Water 10 c.c.
(b) _Stain._--
Saturated aqueous solution of alum 10 c.c.
Saturated alcoholic solution of gentian violet 1 c.c.
Distilled water 5 c.c.
Mix equal parts of a and b before using.
1. Prepare and fix the film in the manner described above.
2. Boil the mixture and immerse the cover-slip in it, whilst
still hot, for one minute.
3. Wash in water.
4. Examine in water; if satisfactory, dry and mount in
Canada balsam.
~3. MacCrorrie's Method.~--
_Mordant-Stain._--
Measure out and mix.
Night blue, saturated alcoholic solution 10 c.c.
Potash alum, saturated aqueous solution 10 c.c.
Tannin, 10 per cent. aqueous solution 10 c.c.
NOTE.--The addition of gallic acid, 0.1 to 0.2 gramme, may
improve the solution, but is not necessary.
METHOD.--
1. Prepare and fix the films as above.
2. Pour some of the mordant-stain on the film and warm
gently, high above the flame, for two minutes (or place in
the "hot" incubator for a like period).
3. Wash thoroughly in water.
4. Dry and mount.
~4. Loeffler's Method.~--
(a) _Mordant._--
Tannic acid, 20 per cent. aqueous solution 10 c.c.
Ferrous sulphate, saturated aqueous solution 5 c.c.
Haematoxylin solution 3 c.c.
Carbolic acid, 1 per cent. aqueous solution 4 c.c.
This solution must be freshly prepared.
_Haematoxylin solution_ is prepared by boiling 1 gramme
logwood
with 8 c.c. distilled water, filtering and replacing the loss from
evaporation.
_Alternative Mordant_ (Bunge's Mordant).--
Tannic acid, 20 per cent. aqueous solution 10 c.c.
Ferrous sulphate, saturated aqueous solution 5 c.c.
Fuchsin, saturated alcoholic solution 1 c.c.
(b) _Stain._--
Weigh out
Methylene-blue }
Or methylene-violet } 4 grammes
Or fuchsin }
and dissolve in
Aniline water, freshly saturated and filtered 100 c.c.
METHOD.--
1. Prepare and fix films as above.
2. Pour the mordant on to the film and warm cautiously over the flame
till steam rises; keep the mordant gently steaming for one minute.
3. Wash well in distilled water till no more colour is discharged; if
necessary, wash carefully with absolute alcohol.
4. Filter a few drops of the stain on to the film, warm as before, and
allow the steaming stain to act for one minute.
5. Wash well in distilled water.
6. Dry and mount.
NOTE.--The flagella of some organisms can be demonstrated better by
means of an alkaline stain or an acid stain--a point to be determined
for each. Speaking generally, those bacilli which give rise to an acid
reaction in the culture medium require an alkali; those which form
alkali in cultivation require an acid. According to requirements,
therefore, Loeffler recommends the addition of sodium hydrate, 1 per
cent. aqueous solution, 1 c.c.; or an equal quantity of an exactly
comparable solution of sulphuric acid.
~5. Van Ermengem's Method.~--This method, being merely a precipitation of
a silver salt on the micro-organisms and not a true stain, creates a
false impression as to the relative proportions of bacteria and
flagella.
(a) _Fixing Fluid._--
Osmic acid, 2 per cent. aqueous solution 10 c.c.
Tannic acid, 20 per cent. aqueous solution 20 c.c.
Acetic acid, glacial 1 c.c.
The fixing fluid should be prepared some days before use and
filtered as required. In colour it should be distinctly
violet.
(b) _Sensitising Solution._--
Silver nitrate, 0.5 per cent. aqueous solution.
This solution must be kept in a dark blue glass bottle or in
a dark cupboard.
Filter immediately before use.
(c) _Reducing Solution._--
Weigh out
Gallic acid 5 grammes
Tannic acid 3 grammes
Potassium acetate, fused 10 grammes
and dissolve in
Distilled water 350 c.c.
Filter.
This solution will keep active for several days, but fresh
solution must be used for each preparation.
