Practical pathology

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.

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.

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.

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.

3. Unna’s Method for Collagen.

Nuclei dark blue; protoplasm light blue; collagen dark red; plasma-cell granules greenish-blue; mast-cell granules red.

4. Mallory’s Fibroglia Method.

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.

Preparation. Boil in a porcelain dish resorcin 4 grms., fuchsin (Grübler) 2 grms., and water 200 cc. After the mixture has boiled a few seconds add 25 cc. of liquor-ferri sesquichlor., Pharm. Germ. III. Stir well and boil for 5 minutes. When cool, filter. Carefully loosen the filter from the funnel, transfer it to the same porcelain dish which still contains a small amount of sediment, and add 200 cc. of 94 per cent alcohol. Boil and stir carefully. Remove the filter-paper when all the sediment is dissolved. Cool, filter; make up the filtrate to 200 cc. with 94 per cent alcohol, and to these 200 cc. add 8 cc. of hydrochloric acid. Resorcin-fuchsin may be obtained from Grübler, but the freshly-prepared stain gives better results.

Nuclei are red; elastic fibres blue-black. Should the stain when old give a diffuse staining differentiate for a longer time in acid alcohol.

c. Fat Tissue. Use same methods as advised for the demonstration of fatty degeneration and infiltration (osmic acid, scharlach R, sudan III).

V. EAR.

Remove temporal bone; fix in formol-Müller’s; decalcify in trichloracetic acid; wash; after-harden in alcohol; imbed in celloidin. For nerve-endings use Golgi’s and methylene-blue methods.

VI. EYE.

Fix in Müller’s, formol-Müller’s, Zenker’s, formol, Flemming’s or Marchi’s solution. Aid fixation by incisions into sclera. The eye should not be left in formol for more than 3 days. Section as desired; imbed larger pieces in celloidin, small ones in paraffin. Use alum-carmine, iron-hæmatoxylin, Van Gieson’s, Weigert’s elastic stain, Levaditi’s silver-method, Golgi’s nerve-stain, methylene-blue method, etc., according to the object of the investigation.

VII. LIVER.

For the demonstration of the bile-capillaries Weigert’s neuroglia method gives the best results. (See Page 300.) This method may be used with sections cut on the freezing-microtome after fixation in formol. Such sections are placed in a ½ per cent solution of chromic acid for 1 hour, transferred to the neuroglia mordant for 5-6 hours, washed well with water, and then treated as for the neuroglia method. Van Gieson’s method may also be used for frozen sections of formol-fixed tissue. The walls of the capillaries show as fuchsin-colored streaks.

VIII. MUSCLE.

Van Gieson’s is the best general stain for both striped and unstriped muscle, as it differentiates the muscle perfectly from the connective-tissue. Mallory’s reticulum stain may also be used for the same purpose. For the study of myoglia fibrils the tissue must be fixed within a few minutes after its removal from the living body. Autopsy material cannot be used. These fibrils can be demonstrated by Mallory’s fibroglia stain, or by Mallory’s phosphotungstic-acid hæmatoxylin stain for neuroglia. (See below.)

1. METHODS FOR STAINING GANGLION CELLS.

A. Lenhossék’s Method.

1. Fix in equal parts of saturated watery picric acid and mercuric chloride (Rabl’s mixture); after-harden in absolute alcohol; imbed in paraffin; cut.

2. Stain in saturated watery solution of toluidin blue, or thionin blue, for 12 hours.

3. Wash rapidly in water.

4. Differentiate carefully in absolute alcohol, or in aniline-alcohol (1-10).

5. Carbol xylol; xylol (quickly); balsam.

Nissl’s granules are blue. This method is easy, and the best for general work.

B. Nissl’s Method.

1. Fix in 96 per cent alcohol for 2-5 days; mount tissue in gum arabic on block; harden in 96 per cent alcohol; cut; place sections in 96 per cent alcohol.

2. Stain in methylene-blue soap mixture (methylene-blue B 3.75 grms., Venetian soap shavings 1.75 grms., water 1,000 cc. Shake well. Keep 3 months before using. Shake and filter before using.), warming, until bubbles arise.

3. Differentiate in aniline alcohol (aniline oil 10 parts, 96 per cent alcohol 90 parts) very rapidly.

4. Arrange section on slide; dry with blotting-paper; cover with cajuput oil.

5. Blot; wash off oil with benzene.

6. Remove benzene; mount the wet section in xylol colophonium, slightly warming; press on cover, and remove excess of colophonium.

Nuclei of ganglion cells light blue: granules dark blue. Toluidin blue, thionin, dahlia, Bismarck brown or neutral red may be used instead of methylene-blue, and often give better results.

