Practical pathology

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).

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).

3. BACTERIA STAINING BY THE TUBERCLE-BACILLUS METHOD. (ZIEHL-NEELSEN.)

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.

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:—

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.

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.

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.