Method.=
1. Stain in Weigert’s iron-hæmatoxylin, or overstain if any other
hæmatoxylin is used. (Weigert’s gives the best results, as it
does not decolorize so readily.)
2. Wash thoroughly.
3. Stain in Van Gieson’s mixture (acid fuchsin 1.5 grms.,
saturated water solution of picric acid [0.6 per cent] 150 cc.
This mixture keeps well. Add 1 cc. of this stock solution to 10
cc. of saturated water solution of picric acid. Stain in this for
10 seconds).
4. Wash quickly; dehydrate in alcohol; clear in xylol or
carbol-xylol; mount in balsam.
I have obtained the best results by making the Van Gieson’s mixture
by taking an ordinary small staining-dish nearly full of saturated
water solution of picric acid, and adding to this, drop by drop,
sufficient saturated water solution of acid fuchsin to make the
solution just dark enough so that the finger cannot be seen through
the staining-dish. The hæmatoxylin-stained section is put into this
mixture for a few seconds, until it appears to become lighter. The
section is then washed in 95 per cent alcohol, dehydrated, cleared
and mounted.
The Van Gieson method is extremely valuable in pathologic work,
because of its varied differential reactions. The nuclei are brown
or black, protoplasm is ochre-yellow, connective-tissue light
red, voluntary and involuntary muscle yellow, axis-cylinders red,
connective-tissue hyalin deep rose-red, epithelial hyalin yellow,
orange or brownish, amyloid yellow or brownish pink, mucin yellow
or brownish, fibrin yellow or brown, necrotic areas yellow or
brownish, lime-salts brown to brownish blue or violet.
Combinations of rubin and orange (Delépine) and erythrosin and
picric acid (Powell White) are also advised as differential
combination stains, but are not so useful as Van Gieson’s. Other
combination methods are to be found in the various modifications of
the Ehrlich triple stain.
CHAPTER XXVI.
SPECIAL STAINING METHODS FOR DEMONSTRATION OF PATHOLOGIC CONDITIONS
IN CELLS OR TISSUES.
=I. AMYLOID.= The best selective staining of amyloid is obtained
with relatively fresh tissues; long preservation in alcohol or
formol tends to weaken the reactions. Probably the best effects are
obtained by formol-fixation for 24 hours, and sectioning on the
freezing-microtome. Good results may be produced, however, after
any of the ordinary fixing and hardening methods by cutting the
sections on the freezing-microtome, without imbedding, or imbedding
in paraffin. The metachromatic reactions are not satisfactory with
celloidin sections. With hæmatoxylin and eosin the amyloid substance
stains a light red or bluish-pink; Van Gieson’s stains it a yellow
or brownish-pink color, giving it practically the same color that it
does epithelial hyalin. The different tissue-relations of the two
substances serve to distinguish them. The most important specific
amyloid stains are:—
1. =Iodine.=
1. Stain in Lugol’s solution 5-10 minutes.
2. Dehydrate in absolute alcohol 4 parts, tincture of iodine 1
part.
3. Clear and mount in origanum oil. Seal preparation with
paraffin, gold size or shellac.
The iodine reaction is also applied to fresh tissues by pouring
Lugol’s solution over a freshly cut surface, and is a good gross
test for amyloid. In both microscopic and macroscopic preparations
iodine gives a mahogany brown color to amyloid; other tissue is
yellow. The iodine reaction may be intensified by placing the
sections in a 1 per cent sulphuric acid; the brown color may be
changed to blue, violet or green.
2. =Methyl Violet.=
1. Stain in 0.5 per cent methyl-violet solution ½ to several
minutes. Examine in water.
2. Wash in water.
3. Differentiate in 2 per cent acetic or dilute hydrochloric acid
1-2 minutes.
4. Wash thoroughly in water.
5. Mount in lævulose or glycerin-gelatin. Amyloid ruby red;
tissue blue-violet.
3. =Gentian Violet.=
Use same method as for methyl violet. The same color-effects are
produced.
4. =Methyl Green.=
Use methyl green in the same way as methyl violet. Amyloid sky-blue
or violet; tissue is green.
5. =Iodine Green.=
1. Stain for 24 hours in a ⅓ per cent water solution of iodine
green.
2. Wash in water.
3. Mount in lævulose, glycerin or glycerin-gelatin. Amyloid red
violet; tissue green.
6. =Birch-Hirschfeld’s Method.=
1. Stain in a 2 per cent alcoholic solution of Bismarck brown for
5 minutes.
2. Wash in absolute alcohol.
3. Wash in water.
4. Stain in a 2 per cent water solution of methyl-violet (or a 20
per cent gentian violet) for 5 minutes.
5. Differentiate in 1 per cent acetic acid until the non-amyloid
parts are brown.
6. Wash thoroughly in water.
7. Mount in lævulose, glycerin or glycerin-gelatin. Nuclei are
brown; amyloid ruby red.
7. =Green’s Method.=
To a few cc. of hæmalum in a watch-glass add a saturated solution
of methyl-violet, drop by drop, until the mixture shows a faint
purple-red tinge at the edge of the glass.
1. Stain sections 15-30 minutes.
2. Differentiate in acid alcohol until the purple begins to fade.
3. Wash thoroughly in water.
4. Mount in glycerin. (Sections may be blotted and dehydrated in
pure liquid paraffin; the latter is then removed by blotting with
xylol, and then mount in pure white vaseline.)
Nuclei are blue, amyloid ruby-red.
8. =Kresyl-echt-violett (Morse’s Method).=
Kresyl-echt-violett (R. extra) 1 grm., 5 per cent carbolic acid
80 cc., alcohol 20 cc. Mix, stirring well; filter. Solution keeps
well, and can be diluted as desired without precipitating.
1. Stain 1-5 minutes.
2. Wash thoroughly in distilled water, differentiating, if necessary.
3. Blot with filter paper.
4. Dehydrate in absolute alcohol as quickly as possible.
5. Clear in turpentine. Blot nearly dry before mounting.
6. Mount in balsam.
Formol, mercuric chloride and Zenker’s all give good results.
Paraffin imbedding, with staining of sections on the cover-glass
(albumin-fixative method), is the best method of staining for
permanent mounts, although good preparations can be obtained by
the use of the freezing-microtome. Carbol-xylol cannot be used for
clearing. Nuclei are blue, protoplasm pale blue, amyloid ruby-red.
As a specific reaction for amyloid and mucin this method has been
used in my laboratory for the last ten years in preference to any
other. The stains are permanent if not exposed to the action of
light.
=Thionin=, =toluidin-blue=, =polychrome-methylene-blue=, and other
metachromatic dyes are also used to give similar reactions with
amyloid, but are not as satisfactory as the kresyl-echt-violett
method. Amyloid may also be stained with =scarlet R= or =sudan
III=, according to the method of Herxheimer, but the results are
rarely satisfactory.
=II. ATROPHY.= Good pictures of atrophic tissues are obtained
with formol-Müller’s, mercuric chloride or Zenker’s fixation, and
staining with Van Gieson’s, to bring out the stroma which is usually
relatively or absolutely increased. In the case of pigment-atrophy
the sections should be very thin and stained with alum- or
lithium-carmine.
=III. CALCIFICATION.= Deposits of lime-salts appear in fresh tissue
as gritty, refractive areas that are bright and shining by reflected
light, and dark by transmitted. They are soluble in acids, solution
of the carbonate being accompanied by the formation of bubbles of
carbonic acid gas. Hæmalum and the alum-hæmatoxylins show a specific
reaction with the phosphates and carbonates of lime, giving them a
deep blue or reddish-violet stain. Fresh calcification usually stains
diffusely blue; older deposits are deep-blue about the borders of the
deposits, lighter or unstained in the center of the mass. Tissues
containing much calcium must be decalcified before imbedding. If
the process of decalcification is not carried too far the specific
staining reaction is not lost.
v. Kossa’s Silver Method for Calcium Phosphate.
1. Fix in alcohol or formol; imbed; cut.
2. Place section in 1-5 per cent silver nitrate solution, and
expose to daylight 5 minutes to 1 hour.
3. Wash in distilled water.
4. Transfer section to a 5 per cent solution of sodium
hyposulphite, to remove excess of silver nitrate.
5. Wash thoroughly in water.
6. Dehydrate in absolute alcohol.
7. Clear in xylol; mount in balsam.
Calcareous deposit black, as the result of the formation of
phosphate of silver and its reduction by the action of light. Alum
carmine may be used as a nuclear stain before the sections are
treated with silver nitrate, or safranin may be used after the
sodium sulphite has been washed out.
=IV. CELL GRANULES AND CELL INCLUSIONS.= The granules and
cell-inclusions here included fall within the class of special
protoplasmic structures found particularly in neoplasms and inflamed
tissues, and which have been supposed to be parasites. For the
staining of other cell-granules see Blood and Blood-forming organs.
1. =Altmann’s Granules.=
1. Fix small, thin pieces of fresh tissue in equal parts of 5
per cent potassium bichromate and 2 per cent perosmic acid for
24 hours. Wash in running water for several hours. After-harden
in alcohol, and imbed in paraffin. Cut very thin and mount on
cover-glass; remove paraffin.
2. Stain in aniline-water-acid-fuchsin (acid fuchsin 20 grms.,
aniline water 100 cc.), warming until vapor is given off.
3. When cool remove the fuchsin with a mixture of 1 part
saturated alcoholic picric acid and 2 parts of water.
4. Renew the picric acid solution and warm on the paraffin oven
for 30-60 seconds.
5. Dehydrate in alcohol; clear in xylol; mount in balsam.
Protoplasm yellow: Altmann’s granules red: fat black.
2. =Russell’s Bodies.=
1. Fix and harden in Müller’s; wash; after-harden in alcohol;
imbed in paraffin; mount on cover-glass.
2. Stain sections in a saturated solution of fuchsin in 2 per
cent carbolic acid 10 minutes or longer.
3. Wash in water.
4. Wash in absolute alcohol for 30 seconds.
5. Counterstain in iodine green (1 grm. in 100 cc. of a 2 per
cent carbolic acid) for 5 minutes.
6. Dehydrate quickly in absolute alcohol.
7. Clear in xylol; mount in balsam.
Nuclei are green; Russell’s fuchsin-bodies light-red; Altmann’s
granules light-red.
3. =Pianese’s Method.=
1. Fix in Pianese’s solution (see methods of fixation) 6 hours;
wash in running water for 12 hours; after-harden in graded
alcohols; imbed in paraffin; mount on cover-glass.
2. Stain 30 minutes in a staining mixture consisting of malachite
green 0.5 grm., acid fuchsin 0.1 grm., Martius yellow 0.01 grm.,
distilled water 150 cc., 96 per cent alcohol 50 cc.
3. Dehydrate in absolute alcohol; clear in xylol; balsam.
Nuclei are green; protoplasm reddish; cell-inclusions light-red.
4. =Method for Staining “Plimmer’s Bodies.”=
1. Fix in Hermann’s fluid for 12-24 hours. Imbed in paraffin.
Mount sections on cover or slide.
2. Transfer sections to hydrogen peroxide for 15-30 seconds.
3. Wash in water.
4. Transfer to a 4 per cent ferric alum solution for 2 hours.
5. Wash in water.
6. Stain in 0.5 per cent watery hæmatoxylin solution for 30
minutes. Differentiate in the ferric alum solution until the
nuclei are dark and protoplasm colorless; control under the
microscope.