2. METHODS FOR STAINING MYELIN SHEATHS.

A. Weigert’s Method.

1. Fix in formol 2-3 days.

2. Primary mordant (potassium bichromate 5 grms., fluorchrom 2.5 grms., water 100 cc.: boil and filter) 4-6 days.

3. Without washing after-harden in graded alcohols, in the dark.

4. Imbed in celloidin.

5. Secondary mordant (neutral copper acetate 5 grms., fluorchrom 2.5 grms., water 100 cc., boil and add 36 per cent acetic 5 cc.) for 1 day at 37°C.

6. Transfer to 70-80 per cent alcohol.

7. Cut.

8. Stain in Weigert’s iron-hæmatoxylin, 24 hours.

9. Wash in water, 30 minutes or longer.

10. Differentiate in borax-potassium ferricyanide (potassium ferricyanide 2.5 grms., borax 2 grms., water 100 cc.) until the gray substance appears yellow to white. Control under microscope.

11. Wash thoroughly in water.

12. Dehydrate in absolute alcohol; clear in xylol; mount in balsam.

Myelinated fibres, blue-black, upon a colorless or light yellow background; red blood cells may be blue-black. Weigert’s method is better than any of the numerous modifications.

B. Orr’s Osmic-Acid Method.

1. Place fresh tissue from cerebral cortex or cord, not more than ⅛ inch in thickness, in 1 per cent acetic, 2 cc., and 2 per cent osmic acid 8 cc., for 48 hours. If mixture is darkened at end of 24 hours, renew.

2. Transfer to 10 per cent formol for 3 days, in order to complete reduction and hardening.

3. Imbed in celloidin or paraffin; cut.

4. Remove paraffin; alcohol; water; differentiate in ⅛-¹/₁₂ per cent potassium permanganate.

5. Transfer to a 1 per cent oxalic acid solution, until sections become yellowish-green.

6. Wash; dehydrate; xylol; balsam.

Nerves and fat black: tissue yellowish-green. A very reliable method.

3. STAINING OF AXIS CYLINDERS.

Stain with Van Gieson’s (red), Mallory’s aniline blue method (red), lithium carmine (red), or

Williamson’s Modification of Bielschowsky’s Method.

1. Fix in Müller’s; imbed; cut.

2. Place sections in 10 cc. of tap water containing a few drops of formalin, 5 minutes.

3. Wash in water.

4. Place in the following silver bath 5-10 minutes: 3 drops of liq. ammoniæ (B.P.) are dropped into a test tube. Add 10 per cent silver nitrate solution, drop by drop, until a brownish precipitate is formed. Dissolve the latter by adding ammonia, drop by drop, until the fluid is quite clear. Add tap water up to 10 cc.

5. Wash thoroughly in water.

6. Transfer to the dilute formol solution until sections become grayish-black (1-3 minutes).

7. Place in the following solution for a few minutes: To 10 cc. of water add 2 drops of 1 per cent watery solution of chloride of gold, a few drops of saturated borax solution, and a few drops of a 10 per cent solution of potassium carbonate.

8. Transfer to a 10 per cent aqueous solution of sodium hyposulphite for a few minutes.

9. Wash in water; dehydrate in alcohol; clear in oil of cajuput; xylol; balsam.

Axis-cylinders, intracellular fibrils and Golgi’s network are stained.

4. STAINING OF NEURO-FIBRILLAR STRUCTURES.

These are stained by Bielschowsky’s method, and by acid fuchsin after fixation with osmic acid. The special methods (Apathy’s gold method, Bethe’s molybdic method, the silver methods of Ramen y Cajal and Robertson) have little practical application in pathologic work, and are used chiefly in the study of normal histology.

5. THE STAINING OF NEUROGLIA.

A. Weigert’s Method.

1. Fix small pieces of fresh tissue in 10 per cent formol for 24 hours.

2. Mordant. 8 days at room temperature (4 days at 37°C.) in copper acetate 5 grms., fluorchrom 2.5 grms., acetic acid 5 cc., water 100 cc.

Or, harden and mordant at the same time in 9 parts of the copper mordant, and 1 part of commercial formol for 8 days, changing on the second day, and once again later.

3. Wash in water: after-harden in alcohol; imbed in celloidin; cut.

4. Place sections in ⅓ per cent aqueous solution of potassium permanganate.

5. Wash in two changes of water.

6. Place in the following reducing mixture, 2-4 hours: Chromogen 5 grms., formic acid (sp. gr. 1.2) 5 cc., water 100 cc.; filter; to 90 cc. add 10 cc. of 10 per cent sodium sulphite solution just before using.

