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.