7. Wash in water 3-6 hours.
8. Counterstain in 1 per cent solution of Ehrlich’s neutral red
until section is yellow-red.
Nuclei blue-black; cell-inclusions yellow- to copper-red.
=V. CHOLESTERIN.= Cholesterin is soluble in absolute alcohol, xylol,
ether and glacial acetic. It occurs in the tissues in characteristic
rhombic plates often showing a square notch in one corner. In
sections from which the cholesterin has been dissolved its presence
may be told by the appearance of “cholesterin-clefts” in the tissue,
or often in the protoplasm of large foreign-body giant-cells
(“cholesterin-giant-cells”). With concentrated sulphuric acid,
sections or material containing cholesterin become yellow and then
rose-pink. Lugol’s gives it a brown color which turns blue-violet
after the addition of sulphuric acid, and exhibits a play of colors,
blue, green, to red.
=VI. CLOUDY SWELLING.= This is best seen in the fresh state in cells
obtained by scraping or teasing, or by the examination of frozen
sections. Osmic acid, sudan III, scarlet R. ether, alcohol and acetic
acid may be used to differentiate from fatty degeneration. The
ordinary fixing and staining methods give good pictures, except for
slight degrees of the change. These are sometimes wholly lost as the
result of the contraction due to the fixation.
=VII. COLLOID.= (See Epithelial Hyalin.)
=VIII. CORNIFICATION.= Horn takes the plasma stains (eosin,
picric acid, etc.). Van Gieson’s makes a good differential
stain. With Gram’s method horn stains deep blue, and with the
Ehrlich-Biondi-Heidenhain method it stains red. After fixation
in Flemming’s it may be stained with safranin or gentian-violet.
=Keratohyalin= occurs as fine granules in the cells of the stratum
granulosum. They stain by hæmatoxylin, carmine and Gram’s method, or
may be demonstrated by means of special stains. =Eleidin= stains with
carmine and the fat-stains, but not with hæmatoxylin.
1. =Buzzi’s Method of Staining Eleidin and Keratohyalin.=
1. Harden, imbed, cut.
2. Stain in Congo red (2-3 drops of a 1 per cent water solution
added to small basin of water) for 2-3 minutes.
3. Wash thoroughly in water.
4. Stain in hæmatoxylin, and wash.
5. Dehydrate in absolute alcohol; xylol; balsam.
Keratohyalin blue, eleidin red.
2. =Fick’s Method of Staining Keratohyalin and Keratin.=
1. Harden in alcohol, imbed, cut.
2. Stain in saturated water solution of kresyl-echt-violett for
3-4 minutes.
3. Wash thoroughly in water.
4. Differentiate in 95 per cent alcohol until connective-tissue
is colorless.
5. Dehydrate in absolute alcohol; xylol; balsam.
Keratohyalin red, keratin dark violet; nuclei blue-violet, plasma
light blue-violet.
=IX. FAT.= When alcohol has been used in the preparation of the
tissue, the fat-contents of the latter are dissolved out, and their
presence can alone be told by the presence of vacuoles. When osmic
acid is used as a fixing agent the oleates and oleic acid are
blackened. The tissue should then be washed in running water and
cut upon the freezing-microtome, or it may be imbedded in celloidin
or paraffin if this is done as quickly as possible to prevent the
loss of the fat. Chloroform or benzene should be used in place of
xylol, as the last-named dissolves out the fat. Safranin should
be used as a stain after fixation with any fluid containing osmic
acid. Frozen sections are to be mounted in glycerin-gelatin; when
balsam is used it should be warm melted Canada balsam without xylol.
Formol fixation preserves fat, and tissues so fixed may be cut on
the freezing-microtome and the sections stained with osmic acid,
sudan III or scharlach R, with nuclear counterstaining when desired.
For the demonstration of fat-embolism, fatty degeneration or fatty
infiltration the following methods are advised:—
1. =Staining of Fat with Osmic Acid.=
1. Fix in formol for 24 hours.
2. Wash; freeze; cut.
3. Place sections in 1 per cent osmic acid, Flemming’s or Marchi’s
fluid for 1-24 hours.
4. Wash in water, changing frequently.
5. 80 per cent alcohol ½-2 hours.
6. Wash in water.
7. Place section flat on slide; blot; add a drop of warmed
glycerin-gelatin; cover quickly. Ringing or sealing is not
necessary.
Or, to mount section in balsam:—
After 6, counterstain with hæmatoxylin or safranin; wash again;
dehydrate quickly with absolute alcohol; clear in pure benzene;
mount in pure melted Canada balsam (containing no xylol).
2. =Staining of Fat with Sudan III or Scharlach R.=
Staining-solutions of these dyes may be made, as follows:—
_a._ Dissolve stain in 70-80 per cent boiling alcohol, keep in
the incubator over night, and use warm.
_b._ Make a solution of absolute alcohol 70 cc., 10 per cent
caustic soda solution 20 cc., water 100 cc. Saturate this with
the stain, slightly heating.
_c._ Make a mixture of 70 per cent alcohol 50 cc. and pure
acetone 50 cc.; saturate this with the stain.
All solutions of these dyes should be filtered before using,
and should be kept covered to avoid evaporation and subsequent
precipitation.
1. Formalin fixation 24 hours; cut on freezing-microtome.
2. Place sections in 70 per cent alcohol.
3. Stain in the simple solution 20-30 minutes; in the acetone or
alkaline alcoholic solutions 2-3 minutes.
4. Wash in 50-70 per cent alcohol, differentiating as needed.
5. Transfer to water; thence to slide; blot, and mount in
glycerin gelatin.
When a nuclear counterstain is desired, put the sections in water
after 4; then stain in hæmatoxylin; differentiate quickly in acid
alcohol; wash in water; place in weak ammonia or lithium-carbonate
solution; wash in water; transfer to slide; blot; mount in glycerin
gelatin.
Sudan III and scarlet R stain the smallest particles of fat
yellowish-red to deep scarlet; scarlet R on the whole gives the
best results. The contrast with the blue nuclei when stained with
hæmatoxylin gives beautiful preparations.
3. =Staining of Fat with Indophenol.=
Stain sections with lithium-carmine; wash; then stain 20 minutes
in a saturated solution of indophenol in 70 per cent alcohol. Fat
blue; nuclei red.
4. =Staining of Fatty Acids and Soaps.=
_a. Benda’s Method._ Fix in 10 per cent formol. Transfer tissue
to Weigert’s copper-fluorchrom mordant (neutral copper acetate 5
grms., fluorchrom 2.5 grms., water 100 cc.; boil and add 5 cc. of
36 per cent acetic acid) in the incubator for 2-4 days. Cut on the
freezing-microtome. Stain sections in sudan III or scharlach R,
and then in hæmatoxylin. Nuclei are blue, normal fat red, necrosed
fat green due to formation of fatty acid copper salt. Soaps give
the same reaction when converted into insoluble salts by fixing in
formol saturated with calcium salicylate. Through comparison of
tissue hardened in this way with another portion fixed in formol
alone soaps and fatty acids may be differentiated.
_b. Smith’s Method._ Stain in concentrated water solution of Nile
blue sulphate for 10 minutes. Fat stains red, nuclei dark blue,
protoplasm light blue, fatty acids dark-blue. Differentiate in 1
per cent acetic acid; wash in water; mount in glycerin-gelatin.
=X. FIBRIN.= Fibrin stains with the acid aniline dyes, except
in areas of necrosis containing diffused chromatin, under which
conditions it stains deep blue with hæmatoxylin. In Van Gieson’s
mixture it stains yellow or brownish; in Mallory’s reticulum stain
it stains red, and with Mallory’s chloride of iron hæmatoxylin
it is grayish to dark blue. The best selective method by far is
Weigert’s, and it is the only really practical method giving a good
differentiation.
1. =Weigert’s Fibrin Stain.=
I have obtained the best results by making this stain as follows:
10 cc. of aniline oil and 100 cc. of water are shaken together
violently for several minutes, and then filtered through a moist
filter. The filtrate must contain no drops of aniline. Add to the
filtrate sufficient dry gentian-violet or methyl-violet to produce
a metallic shimmer on the surface of the solution after the dye is
dissolved by shaking. The solution will keep for several months.
Weigert advised the use of two stock solutions, I (absolute
alcohol 33 cc., aniline oil 9 cc., methyl violet in excess) and II
(saturated water solution of methyl violet). These solutions will
keep for years. When ready to use stain take 3 cc. of Sol. I and 27
cc. of Sol. II. This staining mixture will keep for about 2 weeks.
1. Fix in alcohol, formol, acetone, mercuric chloride or
Müller’s. Imbed in celloidin or paraffin; the latter preferably.
Mount sections on cover-glass with albumin fixative. Celloidin
sections must be fastened to slide by thin film of celloidin
to prevent shrinkage. Sections fixed in chromic mixtures (and
sometimes after formol fixation) must be oxidized in potassium
permanganate and then reduced in oxalic acid to give good
results. (Transfer sections to a 1 per cent solution of potassium
permanganate to which 2 volumes of water have been added; oxidize
for 10 minutes; then wash in water, and reduce for several hours
in a 5 per cent water oxalic acid solution.)
2. Wash in water.
3. Stain in lithium carmine; differentiate in acid alcohol; wash
thoroughly in water.
4. Stain on the slide or cover-glass in the aniline-methyl-violet
(or gentian-violet) solution for 10 minutes. Wash off stain with
physiologic salt solution.
5. Blot section with absorbent paper.
6. Cover section with Lugol’s (300-2-1) or a 5 per cent watery
potassium iodide saturated with iodine. Leave on section for 1-5
minutes.
7. Blot off iodine.
8. Differentiate in aniline xylol (equal parts of xylol and
aniline oil) until the nuclei become red.
9. Wash in xylol, blotting with absorbent paper. Repeat until
section is transparent; then mount in balsam. All aniline oil
must be removed before using the balsam.
Nuclei are red; fibrin deep blue; bacteria, mucin, keratin and
Altmann’s granules also blue. The differentiation must be carefully
controlled under the microscope, and should be stopped before the
finest threads of fibrin begin to be decolorized.
=XI. GLYCOGEN.= Glycogen is soluble in water; and fixation and
hardening must be carried out with absolute alcohol to prevent
the solution of the glycogen. Tissue must be fixed immediately
after death, as glycogen is quickly broken up. Its reaction with
iodine is similar to that of amyloid, but it does not give the
iodine-sulphuric-acid reaction that the latter substance does.
1. =Best’s Iodine Method.=
1. Fix and harden in absolute alcohol; imbed in paraffin; cut.
2. Stain somewhat deeply with hæmatoxylin.
3. Wash in water.
4. Stain in iodine 1, potassium iodide 2, water 100.
5. Dehydrate in iodine 2, absolute alcohol 100.
6. Differentiate in origanum oil, 1-2 hours.
7. Wash thoroughly with xylol.
8. Arrange on slide and dry in air.
9. Mount in pure melted balsam (no xylol).
Nuclei are blue, glycogen brown.