METHOD.--
1. Prepare emulsion, make and fix films as above in the
preceding method, steps 1 to 4.
2. Pour on the film as much of the fixing solution as the
cover-glass will hold, heat carefully over the flame till
steam rises, and allow the steaming fixing fluid to act for
five minutes.
3. Wash well in water.
4. Wash in absolute alcohol.
5. Wash in distilled water.
6. Pour some of the sensitising solution on the film and
allow it to act for from thirty seconds to one minute; blot
off the excess of fluid with filter paper.
7. Without washing, transfer the film to a watch-glass
containing the reducing solution and allow it to remain
therein for from thirty seconds to one minute; blot off the
excess of fluid with filter paper.
8. Without washing, again treat the film with the
sensitising solution, this time until the film commences to
turn black.
9. Wash in distilled water.
10. Dry and mount.
~To Stain Nuclei of Yeast Cells.~
1. Prepare and fix film in the usual manner.
2. Soak in ferric ammonia sulphate 3 per cent. aqueous solution for two
hours.
3. Wash thoroughly in water.
4. Stain in haematoxylin solution (see page 95) for thirty minutes.
5. Wash in water.
6. Differentiate in ferric ammonia sulphate solution for 1-1/2-2
minutes, examining wet under microscope during the process.
~To Stain Spores.~
~1. Single Stain.~--
1. Prepare cover-slip film in the usual way.
2. In fixing, pass the cover-slip film fifteen or thirty times through
the flame instead of only three. This destroys the resisting power of
the spore membrane and allows the stain to reach the interior.
3. Stain in the usual way with methylene-blue or fuchsin.
4. Wash in water.
5. Dry and mount.
~2. Double Stain.~--
1. Prepare and fix film in the usual way--i. e., pass three times
through flame to fix.
2. Cover the film with hot carbol-fuchsin and hold in the forceps above
a small flame until the fluid begins to steam. Set the cover-slip down
and allow it to cool. Repeat the process when the stain ceases to steam
and continue to repeat until the stain has been in contact with the film
for twenty minutes. (This stains both spores and bacteria.)
3. Wash in water.
4. Decolourise in alcohol, 2 parts; acetic acid, 1 per cent., 1 part.
(This removes the stain from everything but the spores.)
5. Wash in water.
6. Mount the cover-slip in water and examine microscopically with the
1/6-inch objective. (Spores should be red, and the rest of the film
colourless or a very light pink.) If satisfactory, pass on to section 7;
if unsatisfactory, repeat steps 2 to 5.
7. Counterstain in weak methylene-blue. (Now spores red, bacilli blue.)
8. Wash in water.
9. Dry and mount.
The spores of different bacilli differ greatly in their resistance to
decolourising reagents; even the spores of the same species of organisms
vary according to their age. Young spores are more easily decolourised
than those more mature.
Sulphuric acid, 1 per cent. aqueous solution, and hydrochloric acid, 0.5
per cent. alcoholic (90 per cent.) solution, are useful decolourising
reagents.
~3. Moeller's Method.~--
1. Prepare and fix films in the usual manner.
2. Immerse in absolute alcohol for two minutes, then in
chloroform for two minutes; wash in water. This dissolves
out any fat or crystals that might otherwise retain the
"spore" stain.
3. Immerse in chromic acid, 5 per cent. aqueous solution,
for one minute; wash in water.
4. Pour Ziehl's carbolic fuchsin on the film, warm as in
previous methods, and allow it to act for ten minutes.
5. Wash in water.
6. Decolourise in sulphuric acid, 5 per cent. aqueous
solution, for five seconds.
7. Wash in water.
8. Counterstain with Kuehne's carbolic methylene-blue for
one or two minutes.
9. Wash in water.
10. Dry and mount.
(Spores red, bacilli blue.)
~4. Abbott's Method.~--
1. Prepare and fix films in the usual manner.
2. Pour Loeffler's alkaline methylene-blue on the film; warm
cautiously over the flame till steam rises and allow the hot
steam to act for one to five minutes.
3. Wash thoroughly in water.
4. Decolourise in nitric acid, 2 per cent. alcoholic
(alcohol 80 per cent.) solution.
