1. Rest the cover-slip, film side uppermost, on the rubber mat.
2. By means of a drop-bottle, cover the film side of the cover-slip with
the selected stain, allow it to act for a few minutes, then wash off the
excess in running water.
The penetrating power of stains is increased by (a) physical
means--e. g., heating the stain; (b) chemical means--e. g., by the
addition of carbolic acid, 5 per cent. aqueous solution; caustic
alkalies, 2 per cent. aqueous solutions; water saturated with aniline
oil; borax, 0.5 per cent. aqueous solution.
The most commonly used dyes for cover-slip film preparations are the
aniline dyes.
(A) Basic:
(a) Methylene-blue.
(b) Gentian violet.
(c) Fuchsin.
These dyes are kept in saturated alcoholic (90 per cent.) solutions so
that decomposition may be retarded.
Two or three drops of alcoholic solution of these dyes to, say, 4 c.c.
water, usually makes a sufficiently strong staining fluid for cover-slip
film preparations.
Carbolic methylene-blue (C.M.B.) and carbol fuchsin (C.F.) are prepared
by covering the cover-slip with 5 per cent. solution of carbolic acid
and adding a few drops of the saturated alcoholic solution of
methylene-blue or fuchsin respectively to it. For aniline gentian violet
(A.G.V.) the stain is added to a saturated solution of aniline oil in
water.
(d) Thionine blue.
(e) Bismarck brown.
(f) Neutral red.
(B) Acid:
(a) Eosin, aqueous yellowish.
(b) Safranine.
These dyes are kept in 1 per cent. aqueous solution to which is added 5
per cent. of alcohol, as a preservative. They are generally used in this
form.
A few nuclear stains (carmine, haematoxylin) are occasionally used more
especially in "section" work.
_Decolourisation._--After overstaining, films may be decolourised by
washing for a longer or shorter time in one of the following reagents
arranged in ascending order of power
1. Water.
2. Chloroform.
3. Acetic acid, 1 per cent.
4. Alcohol.
5. Alcohol absolute, } equal parts.
Acetic acid, 1 per cent., }
{Hydrochloric, 1 per cent. aqueous solution.
{Hydrochloric, 1 per cent. Alcoholic
{ (90 per cent.) solution.
6. Mineral acids: {Sulphuric, 25 per cent. aqueous solution.
{Nitric, 33 per cent. aqueous solution.
_Counterstaining._--Use colours which will contrast with the first
stain; e. g.,
Vesuvin, }
Neutral red, }for films stained by methylene-blue or
Eosin, }Gram's method.
Fuchsin, }
Methylene-blue, }for films stained by fuchsin.
Gentian violet, }
8. _Mounting._--
1. Wash the film carefully in running water.
2. Blot off the superfluous water with the filter paper, or dry more
completely between two folds of blotting paper.
3. Complete the drying in the air, or by holding the cover-slip in the
fingers at a safe distance above the flame of the Bunsen burner.
4. Place a drop of xylol balsam on the centre of a clean 3 by 1 glass
slide and invert the cover-slip over the balsam, and lower it carefully
to avoid the inclusion of air bubbles.
NOTE.--Xylol is used in preference to chloroform to dissolve
Canada balsam, as it does not decolourise the specimen.
~Impression films~ (_Klatschpraeparat_) are prepared from isolated
colonies of bacteria in order that their characteristic formation may be
examined by higher powers than can be brought to bear on the living
cultivation. They are prepared from plate cultivations (_vide_ page 230)
in the following manner.
1. Remove a clean cover-slip from the alcohol pot with sterile forceps
and burn off the spirit.
2. Open the plate and rest one edge of the cover-slip on the surface of
the medium a little to one side of the selected colony. Lower it
cautiously over the colony until horizontal. Avoid any lateral movement
or the inclusion of bubbles of air.
3. Make gentle vertical pressure on the centre of the cover-slip with
the points of the forceps to ensure perfect contact with the colony.