2. =Best’s Carmine Method for Glycogen.=
1. Fix in absolute alcohol; imbed in celloidin; cut.
2. Stain in hæmatoxylin; differentiate in acid alcohol.
3. Wash in water.
4. Stain in filtered carmine mixture (carmine 1 grm., ammonium
chlorate 2 grms., lithium carbonate O.5 grm., water 50 cc.; bring
to boiling point, and when cool, add 20 cc. of strong liquid
ammonia. Keep in dark; can be used after 2-3 days and gives good
results up to 14 days) 2 parts, strong ammonia 3 parts, methyl
alcohol 6 parts. Make fresh each time it is used, as it soon
precipitates; do not filter; stain few sections at a time ¾-1
hour.
5. Differentiate 1-2 minutes in a mixture of absolute alcohol 4
parts, methyl alcohol 2 parts, water 5 parts.
6. Wash in 80 per cent alcohol.
7. Dehydrate in absolute alcohol.
8. Clear in xylol; mount in balsam.
Glycogen is stained red; nuclei blue; dense connective-tissue,
mast-cell granules, protoplasm of gastric glands, etc., red; but
these can all be distinguished morphologically from glycogen. This
is by far the best method for the staining of glycogen.
=XII. HYALIN.= Epithelial hyalin (colloid) stains red or violet
with hæmatoxylin and eosin; it takes the other acid dyes and stains
to some degree with basic aniline stains. Van Gieson’s stains it a
yellow, orange or brownish-pink. Kresyl-echt-violett gives it a deep
indigo-blue color or a more green robin-egg blue. _Connective-tissue
hyalin_ stains deep brilliant red with Van Gieson’s; this is the best
method for differentiating connective-tissue hyalin from amyloid or
epithelial hyalin. Russell’s method also stains hyalin red.
=XIII. HYDROPIC DEGENERATION.= Fix by heat or formol-alcohol. Imbed
in celloidin; stain with hæmatoxylin and eosin.
=XIV. HYPERTROPHY.= Fix in Müller’s or mercuric chloride for simple
staining; for study of nuclei fix in Flemming’s and stain with
safranin.
=XV. INFLAMMATION.= The process of inflammation may be studied to
advantage in the mesentery, web or tongue of the curarized living
frog, by stretching these parts over a cork-ring attached to a
glass plate on which the animal rests. The exposed tissues must be
kept moist with physiologic salt solution. Heat, chemicals or other
irritants may be employed to produce the inflammatory reaction.
For the study of the inflammatory process in sections the ordinary
fixations may be employed, but for the study of the nuclei, mitotic
figures and cell-granulations fixation in Flemming’s, Zenker’s, etc.,
is advised. Safranin, methylene blue and eosin, the various stains
used in the study of blood-cells, etc., may be used.
1. =Staining of Mast-cells.=
_a. Kresyl-echt-violett_ used as for amyloid or mucin is the best
stain for mast-cells. The cell-granules stain bright rose-red.
_b. Ehrlich’s Dahlia Method._
1. Harden in absolute alcohol; imbed; cut.
2. Stain with saturated water solution of dahlia.
3. Wash in water.
4. Dehydrate in absolute alcohol.
5. Clear in xylol; mount in balsam.
_c. Unna’s Method for Mast and Plasma Cells._
1. Harden in absolute alcohol; imbed; cut.
2. Stain in Unna’s polychrome methylene blue ¼-12 hours.
3. Wash in water.
4. Differentiate in Unna’s glycerin-ether mixture (Grübler) 15
seconds to several minutes.
5. Wash carefully in water.
6. Dehydrate in absolute alcohol; clear in xylol; mount in balsam.
Mast cell granules are red; plasma cell granules blue.
_d._ Various modifications of the _Romanowsky method_ stain mast-
and plasma-cells very well.
=XVI. IODINE.= For the demonstration of iodine in tissues the
following method has been advised by =Justus=. The experience of
other workers with it has not been satisfactory.
1. Harden in absolute alcohol; imbed in celloidin; cut.
2. Soak in water to remove alcohol.
3. Put section in wide-mouthed, stoppered bottle in freshly
prepared green chlorine-water for 1-2 minutes.
4. Transfer section on a glass needle to a vessel containing 500
cc. water and 1 cc. of a 1 per cent solution of silver nitrate for
2-3 hours. The section is colored yellow-green, and a precipitate
of silver chloride appears.
5. Transfer section to a warm saturated solution of sodium chloride
until it becomes light.
6. Wash in distilled water.
7. Transfer to a concentrated solution of mercuric chloride.
8. Examine in pure glycerin.
Iodine should be red.
=XVII. MITOTIC FIGURES.= Various histologic methods devised for the
study of mitoses can be applied to the demonstration of these in
neoplasms, inflammation and regeneration. Flemming’s solution or
mercuric chloride fixation gives best results, although formol, or
even absolute alcohol, when used quickly and carefully gives fair
results if tissue is very fresh.
1. =Flemming’s Solution and Safranin.=
1. Fix small pieces of fresh tissue in Flemming’s, in the dark,
for 24 hours; wash 24 hours; after-harden in graded alcohols;
imbed and cut.
2. Stain in 1 per cent water solution or saturated aniline water
solution of safranin, or 1 per cent water methyl violet for 12-24
hours, or carbol-fuchsin for one hour.
3. Differentiate quickly in a 0.5-0.0001 HCl in 70 per cent
alcohol and then in absolute alcohol until stain no longer comes
away in clouds and nuclei have right shade.
4. Clear in xylol; mount in balsam.
Fat is black; mitoses stand out sharply; tubercle-bacilli may be
stained black or red.
2. Fixation in mercuric-chloride may be followed by
=Ehrlich-Biondi-Heidenhain’s stain= (saturated aqueous orange 100
cc., saturated aqueous acid fuchsin 20 cc., saturated aqueous
methyl green 50 cc.) 12 grms. of Grübler’s prepared stain dissolved
in 100 cc. of distilled water, for stock solution. For staining
take 1 cc. of stock solution, water 30 cc., ½ per cent watery acid
fuchsin 3 cc., and 2 per cent acetic 5-6 drops. Stain 2-24 hours;
wash in 90 per cent alcohol; dehydrate in absolute; clear in xylol;
balsam.
Resting nuclei are bluish; mitoses and fragments of leukocyte
nuclei dark green; red blood cells orange red; protoplasm and
connective-tissue fuchsin red.
3. =Benda’s Iron-Haematoxylin Method.=
1. Fix in osmic acid, mercuric chloride or other fixative.
2. Stain sections by placing them in liq. ferri. sulfur. oxyd.
(Germ. Pharm.) diluted with double its volume of water, for 24
hours; wash carefully in distilled water and then in tap water;
stain in 1 per cent watery hæmatoxylin until section is black.
Wash in water. Differentiate in 10-30 per cent acetic acid, or in
liq. ferri. sulfur. oxyd. diluted with distilled water 1-20. A 10
per cent solution of ferric sulphate may be used instead of the
persulphate.
4. =Heidenhain’s Iron-Haematoxylin.=
1. Imbed in paraffin after fixation in mercuric chloride.
2. Immerse section in a 1.5 per cent solution of iron-alum
sulphate (violet-colored salt) or iron-ammonium sulphate for ½-3
hours.
3. Wash in water.
4. Stain in 0.5 per cent watery hæmatoxylin or hæmatein for 12-18
hours.
5. Wash in water.
6. Differentiate in the iron-alum or iron-ammonium solution until
the section becomes deep blue (control under microscope) and
nuclear structures stand out distinctly.
7. Wash in running water for 15 minutes.
8. Absolute alcohol; xylol; balsam.
Instead of the watery hæmatoxylin solution a mixture of hæmatoxylin
1 grm., alcohol 10 cc., and water 90 cc. may be used. Keep four
weeks before using. Stain 24-36 hours. For contrast staining a
weak solution of Bordeaux red may be used before the iron-alum and
hæmatoxylin, staining 24 hours.
=XVIII. MUCIN.= Mucin stains a deep blue or reddish-violet with
an over-ripe hæmatoxylin. When counterstained with picric acid
very beautiful preparations can be obtained. Mucin also gives
a metachromatic reaction with kresyl-echt-violett, thionin,
toluidin-blue and polychrome methylene-blue, staining red with these
stains. Water or carbolic-acid solutions of these stains may be used;
dehydrate in absolute alcohol, clear in xylol, and mount in balsam.
In my opinion =Morse’s Carbol-kresyl-echt-violett= method as given
above for amyloid is the best of these metachromatic reactions.
Muchæmatein and mucicarmin give the most delicate reactions.
1. =Mayer’s Muchaematein.=
1. Absolute alcohol fixation is preferable.
2. Stain sections in Mayer’s solution (hæmatein O.2 grm. mixed
with a few drops of glycerin, O.1 grm. of aluminum chloride, 40
cc. of glycerin, 60 cc. of water) for 5-10 minutes.
3. Wash in water.
4. Dehydrate in absolute alcohol; xylol; balsam.
Carmine may be used for counterstaining; mucin is blue. Should the
mucin swell in the stain replace water and glycerin with 100 cc. of
70 per cent alcohol and 1-2 drops of nitric acid.
2. =Mayer’s Mucicarmin.=
Make staining solution by mixing 1 grm. carmine, O.5 grm. aluminum
chloride, 2 cc. water and 100 cc. of 50 per cent alcohol, heating
over the flame for 2-3 minutes until mixture darkens. Let stand 24
hours and filter. The stock solution may be diluted 1-10. Stain
10 minutes. If it does not stain well add 0.5-1 grm. of aluminum
chloride. Mucin alone should be stained red. Counterstain with
hæmatoxylin.
=XIX. MYELIN.= This appears in the form of doubly refractive
granules, that stain with less intensity with the fat dyes, but may
be differentiated from fat in that it loses the power of reducing
osmic acid after being mordanted for eight days or more in bichromate
solutions, while fat does not.
=XX. NECROSIS.= Hæmatoxylin and eosin, and Van Gieson’s give good
pictures. Use Weigert’s fibrin stain for coagulation-necrosis, and
Benda’s method for the demonstration of fatty acids for the staining
of fat-necrosis. Recent necrotic areas stain diffusely blue with
hæmatoxylin; older areas may take the plasma stains alone. Use
various methods for the demonstration of micro-organisms in the
necrotic areas.
=XXI. NEOPLASMS.= Use hæmatoxylin and eosin, and Van Gieson’s for
ordinary diagnosis. To differentiate sarcoma and carcinoma use Van
Gieson’s, Mallory’s or other reticulum stains. For the study of
cell-inclusions use Altmann’s, Russell’s, Plimmer’s and Pianese’s
methods. Special fixation or Zenker’s is necessary. Methylene-blue
and eosin after Zenker’s give excellent pictures. For the
demonstration of mitoses the methods given above should be employed.
=XXII. PIGMENT.= Use the carmines for contrasting melanin,
hæmofuscin, lipochromes, hæmatoidin, hæmosiderin, bilirubin and all
yellow, brown, blue, black, etc., extrinsic pigments. In tissue
fixed in mercuric chloride or formol bilirubin is green, and can
thus be differentiated from hæmatoidin. The lipochromes give weak
fat-reactions, and this is used to distinguish them from other
yellow or brown pigments. Alcohol fixation is the best for pigment
study, although the other fixing solutions may be used. Formol
sometimes produces pseudo-pigments by its action upon hæmoglobin.