5. Wash thoroughly in water.
6. Counterstain in eosin, 1 per cent. aqueous solution.
7. Wash.
8. Dry and mount.
(Spores blue, bacilli red.)
DIFFERENTIAL METHODS OF STAINING.
~Gram's Method.~--This method depends upon the fact that the protoplasm of
some bacteria permits aniline gentian violet and Lugol's iodine
solution, when applied consecutively, to enter into a chemical
combination which results in the formation of a new blue-black pigment,
only very sparingly soluble in absolute alcohol. Such organisms are said
to "stain by Gram," or to be "Gram positive."
1. Prepare a cover-slip film and fix in the usual way.
2. Stain in aniline gentian violet three to five minutes. Filter as much
aniline water on to the cover-slip as it will hold; then add the
smallest quantity of alcoholic solution of gentian violet which suffices
to saturate the aniline water and form a "bronze scum" upon its
surface--if too much of the alcoholic gentian violet is added the
alcohol present redissolves this scum.
To prepare aniline water, pour 4 or 5 c.c. aniline oil into
a stoppered bottle and add distilled water, 100 c.c. Shake
vigourously and filter immediately before use. The excess of
oil sinks to the bottom of the bottle and may be used again.
3. Wash in water.
4. Treat with Lugol's iodine solution until the film is black or dark
brown.
To do this treat with iodine solution for a few seconds, wash in water,
and examine the film over a piece of white filter paper. Note the
colour. Repeat this process until the film ceases to darken with the
fresh application of iodine solution.
Lugol's solution is prepared by dissolving
Iodine 1 gramme
Iodide of potassium 3 grammes
In distilled water 300 c.c.
5. Wash in water.
6. Wash with alcohol until no more colour is discharged and the alcohol
runs away clear and colourless.
The following mixture may be substituted for absolute alcohol as a
decolouriser
Acetone 10 c.c.
Absolute alcohol 100 c.c.
7. Wash in water.
8. Counterstain very lightly with aqueous solution of Neutral Red. Other
counterstains may be used such as dilute eosin, dilute fuchsin, or
vesuvin.
NOTE.--This section may be omitted when dealing with films
prepared from pure cultivations.
9. Wash in water.
10. Dry and mount.
~Gram-Claudius Method.~--
1. Prepare a cover-slip film and fix in the usual way.
2. Stain in methyl violet, 1 per cent. aqueous solution for three to
five minutes.
3. Treat with two lots picric acid, saturated aqueous solution.
4. Wash in water and dry.
5. Decolourise with clove oil.
6. Wash off clove oil with xylol.
7. Mount in xylol balsam.
~Gram-Weigert Method.~--
1-5. Proceed as for the corresponding sections of Gram's method (_quod
vide_).
6. Dry in the air.
7. Wash in aniline oil, 1 part, xylol, 2 parts, until no more colour is
discharged.
8. Wash in xylol.
9. Mount in xylol balsam.
~Modified Gram-Weigert Method.~--(To demonstrate trichophyta in hair.)
1. Soak the hairs in ether for ten minutes to remove the fat.
2. Stain thirty minutes in a tar-like solution of aniline gentian violet
(prepared by adding 15 drops of the alcoholic solution of gentian violet
to 3 drops of aniline water).
3. Dry the hairs between pieces of blotting paper.
4. Treat with perfectly fresh iodine solution.
5. Again dry between blotting paper.
6. Treat with aniline oil to remove excess of stain. (If necessary, add
a drop or two of nitric acid to the oil.)
7. Again treat with aniline oil.
8. Treat with aniline oil and xylol, equal parts.
9. Clear with xylol.
10. Mount in xylol balsam.
To obtain the best differentiation the preparation should be repeatedly
examined microscopically (with a 1/6-inch objective) between steps 5 and
9, as the actual time involved varies with different specimens.
~Ziehl-Neelsen's Method.~--(To demonstrate tubercle and other acid-fast
bacilli.)