4. Steady one edge of the cover-slip with the forceps and pass the point
of a mounted needle just under the opposite edge and raise the
cover-slip carefully; the colony will be adherent to it. When nearly
vertical, grasp the cover-slip with the forceps and remove it from the
plate. Re-cover the plate.
5. Place the cover-slip, film uppermost, on the rubber mat, and cover
it with an inverted watch-glass until dry.
6. Fix by immersing in one of the fixing fluids previously mentioned
(_vide_ page 82).
7. Clear with acetic acid and alcohol.
8. Stain and mount as an ordinary cover-slip film preparation, being
careful to perform all washing operations with extreme gentleness.
~Microscopical Examination of the Unstained Specimens.~--
1. Place the body tube of the microscope in the vertical position.
2. Arrange the hanging-drop slide on the microscope stage so that the
drop of fluid is in the optical axis of the instrument, and secure it in
that position by means of the spring clips.
3. Use the 1/6-inch objective, rack down the body tube until the front
lens of the objective is almost in contact with the cover-slip--that is,
well within its focal distance. This is best done whilst bending down
the head to one side of the microscope, so that the eyes are on a level
with the stage.
4. Apply the eye to the ocular and adjust the plane mirror to the
position which secures the best illumination.
5. Rack the condenser down slightly and cut down the aperture of the
iris diaphragm so that the light, although even, is dim.
6. Rack up the body tube by means of the coarse adjustment until the
bacteria come into view; then focus exactly by means of the fine
adjustment.
Some difficulty is often experienced at first in finding the hanging
drop, and if the first attempt is unsuccessful, the student must not on
any account, whilst still applying his eye to the ocular, rack the body
tube down (for by so doing there is every likelihood of the front lens
of the objective being forced through the cover-glass, and not only
spoiling the specimen, but also contaminating the objective); but, on
the contrary, withdraw his eye, rack the tube up, and commence again
from step 2.
~Dark Ground Illumination.~--
1. Set up the microscope stand in the vertical position and insert the
highest eyepiece available.
2. Remove the nosepiece from the microscope tube and fit the 2/3 inch
objective in place.
3. Remove the substage condenser and replace it by the dark ground
condenser.
4. Fit up the source of illumination some 30-50 cm. distant from the
microscope. (This should be the Liliput Arc Lamp (Leitz), Nernst Lamp or
incandescent gas lamp; if either of the two latter are employed, a
bull's eye condenser to produce parallel rays must be interposed between
light and microscope); and adjust illuminant and microscope so that the
substage plane mirror is completely filled with light.
5. Focus the two concentric rings engraved upon the upper surface of the
condenser and centre them accurately by means of the centring screws.
6. Prepare a "fresh" specimen (see pages 74-76) of the material it is
desired to observe, using selected, new, 3 by 1 glass slips of less than
1 mm. thickness, and No. 1 cover-glasses (0.17 mm. thick), which should
be cleaned with a piece of soft washleather and not with the emery
paper, as scratches on the glass produce haziness in the preparation.
7. Deposit a large drop of immersion oil (or pure water) on the upper
surface of the condenser and rack it down a few millimetres.
8. Adjust the fresh preparation on the microscope stage and fasten it in
position with the stage clips.
9. Rack up the condenser until the immersion fluid makes contact with
the under surface of the slide; avoid the formation of air bubbles.
10. Adjust the substage mirror so that the light is reflected upward. A
bright spot will be seen on the fresh preparation near the centre of the
field.
11. Replace the 2/3-inch objective by the 1/12-inch oil immersion lens
which has been fitted with the special stop to reduce its N. A.; place a
drop of immersion oil upon the centre of the cover-glasses of the fresh
preparation and lower the microscope tube until the front lens of the
objective has entered the oil drop.
12. Focus the bright spot referred to in step 10. If it no longer
occupies the centre of the field, alter the angle of the substage mirror
until it does.
13. Now focus the lens accurately on the film, cautiously vary the
height of the dark ground condenser until the best position is found.