The iron-reactions are obtained best in sections cut on the
freezing-microtome, although both paraffin and celloidin imbedding
may be used. In testing for iron glass needles should be used and all
traces of iron should be removed from staining-dishes, slides, etc.,
by treating with hydrochloric acid, distilled water and alcohol.
1. =Potassium Ferrocyanid Test for Iron.=
1. Stain sections in lithium carmine for several hours.
2. Differentiate in acid alcohol, stopping short of the desired
complete differentiation of the nuclei.
3. Wash in water.
4. Saturated solution of potassium ferrocyanid 1-3 hours.
5. Acid alcohol until iron-pigment becomes blue (½-12 hours).
Complete differentiation of nuclei.
6. Wash in water.
7. Dehydrate in absolute alcohol.
8. Clear in xylol; mount in balsam.
Hæmosiderin is blue (Berlin blue); nuclei are red. Lithium carmine
may be used after the iron-test, if desired.
2. =Ammonium Sulphide Test for Iron.=
1. Fix in alcohol; imbed; cut.
2. Treat sections with yellow ammonium sulphide for 5-60 minutes.
3. Wash quickly in water.
4. Dehydrate in absolute alcohol.
5. Clear in xylol; mount in balsam.
Stain with lithium carmine either before or after the reaction with
ammonium sulphide. Iron is grayish-black to black.
3. =Combined or Masked Iron.=
1. Treat tissues with Bunge’s fluid (95 per cent alcohol 95 cc., 25
per cent hydrochloric acid 10 cc.) for 1-2 hours at 50-60°C., until
inorganic iron is all removed.
2. Place tissues in acid alcohol (sulphuric acid 4 cc. in 100 cc.
alcohol 95 per cent.)
5. Wash sections in acid alcohol, then pure alcohol, and finally in
distilled water.
6. Transfer to ammonium sulphide (5-60 minutes) or to potassium
ferrocyanid and O.5 HCl for 5 minutes.
7. Wash in water.
8. Counterstain in eosin or safranin; wash; dehydrate in absolute
alcohol; clear in cedar-oil; mount in benzene balsam. Keep
preparations in the dark.
4. =Staining of Chromophilic Cells.=
1. Fix in a chromic solution. In this the chromophilic cells become
yellow or brown.
2. Stain in polychrome methylene blue; the cells become grass-green
in color.
5. =Tests for Silver, Lead and Mercury.=
Use ammonium sulphide as for iron. Black sulphides are formed.
6. =Test for Copper.=
Treat with potassium ferrocyanid and hydrochloric acid; copper
gives a dark yellow-brown coloration.
=XXIII. PSEUDOMUCIN.= It is not precipitated by acetic acid. It has a
greater affinity for the diffuse stains than mucin, and gives weaker
metachromatic reactions.
=XXIV. REGENERATION AND REPAIR.= For the staining of mitoses, cell
granules and cell-inclusions see methods given above. See also
methods for staining of epithelium, reticulum, neuroglia, etc.
XXV. URIC ACID AND PURIN BASES:—
1. =Courmont and Andre’s Method.=
1. Fix in absolute alcohol; imbed; cut.
2. Treat sections with 1/100 ammonia solution or very weak sodium
hyposulphite solution.
3. Transfer to 1/100 silver nitrate solution.
4. Wash.
5. Develop with a photographic developer.
6. Wash in water; stain with hæmalum and eosin; dehydrate; clear
in xylol; balsam.
Uric acid and xanthin or purin bases appear as black granules.
CHAPTER XXVII.
THE STAINING OF PATHOGENIC MICRO-ORGANISMS IN TISSUES
Rapid fixation and hardening are requisites for the successful
staining of micro-organisms in sections. Alcohol, Zenker’s, mercuric
chloride and formol give best results; Müller’s because of its
slow action is not good, although formol-Müller’s may be used
because of the more rapid fixation with this fluid. In the case
of formalin-fixation staining with Weigert-Gram’s method may not
give good results unless the sections are oxidized in potassium
permanganate solution and then reduced in oxalic acid. (See Staining
of Fibrin.) Preservation of the tissue for a long time in alcohol
impairs the staining power of micro-organisms contained within it.
The tissue should be imbedded preferably in paraffin, as very thin
sections must be obtained. The freezing-microtome may be employed
and the thinnest sections selected for staining. Celloidin stains
very heavily with the aniline dyes and retains the color, so that
bacteria in celloidin sections do not stand out very distinctly.
On the whole paraffin sections, floated on slide or cover, and
fastened by albumin-fixative, give the best results, though for
the micro-organisms stained in carbol-fuchsin and decolorized in
nitric acid it is best to float the sections directly onto the
warm stain without removing the paraffin, and mount without the
use of alcohol. This method may be employed for all stains that
are taken out by alcohol. The stains used for film preparations
are as a rule applicable to sections. The basic aniline dyes,
particularly methylene-blue, fuchsin, methyl or gentian violet,
kresyl-echt-violett, thionin, and Bismarck brown, either in saturated
alcoholic solutions or dilutions of such, or in combination with
alkalies, aniline oil or phenol, are usually employed. The various
modifications of the Romanowsky method are very useful. The time
required for staining in sections is usually much longer than for
films; but the staining can often be accelerated or strengthened by
warming over the flame or in the incubator. Contrast staining of the
nuclei with lithium-carmine or Bismarck brown is advisable after the
use of staining methods in which the nuclei are decolorized. Xylol or
origanum oil should be used for clearing.
I. THE STAINING OF BACTERIA IN TISSUES.
According to their staining-reactions bacteria may be very
conveniently grouped in three classes: 1, _Staining with
Gram-Weigert’s method_; 2, _Not staining with Gram-Weigert’s_; 3,
_Staining with the tubercle-bacillus method_ (_acid-resisting_).
1. BACTERIA STAINING BY THE GRAM-WEIGERT METHOD.
Weigert’s modification of Gram’s method, as given above for the
staining of fibrin, is the best for the staining of bacteria
that stain by this method. (See Fibrin, Chapter XXVI.) The
differentiation with aniline-xylol is slower and safer than
with alcohol. Acetone-xylol (1:5) has been recommended in
place of aniline-xylol. Wolbach recommends the use of a 5-10
per cent colophonium-alcohol for differentiation. Contrast
staining with watery Bismarck brown, dilute carbol-fuchsin or
eosin may be carried out if desired. The aniline-xylol may
be saturated with eosin and the section stained during the
differentiation. _Carbol-gentian-violet_ may be used instead of
aniline-gentian-violet; it keeps much better than the latter.
Staining by Gram’s Method (Gram-positive).
Staphylococcus pyogenes aureus.
Staphylococcus pyogenes albus.
Staphylococcus pyogenes citreus.
Streptococcus pyogenes.
Micrococcus tetragenus.
Diplococcus pneumoniæ.
Bacillus aërogenes capsulatus.
Bacillus of diphtheria.
Bacillus of anthrax.
Bacillus of leprosy.
Bacillus of tetanus.
Bacillus of tuberculosis.
Bacillus of rhinoscleroma.
Bacillus of mouse septicæmia.
Bacillus of swine erysipelas.
Oïdium albicans.
Mycelium of actinomyces.
2. BACTERIA NOT STAINING BY GRAM’S METHOD.
For the bacteria belonging to this class Löffler’s methylene-blue,
carbol methylene-blue, a watery solution of methylene-blue or
gentian-violet, Leishman’s or Wright’s modification of Romanowsky’s
methylene-blue eosin method (see page 290), Unna’s alkaline
methylene-blue solution preceded by eosin after Zenker’s fixation
(see page 260), aniline gentian-violet, Zieler’s method and
carbol fuchsin are most commonly used as stains. Wolbach advises
the use of a 5-10 per cent acetone-colophonium solution for the
differentiation of Gram-negative bacteria in tissue fixed in formol.
1. =Löffler’s Methylene-blue.=
1. Saturated alcoholic solution of methylene-blue 30 cc.;
potassium hydrate solution (1 in 10,000) 100 cc.
2. Stain 5 minutes to 24 hours.
3. Wash in water.
4. Differentiate in 1 per cent acetic acid, 10-30 seconds.
5. Wash in 90 per cent alcohol, 2-5 minutes; dehydrate in
absolute alcohol; clear in xylol; mount in balsam.
2. =Gentian-violet.=
1. Stain sections in a 2 per cent watery gentian-violet for 5-20
minutes.
2. Wash in water.
3. Decolorize in 70 per cent alcohol until stain ceases to come
away.
4. Dehydrate in absolute alcohol; clear in xylol; balsam.
3. =Zieler’s Method.=
1. Fix in Orth’s solution, or any fixing solution except those
containing osmic acid; imbed in paraffin or celloidin.
2. Stain in Pranter’s solution (orcein D 0.1 grm., hydrochloric
acid 2.0 cc., 70 per cent alcohol 100 cc.) for 8-24 hours.
3. Wash rapidly in 70 per cent alcohol.
4. Wash in water.
5. Stain in polychrome methylene-blue 10 minutes to several hours.
6. Wash in distilled water.
7. Differentiate in glycerin ether until no more clouds of color
come away and section is light blue.
8. Wash in distilled water.
9. 70 per cent alcohol for a few seconds; absolute 5-10 minutes;
xylol; balsam.
Protoplasm is gray-brown; bacteria dark-blue; background colorless.
Zieler’s method is especially good for the staining of the
glanders, typhoid and chancroid bacilli and the gonococcus.
For Unna’s methylene-blue eosin and the modifications of the
Romanowsky method see Pages 260 and 290 respectively. Pappenheim’s
methyl-green-pyronin method is also recommended for the staining in
sections of Gram-negative bacteria.
Not Staining by Gram’s (Gram-negative).
Gonococcus.
Micrococcus melitensis.
Meningococcus (in sections).
Bacillus of bubonic plague.
Bacillus of chancroid.
Bacillus coli communis.
Bacillus dysenteriæ.
Bacillus of epidemic conjunctivitis (Koch-Weeks).
Bacillus of influenza.
Bacillus mallei.
Bacillus pneumoniæ.
Bacillus proteus.
Bacillus of malignant œdema.
Bacillus pyocyaneus.
Bacillus of typhoid fever.
Bacillus of fowl cholera.
Bacillus of rabbit septicæmia.
Bacillus of swine plague.
Spirillum of Asiatic cholera.
Spirochæte pallida.
3. BACTERIA STAINING BY THE TUBERCLE-BACILLUS METHOD.
(ZIEHL-NEELSEN.)
1. Tubercle-bacillus.
2. Lepra-bacillus.
3. Smegma-bacillus.
4. Lustgarten’s bacillus.
1. Stain sections by floating thin paraffin sections directly on
to warm carbol-fuchsin (fuchsin 1 grm., absolute alcohol 10 cc.,
cryst. carbolic acid, 5 grms., water 100 cc.) for 1-3 minutes.
2. Transfer on spatula to water, agitating so as to wash off excess
of stain.
3. Transfer section to 30 per cent nitric acid and water
alternately, until section has a pale lilac tint.
4. Wash in water.
5. Float on warm watery methylene-blue for 1 minute.
6. Wash in water.
7. Float section on slide; dry over flame or in oven; melt over
flame, and put section at once into xylol to remove paraffin.