1. Smear a thin, even film of the specimen on the cover-slip by means of
the platinum loop. (In the case of sputum, if it is a very watery
specimen, allow the film to dry, then spread a second and even a third
layer over the first.)
2. Fix by passing three times through the flame.
3. Stain in hot carbol-fuchsin (as in staining for spores) for five to
ten minutes. (This stains everything on the film.) Avoid over-heating.
4. Decolourise by dipping in sulphuric acid, 25 per cent. (This removes
stain from everything but acid-fast bacilli; e. g., tubercle, leprosy,
and smegma bacilli and the film turns yellow.)
5. Wash in water. (A pale red colour returns to the film).
6. Wash in alcohol till no more colour is discharged. (This often, but
not invariably, removes the stain from acid-fast bacilli other than
tubercle; e. g., smegma bacillus.)
7. Wash in water.
8. Counterstain in weak methylene-blue. (Stains non-acid-fast bacilli,
leucocytes, epithelial cells, etc.)
9. Wash in water, dry, and mount.
~Pappenheim's Method.~--
This method is supposed to differentiate between B. tuberculosis and
other acid-fast micro-organisms.
1. Prepare and fix film in the usual way.
2. Stain in carbol-fuchsin _without heat_ for three minutes.
3. Without previously washing in water treat the film with three or four
successive applications of corallin (Rosolic acid) solution.
Corallin 1 gramme
Methylene-blue
(saturated alcoholic solution) 100 c.c.
Glycerine 20 c.c.
4. Wash in water.
5. Dry and mount.
~Neisser's Method--Modified.~--(To demonstrate diphtheroid bacilli.)
_Stain I._--
Measure out and mix
Methylene-blue, saturated alcoholic solution 4.0 c.c.
Acetic acid, 5 per cent. aqueous solution 96.0 c.c.
Filter.
_Stain II._--
Weigh out
Neutral red 2.5 grammes
and dissolve in
Distilled water 1000 c.c.
Filter.
METHOD.--
1. Prepare and fix films in the usual way.
2. Pour stain I on the film and allow it to act for two minutes.
3. Wash thoroughly in water.
4. Treat with Lugol's iodine for ten seconds.
5. Wash thoroughly in water.
6. Pour stain II on to the film and allow it to act for thirty seconds.
7. Wash thoroughly in water.
8. Dry and mount.
NOTE.--The cultivation from which the films are prepared
must be upon blood-serum which has been incubated at 37 deg. C.
for from nine to eighteen hours.
The bacilli are stained a light red by the neutral red, which contrasts
well with the two or three black spots, situated at the poles and
occasionally one in the centre representing protoplasmic aggregations (?
metachromatic granules) stained by the acid methylene-blue.
~Wheal and Chown (Oxford) Method.~--(To demonstrate
actinomyces.)
1. Stain briefly with Ehrlich's haematoxylin (until nuclei
are faint blue after washing with tap water).
2. Wash in tap water.
3. Stain in hot carbol-fuchsin (as for tubercle bacilli) for
five to ten minutes.
4. Wash in tap water.
5. Decolourise with Spengler's picric acid alcohol. This is
prepared by mixing:
Alcohol, absolute 20 c.c.
Picric acid, saturated aqueous solution 10 c.c.
Distilled water 10 c.c.
During the progress of steps 1-5 the preparation must be
repeatedly examined microscopically with the 1/6-inch
objective.
When properly differentiated the clubs appear brilliant red
on greenish ground.
6. Dehydrate in alcohol.
7. Clear in xylol.
8. Mount in xylol balsam.
This method serves equally well for films and for sections.
VII. METHODS OF DEMONSTRATING BACTERIA IN TISSUES.
For bacteriological purposes, sections of tissue are most conveniently
prepared by either the ~freezing method~ or the ~paraffin method~.
The latter is decidedly preferable, but as it is of greater importance
to demonstrate the bacteria, if such are present, than to preserve the
tissue elements unaltered, the "frozen" sections are often of value.
Whichever method is selected, it is necessary to take small pieces of
the tissue for sectioning,--2 to 5 mm. cubes when possible, but in any
case not exceeding half a centimetre in thickness. Post-mortem material
should be secured as soon after the death of the animal as possible.