The intensely illuminated bacteria will stand out in vivid contrast to
the dark background.
[Illustration: FIG. 70.--Immersion oil bottle.]
~Microscopical Examination of the Stained Specimen.~--(The body tube of
the microscope may be vertical or inclined to an angle.)
1. Secure the slide on the stage of the microscope by means of the
spring clips.
2. Place a drop of cedarwood oil on the centre of the cover-slip.
The immersion oil is pure cedarwood oil, and is kept in a
small bottle of stout glass (Fig. 70), the cavity of which
is shaped like an inverted cone, and is provided with a
safety funnel (so that the oil does not escape if the bottle
is accidentally overturned) and a dust cap of boxwood fitted
with a wooden rod with which the drop of oil is applied to
the cover-glass or lens.
3. Use the 1/12-inch oil immersion lens of the microscope. Rack down the
body tube till the front lens of the objective is in contact with the
oil and nearly touching the cover-slip.
4. Rack up the condenser until it is in contact with the under surface
of the slide.
5. Apply the eye to the ocular and arrange the plane mirror so as to
obtain the greatest possible amount of light.
6. Rack up the body tube until the stained film comes into view.
7. Focus the condenser accurately on the film.
8. Focus the film accurately by means of the fine adjustment.
VI. STAINING METHODS.
In the following pages are collected the various "stock" stains in
everyday use in the bacteriological laboratory, together with a
selection of the most convenient and generally useful staining methods
for demonstrating particular structures or differentiating groups of
bacteria. The stains employed should either be those prepared by
Gruebler, of Leipzig, or Merck, of Darmstadt. The methods printed in
ordinary type are those which a long experience has shown to be the most
reliable, and to give the best results--those relegated to small type
comprise such as are not so generally useful, but give excellent results
in the hands of the experienced worker.
BACTERIA STAINS.
~Methylene-blue.~--
1. _Saturated Aqueous Solution._
Weigh out
Methylene-blue 1.5 grammes
Place in a stoppered bottle having a capacity of from 150 to 200 c.c.
and add
Distilled water 100.0 c.c.
Allow the water to remain in contact with the dye for two weeks, shaking
the contents of the bottle vigourously for a few moments every day.
Filter.
2. _Saturated Alcoholic Solution._
Weigh out
Methylene-blue 1.5 grammes
Place in a stoppered bottle of 150 c.c. capacity and add
Alcohol, 90 per cent 100.0 c.c.
Allow the alcohol to remain in contact with the dye for two hours,
shaking vigourously every few minutes. Filter.
3. _Carbolic Methylene-blue_ (Kuehne).
Weigh out
Methylene-blue 1.5 grammes
Carbolic acid 5.0 grammes
and dissolve in
Distilled water 100.0 c.c.
and add
Absolute alcohol 10.0 c.c.
Filter.
4. _Alkaline Methylene-blue_ (Loeffler).
Measure out and mix
Methylene-blue, saturated alcoholic solution 30.0 c.c.
Caustic potash, 0.1 per cent. aqueous solution 100.0 c.c.
Filter.
~Gentian Violet.~--
5. _Saturated Aqueous Solution._
Weigh out
Gentian violet 2.25 grammes
and proceed as in preparing the corresponding solution of
methylene-blue.
6. _Saturated Alcoholic Solution._
Weigh out
Gentian violet 5.0 grammes
and proceed as in preparing the corresponding solution of
methylene-blue.
7. _Carbolic Gentian Violet_ (Nicolle).
Measure out and mix
Gentian violet, saturated alcoholic solution 10.0 c.c.
Carbolic acid, 1 per cent. aqueous solution 100.0 c.c.
Filter.
8. _Anilin Water Solution_ (Koch-Ehrlich).
Measure out
Distilled water 100 c.c.
Add anilin oil drop by drop (shaking well after the addition of each
drop) until the solution is opaque.
Filter until clear.
and add
Absolute alcohol 10 c.c.