8. Balsam.
Various staining-methods have been recommended for the staining of
the most important pathogenic bacteria in tissues. The most useful of
these methods are here given:—
_a. Cocci._
1. =Pyogenic Cocci.= Stain by Gram-Weigert’s, contrast with
Bismarck brown or lithium-carmine.
2. =Pneumococcus.= Stains with ordinary water solutions,
carbol-fuchsin and Gram-Weigert’s. The staining of the capsule in
sections is not very satisfactory.
3. =Gonococcus.= Gram-negative. Stains in sections with Zieler’s
method, Löffler’s methylene-blue, or dilute carbol-fuchsin with
differentiation in alcohol.
4. =Micrococcus Catarrhalis.= Stains like the gonococcus.
5. =Diplococcus Intracellularis Meningitidis.= Smear preparations
often Gram-positive, in sections usually Gram-negative. Use same
stains as for gonococcus.
6. =Micrococcus Tetragenus.= Stain with Gram’s or watery solutions
of basic aniline dyes.
_b. Bacilli._
1. =Anthrax-bacillus.= Stain with Gram-Weigert’s and contrast with
Bismarck brown or lithium-carmine. Stains also with strong watery
gentian-violet solution, with differentiation in strong alcohol.
2. =Bacillus of Malignant Oedema.= Gram-negative. Stain with watery
solution or gentian-violet.
3. =Bacillus of Tetanus.= Gram-positive. Stains with watery
solutions of basic aniline dyes.
4. =Bacillus Aërogenes Capsulatus.= Gram-positive. Stains with
other aniline stains.
5. =Bacillus Pyocyaneus.= Stains with Gram’s and other aniline dyes.
6. =Bacillus of Influenza.= Gram-negative. Fix tissue in alcohol.
Stain with dilute carbol-fuchsin and differentiate in dilute acetic
acid.
7. =Koch-Week’s Bacillus.= Gram-negative.
8. =Bacillus of Bubonic Plague.= Gram-negative. Stain by
_Gaffky’s method_ (Fix in alcohol or a mixture of glacial acetic
acid 10.0, chloroform 30.0, and 96 per cent alcohol 60.0, imbed
in paraffin, stain 2-3 hours in weak watery methylene-blue,
dehydrate quickly in absolute alcohol, xylol, balsam). It may
also be stained by 24 hours in concentrated solution of fuchsin
in glycerin, rapid differentiation in weak acetic; alcohol;
xylol; balsam. Alcohol or mercuric chloride fixation should be
used, as formol fixation does not give good staining.
9. =Typhoid Bacillus.= Löffler’s methylene-blue or carbol-fuchsin,
staining 24 hours, decolorizing in dilute acetic and washing
rapidly in alcohol. Zieler’s method may also be used. It is
Gram-negative.
10. =Paratyphoid Bacillus.= Stains like the typhoid bacillus.
11. =Colon Bacillus.= Gram-negative. May be stained with Löffler’s
methylene-blue or carbol-fuchsin.
12. =Diphtheria Bacillus.= May be stained in sections of
diphtheritic membranes with Löffler’s methylene-blue, watery
aniline stains, or with Gram’s if the decolorization is not carried
too far.
13. =Bacillus of Chancroid.= Gram-negative. Stain according to
_Unna’s method_:—
1. Fix in alcohol.
2. Stain 5-10 minutes in a mixture of a solution of
methylene-blue 1 grm., potassium carbonate 1 grm., alcohol 20
cc., water 100 cc., and a solution of methylene-blue 1 grm.,
borax 1 grm., water 100 cc.
3. Place sections on slide; blot.
4. Decolorize in Unna’s glycerin-ether mixture.
5. Dry; dehydrate in alcohol.
6. Xylol; balsam.
14. =Bacillus of Glanders.= Gram-negative. Stain with Zieler’s
method or with Löffler’s methylene-blue, differentiating in weak
acetic.
_Noniewicz’s Method._
1. Stain with Löffler’s methylene-blue 2-5 minutes.
2. Wash in water.
3. Differentiate for about 5 seconds in a mixture of ½ per cent
acetic acid 75 cc., ½ per cent watery solution tropaeolin 25 cc.
4. Wash in water; dry by blotting; xylol; balsam.
Bacilli deep-blue; tissues light-blue.
15. =Bacillus of Rhinoscleroma.= Gram-positive. Fix in alcohol for
_Wolkowitsch’s method_:—
1. Stain in aniline gentian-violet 24-48 hours.
2. Wash in water.
3. Treat with Lugol’s 1-4 minutes.
4. Decolorize in absolute alcohol.
5. Remove more color by oil of cloves.
6. Xylol; balsam.
In tissues fixed in osmic acid and then stained in hæmatoxylin
the bacilli are dark blue with light blue capsules. The hyaline
substance of rhinoscleroma stains with basic stains.
16. =Friedländer’s Bacillus.= Gram-negative. Stains with ordinary
aniline dyes. For staining the capsules the following method is
advised:—
1. Stain for 24 hours in the incubator in a mixture of a
concentrated alcoholic solution of gentian-violet 50 cc., glacial
acetic acid 10 cc., and distilled water 100 cc.
2. Wash in a 1 per cent acetic acid solution.
3. Alcohol; xylol; balsam.
Bacilli deep-blue; capsules light-blue.
17. =Tubercle-bacillus.= Gram-positive. Stain in sections on
warm carbol-fuchsin without removing paraffin, as given above.
Alcohol and mercuric chloride fixation give best results.
Aniline-gentian-violet may also be used, staining with a warm
solution for 15-30 minutes, and decolorizing in 20 per cent nitric
acid followed by 70 per cent alcohol, counterstaining in Bismarck
brown, dehydrating in alcohol, clearing in xylol and mounting in
balsam. The Weigert-Gram method may be used for the demonstration
of the branched or streptothrix forms of the tubercle bacillus. For
celloidin sections _Mallory and Wright_ advise the following:—
1. Stain rather lightly in alum-hæmatoxylin.
2. Wash in water.
3. Dehydrate in 95 per cent alcohol.
4. Attach sections to slide by ether-vapor method.
5. Stain in steaming carbol-fuchsin 2-5 minutes.
6. Wash in water.
7. Acid alcohol ½-1 minute.
8. Wash thoroughly in several changes of water to remove acid
completely and to bring back blue color to nuclei.
9. 95 per cent alcohol to remove fuchsin.
10. Aniline-oil, followed by xylol, blotting.
11. Xylol; balsam.
Celloidin is colorless, nuclei blue, tissue colorless,
tubercle-bacilli red. Orange G may be used as a diffuse stain.
18. =Lepra Bacillus.= Gram-positive. Stain paraffin sections on
warm carbol-fuchsin, as for the tubercle-bacillus. To differentiate
from the tubercle-bacillus, stain 6-7 minutes in a dilute alcoholic
solution of fuchsin, and decolorize in acid alcohol (nitric acid 1,
alcohol 10). Lepra-bacilli stain; tubercle-bacilli do not.
_c. Trichomycetes._
1. =Actinomyces.= Alcohol and formol fixation are best. Good
preparations can be obtained with hæmatoxylin and eosin, Van
Gieson’s or Weigert-Gram’s. The special staining methods advised
give no better results than these simpler stains. Differential
staining of clubs and mycelium may be obtained by _Mallory’s
method_:—
1. Stain lightly in alum-cochineal. (Powdered cochineal 6 grms.,
ammonia alum 6 grms., water 100 cc. Boil half an hour, add water
lost by evaporation, filter, add crystals of thymol.)
2. Saturated watery eosin 10 minutes.
3. Wash in water.
4. Stain in aniline gentian-violet 2-5 minutes.
5. Wash in physiologic saline solution.
6. Transfer sections to Lugol’s for 1 minute.
7. Pass rapidly through water.
8. Dry thoroughly between folds of filter-paper.
9. Cover section with aniline-oil until clear.
10. Xylol; balsam.
Clubs pink; mycelium blue.
2. =Nocardia, Cladothrix, Streptothrix and Leptothrix.= Löffler’s
methylene-blue and carbol-fuchsin give good results. The Nocardiæ
are acid-fast with dilute acids. They give good preparations with
Weigert’s fibrin stain and lithium-carmine.
_d. Vibrios._
1. =Cholera Vibrios.= Gram-negative. Sections may be stained with
fuchsin or methylene-blue.
_e. Spirilla and Spirochætes._
1. =Spirillum of Recurrent Fever.= Stain in sections with
Levaditi’s silver-method, or with Nikiforoff’s method:—
1. Fix for 24 hours in equal parts of a 5 per cent water solution
of potassium bichromate and a saturated solution of mercuric
chloride in 0.6 per cent sodium chloride.
2. After-harden in graded alcohols in the incubator.
3. Imbed in paraffin.
4. Stain 24 hours in a mixture of alcoholic 1 per cent solution
of tropæolin 5 cc., concentrated watery methylene-blue solution
10 cc., caustic potash solution (1:1000) 2 drops.
5. Wash in water.
6. Dip several times in a mixture of equal parts of absolute
alcohol and ether.
7. Oil of bergamot; xylol; balsam.
The spirillum of _African relapsing fever_ stains with the same
stains as the spirillum Obermeieri. The _spirochætes of Vincent’s
angina_ and _fowl-spirillosis_, and the _spirochæte refringens_
stain with watery aniline dyes and with Giemsa’s stain; in section
they are stained by the Levaditi method.
2. =Spirochaeta Pallida (Treponema Pallidum).= This organism is
best examined in the living condition by means of the dark-field
illumination (dark-field condenser). A very simple method of
dark-field illumination consists of the use of India ink. The
suspected discharge or serum is placed on a slide and an equal
quantity of ink (Gunther’s or Higgin’s) added. The serum and ink
are rapidly mixed and spread over the slide to dry in a pale brown
smear. The oil for the immersion is placed directly on the smear.
The spirochætes appear as white spirals against a brownish-black
field. The best results are obtained with serum; the presence
of mucus or fibrin interferes with the clearness of the picture
obtained.
Smears of serum from syphilitic lesions may be dried in the air and
fixed in absolute alcohol or equal parts of absolute alcohol and
ether for 15-20 minutes. They may then be stained by Giemsa’s (old
formula) stain (azur II-eosin 3 grms., azur II 0.8 grm., glycerin
[Merck’s chemically pure] 250 grms., methyl-alcohol [Kahlbaum I]
250 grms.). This solution can be obtained from Grübler. Ten drops
of the stain are mixed with 10 cc. of distilled water immediately
before the staining. The fixed preparation is covered with the
diluted staining fluid and warmed over the flame until a slight
steam arises. It is then allowed to cool for about 15 seconds, when
the stain is poured off and replaced by fresh, and the process
repeated four or five times, when the preparation is washed,
dried and mounted in balsam. Spirochætes are dark red. Slide or
cover-glass and forceps must be absolutely clean. Smears may also
be fixed and stained by _Wright’s blood-stain_.
For the demonstration of the treponema in sections the method of
_Levaditi_ gives the most satisfactory results:—
1. Fix thin pieces of tissue 24 hours or longer in 10 per cent
formol. (Formol-Müller’s and alcohol-fixation may also be used.)
2. 24 hours in 96 per cent alcohol.