The tissue is prepared for cutting by--
(a) Fixation; that is, by causing the death of the cellular elements
in such a manner that they retain their characteristic shape and form.
The fixing fluids in general use are: Absolute alcohol; corrosive
sublimate, saturated aqueous solution; corrosive sublimate, Lang's
solution (_vide_ page 82); formaldehyde, 4 per cent. aqueous solution.
(Of these, Lang's corrosive sublimate solution is decidedly the best
all-round "fixative.")
(b) Hardening; that is, by rendering the tissue of sufficient
consistency to admit of thin slices or "sections" being cut from it.
This is effected by passing the tissue successively through alcohols of
gradually increasing strength: 30 per cent. alcohol, 50 per cent.
alcohol, 75 per cent. alcohol, 90 per cent. alcohol, absolute alcohol.
In both these processes a large excess of fluid should always be used.
FREEZING METHOD.
1. ~Fixation.~ Place the pieces of tissue in a wide-mouthed glass bottle
and fill with absolute alcohol. Allow the tissues to remain therein for
twenty-four hours.
2. ~Hardening.~ Remove the alcohol (no longer absolute, as it has taken up
water from the tissues) from the bottle and replace it with fresh
absolute alcohol. Allow the tissues to remain therein for twenty-four
hours.
[Illustration: FIG. 71.--Washing tissues.]
NOTE.--If not needed for cutting immediately, the hardened
tissues can be stored in 75 per cent. alcohol.
3. Remove the alcohol from the tissues by soaking in water from one to
two hours. Remove the stopper from the bottle; rest a glass funnel in
the open mouth and place under a tap of running water. The water of
course, overflows, but the tissues remain in the bottle (Fig. 71).
4. Impregnate the tissues with mucilage for twelve to twenty-four hours,
according to size. Transfer the pieces of tissue to a bottle containing
sterilised gum mixture.
~Formula.~--
Gum arabic 5 grammes
Saccharose 1 gramme
Boric acid 1 gramme
Water 100 c.c.
5. Place the tissue on the plate of a freezing microtome (Cathcart's is
perhaps the best form), cover and surround with fresh gum mixture;
freeze with ether, or for preference, carbon dioxide, and cut sections.
6. Float the sections off the knife into a glass dish containing tepid
water and allow them to remain therein for about an hour to dissolve out
the gum.
(If not required at once, store in 90 per cent. alcohol.)
7. Transfer to a glass capsule containing the selected staining fluid,
by means of a section lifter.
8. Transfer the sections in turn to a capsule containing absolute
alcohol (to dehydrate) and to one containing xylol or oil of cloves (to
clear).
9. Mount in xylol balsam.
_Alternative Rapid Method._--
1. Cut very small blocks of the tissue.
2. Fix in formalin 10 per cent. aqueous solution (fixation
fluid No. 7, page 82) for 24 hours.
3. Transfer block to plate of freezing microtome and freeze
with carbon dioxide vapour.
4. Float the sections off the knife into a glass dish of
tepid water.
5. Stain the sections in glass capsules containing selected
stains.
6. Place the stained section in a dish of clean water and
introduce a glass slide obliquely beneath the section; with
a mounted needle draw the section on to the slide and hold
it there; gently remove the slide from the water, taking
care that any folds in the section are floated out before
the slide is finally removed from the water.
7. Drain away as much water as possible from the section.
Drop absolute alcohol on to the section from a drop bottle,
to dehydrate it.
8. Double a piece of blotting paper and gently press it on
the section to dry it.
9. Drop on xylol to clear the section.
10. Place a large drop of xylol balsam on the section and
carefully lower a cover-glass on to the balsam.
PARAFFIN METHOD.
1. ~Fixation.~ Place the pieces of tissue, resting on cotton-wool, in a
wide-mouthed glass bottle. Pour on a sufficient quantity of the
corrosive sublimate fixing fluid; allow the tissue to remain therein for
twelve to twenty-four hours according to size.