7. Wash twice in water.

8. Place sections in 5 per cent carefully filtered aqueous chromogen solution 10-12 hours. (The glia fibres become darker, and a yellowish contrast is obtained for the ganglion and ependymal cells and thicker axis cylinders. Connective-tissue is stained red.)

9. Wash in water.

10. Place section on a slide freshly cleaned with alcohol; dry with filter paper; stain in the following mixture for about 30 seconds: Saturated solution of methyl violet in 70-80 per cent alcohol 100 cc., oxalic acid 5 per cent solution, 5 cc.

11. Remove excess of stain; dry with filter paper; cover slide with saturated solution of iodine in 5 per cent potassium-iodide solution, 30 seconds.

12. Remove iodine solution; dry with filter paper; differentiate in a mixture of equal parts aniline oil and xylol until no more heavy clouds of stain are given off. Control under microscope.

13. Dry section with filter-paper; add xylol; blot; repeat three times.

14. Mount in balsam or turpentine colophonium.

Neuroglia fibres and nuclei, blue; connective-tissue, blue-violet; thicker myelin sheaths, ganglion and ependymal cells, yellowish. This is the best method, none of the modifications giving as good results. No method, however, will stain every neuroglia-fibre.

B. Mallory’s Neuroglia Method.

1. Fix small pieces in 10 per cent formol, 4 days.

2. After-harden in saturated watery picric solution 4-8 days: or combine 1 and 2 by using formol 10 cc. with 90 cc. saturated picric acid solution.

3. Place in a 5 per cent aqueous solution of ammonium bichromate, 4-7 days at 37°C., changing solution 011 second day; or 3-4 weeks at room-temperature.

4. Without washing, harden in alcohol; imbed in celloidin or paraffin; cut.

5. Place sections in ¼ per cent aqueous solution of potassium permanganate, 15-30 minutes.

6. Wash in water.

7. Immerse in 5 per cent aqueous oxalic acid, 5-30 minutes.

8. Wash in several changes of water.

9. Stain in following solution, 1-several days: Hæmatoxylin 0.1 grm., 10 per cent phosphotungstic acid 20 cc., hydrogen peroxide 0.2 cc., water 80 cc.

10. Wash rapidly in water.

11. Differentiate in freshly prepared 30 per cent alcoholic solution of ferric chloride, 5-20 minutes.

12. Wash in water.

13. Dehydrate in 95 per cent and absolute alcohol or blot with xylol.

14. Clear in xylol; balsam.

When Zenker’s fixation is used, omit 2 and 3, and after cutting sections treat with Lugol’s to remove mercury and then with 95 per cent alcohol to wash out iodine; then wash in water and proceed with 5.

Neuroglia, nuclei and fibrin, dark-blue; all else is pale yellow or gray. If the differentiation in 11 is omitted, the axis-cylinders and ganglion-cells are rose-pink; the connective-tissues, dark red-pink.

6. COMBINED STAINING OF SEVERAL NERVOUS STRUCTURES.

Various methods of impregnation with silver, gold or lead are used in histologic work, the Golgi methods and their modifications in particular. They have but little application in pathologic work, and for that reason are omitted here, as is also a consideration of Ehrlich’s vital methylene-blue method and its modifications. Full details of these methods can be found in laboratory textbooks on histology.

7. METHODS FOR THE DEMONSTRATION OF NERVE-DEGENERATION.

A. Marchi’s Method.

1. Harden small fresh pieces of tissue in Müller’s fluid for at least 8 days. Handle tissue very carefully to prevent mechanical injury. The tissue may be placed first in formol, and then later transferred to Müller’s fluid.

2. Place in freshly prepared Marchi’s fluid (Müller’s fluid 2 parts, 1 per cent osmic acid solution 1 part) for about 8 days in the incubator at 37°C. The brain requires a longer time. When the mixture loses the osmic acid smell renew it.

3. Wash in running water, 24 hours.

4. Harden in graded alcohols.

5. Imbed in celloidin; cut; dehydrate; clear; mount.

Degenerated nervous tissue (fat) is black: all else brownish gray. Contrast stain in Van Gieson’s, lithium carmine, etc. This method is good for the demonstration of early degenerations.

B. Donaggio’s Methods for Early Degeneration.

Method I—

1. Fix in Müller’s fluid or in 4 per cent potassium-bichromate solution. The tissue may remain in the fluid for any length of time.

2. Transfer directly, without washing, to alcohol. Dehydrate; imbed in celloidin; cut sections 20-30 microns. Place sections in distilled water for a few seconds.