3. Transfer to distilled water until tissue sinks.
4. Impregnation for 3 days in incubator, in a 1.5-3 per cent
silver nitrate solution.
5. Wash for a short time in water.
6. Reduce for 48 hours, in the dark, at room-temperature, in
pyrogallic acid 4 grms., 40 per cent formol 5 cc., distilled
water 100 cc.
7. Wash in water. Cut on freezing-microtome, or imbed in
celloidin or paraffin. Toluidin-blue or safranin may be used as a
contrast-stain.
The spirochætes are dark brown to black. Silver precipitates occur
chiefly in the outer portions of the tissue. The reticulum is
brown; other parts of the tissue are yellowish. Levaditi’s more
recent modification of this method does not give so good results as
the original.
_Schmorl’s Staining of Sections with Giemsa’s Stain._
1. Fix in 10 per cent formol. Cut very thin sections on
freezing-microtome.
2. Place the sections in a staining dish containing a measured
amount of distilled water. To each cc. of water add one drop
of Giemsa’s stain. Use clean glass-needles to manipulate the
sections. After 1 hour transfer sections to a fresh solution, in
which they are left 5-12-24 hours.
3. Wash quickly in a concentrated solution of potassium alum,
then quickly in water.
4. Mount in glycerin-gelatin; or dry on the slide until nearly
perfectly dry, then xylol, and balsam, or cedar oil. Alcohol must
not be used.
II. THE STAINING OF PATHOGENIC YEASTS AND MOULDS IN SECTIONS.
1. =Blastomycetes.= The parasites of blastomycetic dermatitis
can be demonstrated unstained in pus treated with a weak sodium
hydroxide. In sections they are easily found after treatment
with ordinary staining methods. The various modifications of the
Romanowsky method, or other methylene-blue-eosin staining, give
better staining of the parasite than can be obtained by hæmatoxylin
and eosin.
2. =Oïdium Albicans.= Staining with Weigert-Gram’s and
lithium-carmine gives beautiful preparations.
3. =Moulds.= These are best examined in the unstained condition,
by treating the material with equal parts of alcohol and ether,
followed by a 3 per cent potassium hydroxide solution. The
organisms and spores are brought out distinctly. Löffler’s
methylene-blue may be used for staining. In the case of sections
stain 1-2 hours and contrast with eosin. For the examination of
hairs or horny scales for fungi, _Unna’s method_ may be used:—
1. Add glacial acetic acid to hair or epidermis; make cover-glass
preparations, drying by heat.
2. Ether and alcohol equal parts.
3. Stain in borax 1 grm., methylene-blue 1 grm., water 100 cc.,
½-5 minutes.
4. Wash in water; dry; balsam.
If the horny elements are too deeply stained, decolorize in 1 per
cent acetic for 10 seconds, or in 1 per cent oxalic, citric, or
arsenious acid for 1 minute.
III. THE STAINING OF ANIMAL PARASITES.
1. =Amoeba Coli.= Examine fresh material from fæces, abscesses or
cultures, in physiologic saline solution, on a warm stage. Stain
under the cover with methylene-blue and carmine. Make permanent
mounts by removing excess of stain and running in 50 per cent
glycerin. In fixed preparations the nuclei of the amoebæ do not
stain with ordinary nuclear stains. _Mallory’s method_ may be used:—
1. Fix in alcohol.
2. Stain sections in a saturated aqueous solution of thionin 3-5
minutes.
3. Differentiate in a 2 per cent aqueous solution of oxalic acid
for ½-1 minute.
4. Wash in water; dehydrate in absolute alcohol; clear in xylol;
mount in xylol-balsam.
Nuclei of the amœbæ and granules of the mast-cells are
brownish-red; nuclei of cells blue.
2. =Trichomonas vaginalis and intestinalis=; =Cercomonas coli=;
=Megastoma entericum=; =Balantidium coli=; =Pyrosoma bigeminum=;
=Trypanosoma=; =Leishman-Donovan bodies=, and allied forms are
best stained with the modifications of Romanowsky’s stain; the
chromatin is red-violet (macro-nucleus red, micro-nucleus black,
flagellum red, protoplasm blue, basophilic granules black). For
staining in sections mercuric chloride or Zenker’s fixation
followed by staining with polychrome methylene-blue, Giemsa’s or
the modifications of the Romanowsky method may be employed.
3. =Plasmodium Malariae.= For films make medium smears (not too
thin); fix with equal parts of absolute alcohol and ether for ½-1
hour; or fix and stain in the same solution (Leishman-Romanowsky,
Wright’s stain, etc.). For single staining methylene-blue,
carbol-thionin, etc., may be employed; for double staining
eosin and methylene-blue. Ehrlich’s tri-acid, or any of the
eosin-methylene-blue combinations may be used (particularly the
Leishman-Romanowsky or Wright’s). With the Romanowsky methods the
body of malarial organism is stained blue, the chromatin varying
shades of lilac, red, purplish-red or almost black. When the blood
contains but few parasites 1 cc. may be drawn, mixed with 20 cc.
of distilled water and centrifugated. Smears are then made of the
sediment. For the staining of the plasmodium in imbedded tissues
the following method is recommended by _Bignami_. The tissue should
be fixed in formol or mercuric chloride, preferably a mixture of
mercuric chloride 1 grm., sodium chloride 0.75 grm., acetic acid
0.75 grm., water 200 cc. Fix for 2 hours; after-harden in alcohol
and iodine-alcohol, changing the alcohol each day for seven days.
Dehydrate in absolute alcohol, and imbed in celloidin or paraffin.
Stain in a saturated watery solution of magenta or in a mixture
of equal parts of saturated alcoholic mixtures of magenta and
orange G. Good results may, however, be obtained with Löffler’s
methylene-blue. Clear in xylol; mount in balsam.
4. =Coccidia and Sarcosporidia.= The ordinary fixations give good
results. Imbed in paraffin or celloidin. Weigert’s iron-hæmatoxylin
and Van Gieson’s give as good pictures as any of the special
methods advised.
5. =Negri Bodies of Rabies.= Examine in smears or make sections.
Take portions of gray brain-substance from the cortex in the
region of the fissure of Rolando (in the dog from around the
crucial sulcus), from the hippocampus, and from the cerebellum.
Smear cover or slide by taking a thin slice of the gray matter
and compressing it between two slides, or cover and slide, or by
drawing the cover across the cut surface in order to get some
of the cells. Dry in the air; stain with watery methylene-blue;
wash; stain with watery acid fuchsin; wash in water; blot dry;
mount in balsam. Negri bodies fuchsin-red (about size of red
blood cells); everything else blue. When dried in the air and
then fixed in methyl alcohol for 5 minutes the smears may be
stained by Giemsa’s method. For the demonstration of the bodies
in sections fix in Zenker’s, imbed in paraffin, and stain by the
eosin-methylene-blue method. The bodies take the eosin stain.
Formol fixation, freezing-microtome and Romanowsky stain give
quick results. _Mann’s method_ for the staining of Negri bodies in
sections is strongly recommended by many workers. Fix material in
mercuric chloride or Mann’s fluid (1 grm. picric acid and 2 grms.
tannin dissolved in 100 cc. concentrated water solution of mercuric
chloride) for 24 hours; wash thoroughly in running water; imbed in
paraffin. Stain in Mann’s mixture (1 per cent aqueous methyl-blue
[not methylene-blue] 35 parts, 1 per cent aqueous eosin 35 parts)
for 24 hours; wash in water; rinse in absolute alcohol; place in
alkaline alcohol (absolute alcohol 50 cc., 4 drops of a 1 per cent
solution of sodium hydroxide) for 15-20 seconds until sections
become reddish; wash quickly in alcohol; wash about 2 minutes in
water until superfluous color is removed; place in weak acidulated
water (acetic acid) 1-2 minutes until sections are blue; quick
dehydration in alcohol; xylol; balsam. Cells are blue, nucleoli and
blood-vessels red; Negri bodies bright red. For quick diagnosis use
acetone fixation and imbedding, stain in Mann’s fluid 2-4 minutes,
and proceed as in the Mann’s method. While these bodies possess
a great diagnostic importance for rabies, their exact nature
must still be regarded as unsettled; they are most probably not
parasites.
6. =Vaccine Bodies.= Fix in Flemming’s, mercuric chloride or
Zenker’s; imbed in celloidin or paraffin. Stain with Heidenhain’s
iron-hæmatoxylin (bodies black) or Biondi-Heidenhain mixture
(bodies blue, nuclei of leukocytes and mitoses green, nuclei
of epithelium and connective-tissue blue, protoplasm and
connective-tissue red). These bodies are probably not parasites,
but may be products of cell-degeneration.
7. =Vermes.= The _heads_, _proglottides_ and _ova_ are best
examined in the fresh state, in physiologic saline or glycerin.
Acetic acid may be used to bring out details. Berlin-blue or
methylene-blue may be injected through the genital pore for the
demonstration of the excretory and genital organs. _Scolices_
and _hooklets_ of _echinococcus_ may be obtained by scraping
the cyst-wall; examine in glycerin. Permanent preparations of
_cestodes_, _nematodes_ and _trematodes_ may be made by fixing
in mercuric chloride, formol or Flemming’s, after-hardening in
alcohol, staining in orange G, borax carmine, alum hæmatoxylin,
hæmatoxylin and eosin etc., mounting in glycerin gelatin; or
dehydrating, clearing in xylol and mounting in balsam. For sections
imbed in paraffin or celloidin. _Trichinæ_ may be studied by
teasing the fresh muscle; by digesting with pepsin and hydrochloric
acid and examining the freed trichinæ on a warm stage; or by
imbedding in paraffin or celloidin and staining with hæmatoxylin
and eosin. Permanent mounts of the embryos of _filaria_ may be
made by fixing cover-glass preparations of blood or chylous fluid
by heat or mercuric chloride, and staining for a few seconds with
Löffler’s or a 2 per cent aqueous thionin.
CHAPTER XXVIII.
THE STAINING OF SPECIAL ORGANS AND TISSUES.
I. BLOOD AND BLOOD-FORMING ORGANS.
The =blood= may be examined by means of _films_, _stained_ or
_unstained_, or by _sections_, _celloidin_ or _paraffin_.
_A. FILMS._ The blood may be obtained from the pulp of the ring
finger, from the skin over the knuckles, or from the posterior aspect
of the lobe of the ear. The place selected should be carefully
cleansed with water, soap and 1/1000 mercuric chloride solution, and
finally with alcohol and ether. A puncture is made with a sterilized
triangular needle or knife, or a stub-pen with one point broken
off. The last-named makes a most useful and inexpensive instrument
for this purpose. The puncture should be made by a quick and deep
stab, so that sufficient blood can be obtained from one stab-wound.
Pressure should not be employed to force blood from the wound.
Bleeding may be encouraged by letting the arm hang down, or by
applying pressure in the furrow of the terminal joint of the finger.
The first drop of blood should be wiped away with a clean towel.
