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.