3. Transfer to the following mixture for 10-20 minutes: To 20 per cent solution of ammoniated chloride of tin add an equal amount of 1 per cent aqueous hæmatoxylin. Allow to stand for five days. Keep in the dark, and in a cool place.

4. Wash rapidly in distilled water.

5. Differentiate in Pal’s solution (oxalic acid 0.5 grm., potassium sulphite 0.5 grm., water 100 cc.) until the normal fibres are entirely decolorized.

6. Dehydrate; xylol; neutral balsam.

Degenerated fibres blue; normal, decolorized.

Method II—

1. Fix in Müller’s fluid: imbed as in Method I.

2. Place sections in 0.5-1 per cent aqueous hæmatoxylin solution, 10-20 minutes.

3. Transfer directly to a saturated aqueous solution of neutral acetate of copper, 30 minutes. Renew copper solution once.

4. Decolorize as in Method I.

5. Wash rapidly in distilled water.

6. Dehydrate in graded alcohols; xylol; balsam.

Degenerated fibres black; normal fibres unstained, except for a narrow circle at periphery.

Method III—

1. Fix and imbed as in Method I.

2. Stain in 0.5-1 per cent aqueous hæmaloxylin solution, 10-20 minutes.

3. Transfer directly to 10-20 per cent solution of perchloride of iron. The section becomes black. After a few seconds they lose their color. If washed in water, they regain their color.

4. Without washing, differentiate in acid alcohol (0.75 cc. HCl in 100 cc. alcohol).

5. Dehydrate in absolute alcohol; xylol; balsam.

Degenerated fibres appear as small black streaks or circular areas.

C. Staining of Fat-granule Cells.

Fix in Flemming’s or Marchi’s mixtures; or in formol, staining with sudan III or scharlach R. The tissues may be examined also in the fresh condition.

D. Old Degenerations.

Use Weigert’s myelin method to show absence of myelinated fibres. Van Gieson’s method stains the neuroglia of degenerated areas a deep red; it is very useful combined with Weigert’s myelin stain. Weigert’s neuroglia stain may be used to demonstrate the relative parts played by neuroglia and connective-tissue in the formation of sclerotic patches. When Weigert’s iron-hæmatoxylin is used with Van Gieson’s the neuroglia remains unstained, while the connective-tissue stains red. With other hæmatoxylins the neuroglia cannot be sharply differentiated from connective-tissue when stained with Van Gieson’s.

8. PERIPHERAL NERVES.

Use any of the above methods for the staining of myelin sheaths, ganglion cells, axis cylinders, etc. Van Gieson’s is good for the demonstration of connective-tissue increase. For the demonstration of peripheral fibrils and nerve-endings consult textbooks on histology for Golgi methods, Ehrlich’s vital methylene-blue method, and the modifications of May, Drasch, and others.

Platner’s Method.

1. Harden in 25 per cent solution of liq. ferri sesquichlor., 1-5 days.

2. Wash in water, until the addition of KCNS to the water yields no reaction.

3. Place in 75 per cent alcohol containing an excess of di-nitroresorcin, 2-30 days, according to the size of the piece of tissue.

4. Dehydrate in absolute alcohol.

5. Imbed; cut; dehydrate; clear; mount.

Axis cylinders, emerald green. A good method for the rapid demonstration of pathological processes connected with the peripheral nerves.

9. CHROMAFFINIC TISSUES.

Wiesel’s Method.

1. Fix 1-4 days in 10 parts of a 5 per cent potassium bichromate solution, 20 parts of 10 per cent formol, 20 parts distilled water.

2. 1-2 days in 5 per cent potassium bichromate.

3. Wash thoroughly in running water, 24 hours; harden in graded alcohols; imbed in paraffin.

4. Stain sections in a 1 per cent aqueous toluidin-blue or water-blue solution.

5. 5 minutes in tap-water.

6. Stain 20 minutes in a 1 per cent watery safranin solution.

7. 96 per cent, and then absolute alcohol, until the blue color appears.

8. Xylol; balsam.

Chromaffinic cells green; other cells light blue, nuclei red.

X. SKIN.

Skin should be fixed in formol-Müller’s or formol, and should not be left too long in alcohol or xylol. Imbed in celloidin preferably. For general use the ordinary stains suffice; for the study of pigment stain with alum or lithium-carmine. Use Weigert’s elastic-tissue stain for the demonstration of elastic fibrils. The intercellular bridges may be demonstrated by Van Gieson’s (remaining unstained), or by various special staining methods.

Herxheimer’s Method for Epithelial Fibrillae.

1. Harden; imbed; cut.

2. Stain in a saturated watery solution of kresyl-echt-violett.

3. Dehydrate in absolute alcohol; clear in clove oil; balsam.