When the second drop reaches the size of a pin-head touch it with
the under side of a perfectly clean cover-glass, held by forceps,
not by the fingers; place this cover-glass immediately upon another
clean cover, so that the blood will spread out between the two covers
in a thin film. The covers are then separated by sliding them apart
without pressing or squeezing; place covers with film side upward,
and dry in the air. The films should not be touched with the hands;
forceps alone should be used to handle them. If the blood does not
dry as quickly as it is spread the film will be too thick. Films
may be made upon slides in the same way, or the drop of blood may
be caught upon the edge of a clean cover, slide or “spreader” and
then drawn rapidly across a slide. The dried film may be marked
by scratching with a needle-point the number and date on the film
itself. Blood-films may be fixed _without drying_ by exposure to the
vapor of formol or osmic acid for several seconds and then dropping
into absolute alcohol. Formol alcohol, saturated mercuric chloride
solution or Flemming’s solution may also be used for the fixation of
wet films, fixing for 5-10 minutes, and washing thoroughly after
each of the last two solutions. The _dried film_ may be fixed by
exposure to heat (110-115°C.) for 5-10 minutes for Ehrlich’s triple
stain, and for 2 hours for the methylene-blue-eosin methods; 30-60
seconds at a temperature of 120°C. may suffice; the film should be
brought at once into the required temperature. Heat-fixed films are
improved by dipping them for a few minutes in mercuric chloride
solution and then washing well before staining. _Acetone-free methyl
alcohol_ (1-2 minutes), _absolute alcohol and ether in equal parts_
(½-12 hours), _formol-alcohol_ (1-2 minutes), _alcoholic mercuric
chloride_ (absolute alcohol 25 cc., ether 25 cc., 5 drops of a 2
grms. mercuric chloride solution in 10 cc. of absolute alcohol)
for 2-5 minutes, and _formol-vapor_ are the chief solutions used
for fixing the air-dried film. For ordinary work methyl alcohol,
formol alcohol, and the absolute-alcohol and ether mixture give good
results; heat fixation brings out the granules well, and mercuric
chloride is a good fixative for the leukocytes. The combination of
fixation and staining, as in Leishman’s or Wright’s modification of
the Romanowsky method, is also recommended for general work.
For the _staining of blood-films_ an almost endless variety of
staining-methods can be found in the literature. Many of these
represent slight deviations in the method of making the stain or in
its application, such deviations marking stages of improvement in the
development of the method. It is not necessary, therefore, to give
all of these methods, but to consider only the latest modifications
of value. In a general way _blood-stains_ may be divided into five
classes:—
1. =HAEMATOXYLIN AND EOSIN.=
Fix in equal parts of absolute alcohol and ether for at least
30 minutes; stain with hæmalum and eosin, or Ehrlich’s acid
hæmatoxylin and eosin. By adding O.5 grm. of eosin to the formula
for Ehrlich’s acid hæmatoxylin a combination stain can be made that
is very good for blood-films fixed by heat or absolute alcohol and
ether. Stain 2-24 hours, wash, dry and mount in xylol balsam.
2. =EOSIN AND METHYLENE-BLUE.=
Fix by formol (dried film over 40 per cent formol for 1 minute);
absolute alcohol for 1 minute; stain 5 minutes in a 1 per cent
watery eosin; then without removing eosin place in watery
methylene-blue for 2 minutes; wash quickly; dry in air; balsam.
3. =MIXTURES OF EOSIN AND METHYLENE-BLUE.=
The numerous mixtures of methylene-blue and eosin are not very
stable, can be kept for a few days only, and give varying results.
_Jenner_ improved this method of staining greatly by collecting the
precipitate formed by the addition of eosin to methylene-blue, and
dissolving it in pure methyl alcohol, thus giving a solution that
fixes and stains at the same time. The _May-Grünwald_ method is
practically the same.
Jenner’s Method.
a. Water-soluble eosin, 1.25 grms.
Distilled water, 100 cc.
b. Medicinal methylene-blue, 1 grm.
Distilled water, 100 cc.
Mix equal parts of _a_ and _b_ in an open basin, stirring with
a glass rod. Let stand for 24 hours; filter; dry the residue at
50°C. Wash residue thoroughly with distilled water and again dry
thoroughly. Take 0.5 grm. of the dried powder and dissolve in 100
cc. of pure methyl alcohol. Filter. Solution keeps well.
1. Make blood-film. Dry in air. Do not fix.
2. Cover film with stain, keeping under watch-glass to prevent
evaporation Stain 2 minutes.
3. Wash in distilled water until the film has a pink color. Dry
in air. Mount in xylol-balsam.
Neutrophile granules are red, eosinophile rose red, basophile
granules violet, red blood cells and central portion of
blood-platelets are terra-cotta, leukocyte nuclei and granules in
red blood cells are blue, protoplasm of nuclei and outer portion of
platelets light blue.
4. =MODIFICATIONS OF THE ROMANOWSKY METHOD.=
A large group of stains has resulted from various applications
of the Romanowsky idea of uniting equimolecular proportions of
methylene-blue and eosin, and the solution of the dyes so obtained
in some suitable solvent. These dyes consist of mixtures of
methylene violet, methylene azure, eosinate of methylene blue,
etc., and can be obtained from Grübler and Co. under various names,
such as _Azur-blau_, _Bleu Borrel_, _Giemsa’s stain_, _Leishman’s
stain_, etc. _Hastings_, _Leishman_, _Wright_ and others have
combined the Romanowsky method with that of Jenner by dissolving
the new dyes obtained by their various modifications in pure methyl
alcohol, so as to form a solution that will fix and stain at the
same time. _Hastings’ stain_ is a modification of _Nocht’s stain_;
_Wright’s stain_ is a modification of the _Leishman-Romanowsky_
method. The revised directions given by _Wright_ for making and
using his stain are here given. Wright’s method and the Giemsa
stain possess all of the staining advantages afforded by the
variations of the Romanowsky method, and are alone given here. The
former is recommended for blood-work, the latter for the staining
of protozoa.
Wright’s Blood-stain.
To a 0.5 per cent aqueous solution of sodium bicarbonate add
methylene-blue (B.X or medicinal) in the proportion of 1 grm. of
the dye to each 100 cc. of the solution. Heat the mixture in a
steam sterilizer at 100°C. for one full hour, counting the time
after the sterilizer has become thoroughly heated. The mixture
should be placed in a flask of such size and shape that the
fluid will not be more than 6 cm. deep. After heating, allow the
mixture to cool, placing the flask in cold water if desired, and
then filter it to remove the precipitate. When cold the fluid
should have a deep purple-red color when viewed in a thin layer by
transmitted yellowish artificial light. It does not show this color
while warm.
To each 100 cc. of the filtered mixture add 500 cc. of a 0.1 per
cent aqueous solution of yellow water-soluble eosin, and mix
thoroughly. Collect on a filter the abundant precipitate which
immediately appears. When the precipitate is dry, dissolve it in
pure methyl alcohol (Merck’s) in the proportion of 0.1 grm. to 60
cc. of the alcohol. To facilitate solution the precipitate is to be
rubbed up in a porcelain dish or mortar with a spatula or pestle.
This alcoholic solution is the staining solution. It should be
kept in a tightly-stoppered bottle. Should it become concentrated
through evaporation methyl alcohol in proper quantity should be
added.
1. Cover film with a given quantity of staining fluid by means of
a medicine dropper.
2. After 1 minute add to the staining fluid on the film the same
quantity of distilled water by means of the medicine dropper,
and allow the mixture to remain for 2-3 minutes according to the
intensity of the stain desired. A longer period of staining may
produce a precipitate. Eosinophile granules show best after short
staining. The quantity of diluted stain on the preparation should
not be so great that some of it runs off.
3. Wash the preparation in water for 30 seconds or until the
thinner portions of the film become yellow or pink in color.
4. Dry, and mount in balsam.
Films more than a few hours old do not stain as well as fresh ones.
The _red cells_ are orange or pink in color. Polychromatophilia
and punctate basophilia or granular degeneration are well shown.
Nucleated reds have deep-blue nuclei, and their cytoplasm is
usually bluish. The _lymphocytes_ have dark purplish-blue nuclei
and cytoplasm of a robin’s-egg blue, in which a few dark-blue
or purplish granules are sometimes present. The nuclei of the
_polynuclear neutrophilic leukocytes_ are dark-blue or dark
lilac-colored, the granules reddish-lilac. The _eosinophiles_ have
blue or dark lilac nuclei, a blue cytoplasm and eosin-red granules.
The _large mononuclear leukocytes_ have a dark lilac or blue
nucleus, cytoplasm pale blue or blue with dark-lilac or deep purple
granules. Mast-cells have purplish or dark-blue nuclei, bluish
protoplasm and coarse dark purple or black granules. _Myelocytes_
have dark blue or lilac nuclei, blue cytoplasm, and dark-lilac
or reddish-lilac granules. _Blood platelets_ are blue with small
violet or purplish granules in their central portions. _Malarial
parasites_ have a blue body and lilac or red chromatin. _Spirochæte
pallida_ is pale blue.
Giemsa’s Method.
_a_, One per cent water solution of azur-blau; _b_, one per cent
watery solution of eosin. For staining take 1 cc. of _b_, add 10
cc. of water, and then 1 cc. of the azur-blau solution. Stain 10
minutes to 1 hour.
Giemsa’s Old Method.
Azur II—Eosin 3.0 grm.
Azur II 0.8 grm.
Glycerin (Merck’s pure) 250.0 cc.
Methyl-alcohol (Kahlbaum I) 250.0 cc.
To 1 cc. of distilled water in a small, perfectly clean graduate
add 1 drop of the stain, shaking very gently. Make very thin
film; dry in air; fix 15-20 minutes in absolute alcohol. Cover
preparation with a thin layer of the freshly diluted stain for
10-15 minutes, renewing stain at end of 10 minutes. Wash in a
stream of water. Differentiate over-stained preparations in
distilled water. Dry with absorbent paper; mount in balsam. Stains
the _spirochæte pallida_ and _malarial organisms_. The Giemsa
solution may be obtained from Grübler. A more intense staining can
be obtained by adding to the water used for diluting the stain 1-2
drops of a 0.1 per cent solution of potassium carbonate.
5. SPECIAL ELECTIVE STAINS FOR THE BLOOD.
1. =Ehrlich’s Triple Stain.=
Saturated watery solution of Orange G 120 cc.
Saturated watery solution of acid fuchsin 80 cc.
Saturated watery solution of methyl green 100 cc.
Glycerin 50 cc.
Distilled water 300 cc.
Absolute alcohol 180 cc.
Mix gradually; allow to stand for several months; do not shake or
filter. Remove stain with pipette. Fix by heat, or pure methyl
alcohol for 5 minutes. Stain 5-10 minutes; wash thoroughly, dry
and mount in balsam. Neutrophile granules violet; eosinophile,
a bright red; nuclei of the neutrophilic and eosinophilic cells
greenish-blue; nuclei of the lymphocytes deep-blue; nuclei of the
large mononuclears pale blue; those of red cells intense blue; red
cells copper red. The Aronson-Philipp modification is more variable
and less satisfactory.
Pappenheim’s Stain for Lymphocytes.
3-4 parts of polychrome methylene-blue or methyl green to 1-2 parts
of pyronin. Fix in absolute alcohol. Nuclei blue-green; protoplasm
bright red.
Staining of Blood-platelets.