2. Pour off the fixing fluid and wash thoroughly in running water for
twenty minutes to half an hour to remove the excess of corrosive
sublimate.
[Illustration: FIG. 72.--~L~-shaped brass moulds.]
[Illustration: FIG. 73.--Paraffin kettle.]
3. ~Hardening.~ Place the tissues in each of the following strengths of
alcohol in turn for from twelve to twenty-four hours: 50 per cent., 75
per cent., 90 per cent., absolute.
4. ~Dehydration~ is effected by transferring the tissues to fresh absolute
alcohol.
5. ~Clearing.~ Half fill a wide-mouthed bottle with chloroform. On the
surface of the chloroform float a layer of absolute alcohol about five
to ten millimetres in depth. Place the pieces of tissue in the layer of
alcohol and when they have sunk through this layer, transfer them to
pure chloroform for from six to twenty-four hours according to the size
of the pieces. When "cleared," the tissue becomes more or less
transparent.
6. ~Infiltration.~ Place the cleared tissues in fresh chloroform with
several pieces of paraffin wax and stand in a warm place, such as on the
top of the warm incubator. The warmth gradually melts the paraffin and
the tissues should remain in the mixture about twenty-four hours.
7. Transfer the tissues to a vessel containing pure melted paraffin.
Place this vessel in a paraffin water-bath regulated for 2 deg. C. above
the melting-point of the paraffin used, and allow the tissues to soak for
some four to six hours to ensure complete impregnation. The paraffin
used should have a melting-point of not more than 58 deg. C. For all
ordinary purposes 54 deg. C. will be found quite high enough.
8. Imbed in fresh paraffin in a metal (or paper) mould.
(a) Arrange a pair of ~L~-shaped pieces of metal on a plate of glass to
form a rectangular trough (Fig. 72).
(b) Pour fresh melted paraffin into the mould from a special vessel
(Fig. 73).
(c) Lift the piece of tissue from the paraffin bath and arrange it in
the mould.
(d) Blow gently on the surface of the paraffin in the mould, and as
soon as a film of solid paraffin has formed, carefully lift the glass
plate on which the mould is set and lower plate and mould together into
a basin of cold water.
(e) When the block is cold, break off the metal ~L~'s; trim off the
excess of paraffin from around the tissue with a knife, taking care to
retain the rectangular shape, and store the block in a pill-box.
When several pieces of tissue have to be imbedded at one time, shapes of
stout copper, 10 cm., 5 cm., and 2.5 cm. square respectively, and 0.75
cm. deep (Fig. 74) will be found extremely useful. These placed upon
plates of glass replace the pair of L's in the above process. When the
paraffin has set firmly the screw a should be loosened to allow the
two halves of the flange b to separate slightly--this facilitates
removal of the paraffin block.
[Illustration: FIG. 74.--Paraffin mould.]
8. Cement the block on the carrier of a "paraffin" microtome (the Minot,
the Jung, or the Cambridge Rocker) with a little melted paraffin.
Greater security is obtained if the paraffin around the base of the
block is melted by means of a hot metal or glass rod.
9. Cut sections--thin, and if possible in ribbands.
~Mounting Paraffin Sections.~--
1. Place a large drop of 30 per cent. alcohol on the centre of a slide
(or cover-slip) and float the section on to the surface of the drop,
from a section lifter.
2. Hold the slide in the fingers of one hand and warm cautiously over
the flame of a Bunsen burner, touching the under surface of the glass
from time to time on the back of the other hand. As soon as the slide
feels distinctly warm to the skin, the paraffin section will flatten out
and all wrinkles disappear.
(The slide with the section floating on it may be rested on the top of
the paraffin bath for two or three minutes, instead of warming over the
flame as here described.)
3. Cautiously tilt up the slide and blot off the excess of spirit with
blotting paper, leaving the section attached to the centre of the
slide.
4. Place the slide in a wire rack (Fig. 75), section downward, in the
"hot" incubator for twelve to twenty-four hours. At the end of this time
the section is firmly adherent to the glass, and is treated during the
subsequent steps as a "fixed" cover-glass film preparation.