The blood-platelets may be examined in the fresh state by coating
a cover-slip with Deetjen’s agar-solution (boil 5 grms. agar-agar
in 500 cc. distilled water, filter hot, and to each 100 cc. of the
filtrate add 0.6 grm. sodium chloride, 6-8 cc. of a 10 per cent
solution of sodium phosphate and 5 cc. of a 10 per cent solution
of sodium diphosphate). Place drop of blood on this coating and
examine on warm stage. For permanent stained preparations bleed
into a fixing and staining fluid (equal parts alcohol and ether and
Romanowsky’s stain) or use Wright’s stain.
Bremer’s Diabetic Reaction.
Take a clean cover-glass, smear one-half with normal blood, the
other half with diabetic blood. Fix for 2 hours at 120°C., or in
equal parts of absolute alcohol and ether at 60°C. for 4 minutes.
Stain in a 10 per cent watery methylene-blue for 2 minutes, wash
off the stain in water, and stain for 10 seconds in a ⅛ per cent
watery eosin. Wash, dry and mount in balsam. In diabetic blood the
red cells are green; in normal blood red. While this reaction is
constant in diabetic blood it also occurs in leukæmia, Hodgkin’s
disease, exophthalmic goitre and multiple neuritis. A 1 per cent
solution of _Biebrich scarlet_ stains diabetic blood intensely,
normal blood but slightly. On the other hand, a _1 per cent
methylene-blue_ and a _1 per cent Congo red_ stain normal blood
intensely and diabetic blood slightly.
Staining of Glycogen in Leukocytes.
To a solution of Lugol’s (100:3:1) add sufficient gum arabic to
make a syrupy mixture. Keep tightly corked. Place a drop of this
solution upon an air-dried film. After 1 minute dry with blotting
paper. Examine with oil immersion. A positive reaction is shown
by the presence of a diffuse brown or reddish-brown coloration or
granules in the cell-body of the polymorphonuclear leukocytes.
Staining of Fat in Blood.
Stain in solutions of scharlach R or sudan III in 70 per cent
alcohol.
Staining of Blood-parasites.
The _malarial parasites_, _trypanosomes_, _Leishman-Donovan
bodies_, _sporidia_, _piroplasma bigeminum_, _spirilla_ and
_spirochætes_ and the _filaria_ may all be stained with Wright’s
or Giemsa’s modification of the Romanowsky method, or by any of
the modifications of this method. (See also Staining of Animal
Parasites.)
_B. SECTIONS._ The blood is allowed to drop directly into
Flemming’s solution and allowed to stand for 24 hours. It is then
washed in water by repeated decanting, or the coagulum may be
placed in a bottle covered with muslin, and then exposed to running
water. It is after-hardened in alcohol and imbedded in paraffin.
Safranin should be used to stain the sections. This method is
especially good for the demonstration of mitoses in the blood-cells.
Bone-marrow.
Prepare films and fix and stain, as for blood films. For sections,
fix the marrow in formol-Müller’s, mercuric chloride, Zenker’s,
etc.; imbed in paraffin; cut very thin sections; stain with
Ehrlich’s triple stain or Wright’s modification of Leishman’s
stain. To distinguish the young forms of erythrocytes and
leukocytes _Trambusi_ fixes in Flemming’s, stains the sections
in a 1 per cent thionin solution in aniline-water (4:100),
differentiates in acid-alcohol, and then brings the sections into
a watery eosin and finally an alcoholic eosin, and mounts in
xylol-balsam.
Spleen and Lymphnodes.
Fresh material may be obtained by means of a trocar, and may be
examined in the fresh state, or films may be prepared, fixed
and stained, as for blood-smears. Sections of fixed tissues may
be obtained by the use of the same fixing and staining methods
employed in the study of the blood or bone-marrow. For the study
of the reticulum Mallory’s reticulum stain or the digestion-method
may be used. In ordinary work formol-fixation followed by
eosin-staining is of great value in distinguishing hæmolymphnodes
and lymphatic glands.
II. BONE.
For ordinary work decalcification is necessary except for those
pathologic conditions in which the lime-salts have been lost (see
Chapter XXI). Imbed in celloidin preferably. When decalcification
has been carried out place sections in an alkaline solution before
staining, and stain for a longer period than usual. Methylene-blue
and eosin stain osteoblasts and osteoclasts very well. Van
Gieson’s is an especially useful stain for ordinary work; osteoid
tissue is red, calcified areas yellow. Sections of bone without
decalcification can be prepared by fixing, hardening and staining
in bulk; the bone is then sawn in the dried condition, and the
sections ground down to the required thickness. _Schmorl’s_ methods
for the preparation of bone-sections have practically superseded
all other staining methods.
Schmorl’s Thionin-picric-acid Method.
1. Fix in formol or formol-Müller’s preferably.
2. Decalcify in formol-nitric acid or Ebner’s alcoholic HCl acid
solution.
3. Wash thoroughly in water. After-harden in increasing strengths
of alcohol; freeze or imbed in celloidin (not paraffin); cut.
4. Transfer sections to water for 10 minutes.
5. Stain sections, well spread out, for 5-10 minutes in a
solution of 2 cc. of a saturated solution of thionin in 50 per
cent alcohol and 10 cc. of water, to which 1-2 drops of ammonia
are added.
6. Wash in water.
7. Stain ½-1 minute in hot saturated and cold filtered
watery-solution of picric acid.
8. Wash in water.
9. Differentiate in 70 per cent alcohol until the color ceases to
come away in blue-green clouds, 5-10 minutes or more.
10. Dehydrate in 96 per cent alcohol.
11. Clear in phenol xylol; xylol; balsam.
Lacunæ and canaliculi dark brown to black; bone cells red;
ground substance yellow or brownish yellow. Calcified areas take
a darker yellow than non-calcified. This method consists in an
impregnation with a fine precipitate rather than a staining. Should
the precipitate be too heavy in portions of the section it may be
removed by thorough washing between 9 and 10.
Schmorl’s Thionin and Phosphotungstic or Phosphomolybdic Acid
Method.
1. Fix thin pieces of fresh bone in formol, then in Müller’s for
6-8 weeks, or 3-4 weeks in the incubator. Fixation is best at
37°C.
2. Decalcify in Ebner’s hydrochloric acid solution. Wash
thoroughly. After-harden in alcohol. Imbed in celloidin or
paraffin. Cut very thin sections.
3. Water for 10 minutes.
4. Stain in the alkaline-thionin solution, as in previous method,
for 3 minutes.
5. Wash in water.
6. With glass needles transfer sections to a saturated watery
solution of phosphotungstic or phosphomolybdic acid for a few
seconds.
7. Wash 5-10 minutes, until section is sky-blue in color.
8. Fix the stain for 3-5 minutes in ammonia 1 part, water 10
parts.
9. Transfer directly to 90 per cent alcohol; change twice.
10. Dehydrate; clear in carbol-xylol; balsam.
If the ground-substance is too dark differentiate in acid alcohol
before dehydrating, and then wash thoroughly before beginning the
dehydration.
Outlines of lacunæ and canaliculi are dark blue; ground-substance a
light or greenish blue; cellular elements a diffuse blue. Schmorl
advises this method for growing bone; in rickets the well-ossified
areas alone stain well. Both of the Schmorl methods can be used for
the study of =teeth= as well.
Staining of Sharpey’s Fibres (v. Kölliker).
1. Harden, decalcify, imbed, cut.
2. Place section in strong acetic acid until it becomes
transparent.
3. Stain in a saturated watery solution of indigo carmine for
15-16 seconds.
4. Wash in water; mount in glycerin.
Fibres red; ground-substance blue.
III. CARTILAGE.
Cartilage stains deeply with hæmatoxylin; with Weigert’s fibrin
stain it holds the blue; with thionin and polychrome methylene-blue
stains for mucin cartilage stains metachromatically red.
IV. CONNECTIVE TISSUES.
_a._ =Connective-tissue Fibrils.= The demonstration of
connective-tissue fibrillæ or reticulum is of great importance in
the differential diagnosis of sarcoma and carcinoma. _Van Gieson’s_
method is the best stain for the coarser fibrils, but does not
bring out the finer reticulum as well as Mallory’s aniline-blue
method.
1. =Mallory’s Reticulum Stain.=
1. Fix in mercuric chloride or Zenker’s. After-harden in alcohol;
imbed in celloidin or paraffin; cut.
2. Stain in 1/10 aqueous acid fuchsin 5-10 minutes.
3. Transfer directly to the following solution and stain for 20
minutes or longer:—
Aniline-blue, water-soluble (Grübler) 0.5 grm.
Orange G (Grübler) 2.0 grms.
1 per cent aqueous solution of phosphomolybdic acid 100 cc.
4. Wash and dehydrate in several changes of 95 per cent alcohol;
dry with absorbent paper.
5. Clear in xylol or origanum oil.
6. Mount in balsam.
Fibrils and reticulum of connective tissue, amyloid, mucin
and connective-tissue hyalin are blue; nuclei, protoplasm,
fibroglia fibres, axis cylinders, neuroglia fibres and fibrin
are red; elastic fibres are pink or yellow; red blood-cells and
myelin-sheaths are yellow.
2. =Mall’s Method for Reticulum.=
1. Digest frozen sections of fresh tissue for 24 hours in a
solution of Parke, Davis and Co.’s pancreatin 5 grms., soda
bicarbonate 10 grms., water 100 cc.
2. Wash carefully in water.
3. Place section in test-tube half-full of water and shake
thoroughly to remove cells.
4. Spread out on slide and allow to dry.
5. Allow a few drops of the following solution to dry on
slide:—picric acid 10 grms., absolute alcohol 33 cc., water 300
cc.
6. Stain for 30 minutes in the following mixture: acid fuchsin 10
grms., absolute alcohol 33 cc., water 66 cc.
7. Wash in the picric acid solution for a second.
8. Dehydrate in absolute alcohol; xylol; balsam.
3. =Unna’s Method for Collagen.=
1. Harden in absolute alcohol; imbed; cut.
2. Stain 5-15 minutes in polychrome methylene-blue.
3. Wash in water.
4. Differentiate in 1 per cent neutral orcein in absolute
alcohol, 15 minutes.
5. Dehydrate in absolute alcohol.
6. Clear in xylol; mount in balsam.
Nuclei dark blue; protoplasm light blue; collagen dark red;
plasma-cell granules greenish-blue; mast-cell granules red.
4. =Mallory’s Fibroglia Method.=
1. Fix thin, small, fresh pieces of tissue in Zenker’s fluid;
harden in alcohol; imbed in celloidin or paraffin; cut.
2. Stain sections in 1 per cent aqueous acid fuchsin for 12 hours
in the cold, or 20-30 minutes at 50-56°C.
3. Wash in water for 5 seconds.
4. Differentiate in 0.25 per cent aqueous potassium permanganate
solution 10-20 seconds.
5. Wash in water for 5 seconds.
6. Dehydrate in alcohol; clear in xylol or origanum; mount in
balsam.
Fibroglia fibrils and cell-nuclei intensely red; contractile
elements of striped muscle, smooth muscle, neuroglia fibres,
cuticular surfaces of epithelial cells and fibrin are also red;
connective-tissue fibres brownish-yellow or colorless; elastic
fibres, unless degenerated, bright yellow.
_b._ =Elastic Fibres.= Weigert’s method is so superior to the Unna
orcein-stain that it alone is given here. It is our best elective
stain: gives permanent preparations, and is in every way practical.
The stain keeps well.
Weigert’s Method for Staining Elastic Fibres.