Saturated alcoholic solution gentian violet 11 c.c.
Filter.
NOTE.--This solution will not keep longer than 14 days.
~Thionine Blue (or Lauth's Violet).~--
9. _Carbolic Thionine Blue_ (Nicolle).
Weigh out
Thionine blue 1.0 gramme
Carbolic acid 2.5 grammes
and dissolve in
Distilled water 100.0 c.c.
Filter.
Before use dilute with equal quantity of distilled water and again
filter.
~Fuchsin (Basic).~--
10. _Saturated Aqueous Solution._
Weigh out
Basic fuchsin 1.5 grammes
and proceed as in preparing the corresponding solution of methylene-blue
(_q. v._).
11. _Saturated Alcoholic Solution._
Weigh out
Basic fuchsin 3.5 grammes
and proceed as in preparing the corresponding solution of
methylene-blue.
12. _Carbolic Fuchsin_ (Ziehl).
Weigh out
Basic fuchsin 1.0 gramme
Carbolic acid 5.0 grammes
dissolve in
Distilled water 100.0 c.c.
and add
Absolute alcohol 10.0 c.c.
Filter.
CONTRAST STAINS.
~Eosin.~--There are several commercial varieties of eosin, which, from the
bacteriological point of view, possess very different values. Gruebler
lists four varieties, of which two only are useful for bacteriological
work:
Eosin, aqueous yellowish.
Eosin, aqueous bluish.
13. _Eosin Aqueous Solution_ (Yellowish or Bluish Shade), 1 per cent.
Weigh out
Eosin, aqueous 1.0 gramme
dissolve in
Distilled water 100.0 c.c.
and add
Absolute alcohol 5.0 c.c.
Filter.
14. _Eosin Alcoholic Solution_, 0.5 per cent.
Weigh out
Eosin, alcoholic 0.5 gramme
and dissolve in
Alcohol (70 per cent.) 100.0 c.c.
Filter.
~Safranine.~--
15. _Aqueous Solution._
Weigh out.
Safranine 0.5 gramme
and dissolve in
Distilled water 100.0 c.c.
Filter.
~Neutral Red.~--
16. _Aqueous Solution._
Weigh out
Neutral red 1.0 gramme
and dissolve in
Distilled water 100.0 c.c.
Filter.
~Vesuvin (or Bismarck Brown).~--
17. _Saturated Aqueous Solution._
Weigh out
Vesuvin 0.5 gramme
and dissolve in
Distilled water 100.0 c.c.
Filter.
TISSUE STAINS.
~Aniline Gentian Violet~ (For Weigert's Fibrin Stain).--
Weigh out
Gentian violet 1.0 gramme
and dissolve in
Absolute alcohol 15.0 c.c.
Distilled water 80.0 c.c.
then add
Aniline oil 3.0 c.c.
Shake well and filter before use.
~Haematoxylin~ (Ehrlich).--
1. Weigh out
Haematoxylin 2.0 grammes
and dissolve in
Absolute alcohol 100.0 c.c.
2. Weigh out
Ammonium alum 2.0 grammes
and dissolve in
Distilled water 100.0 c.c.
3. Mix 1 and 2, allow the mixture to stand forty-eight hours, then
filter.
4. Add
Glycerine 85.0 c.c.
Acetic acid, glacial 10.0 c.c.
5. Allow the stain to stand for one month exposed to light; then filter
again ready for use.
~Haematin~ (Mayer's).--
A. Weigh out
Haematin 1.0 gramme
and dissolve in
Alcohol 90 per cent. (warmed to 37 deg. C.) 50 c.c.
B. Weigh out
Potash alum 50 grammes
and dissolve in
Distilled water 100 c.c.
Prepare these two solutions in separate flasks. Take a clean flask of
250 c.c. capacity and insert a large funnel in its neck. Pour the
solutions A and B simultaneously and slowly into the funnel to mix
thoroughly. Store for future use.
NOTE.--If acid haematin is required, introduce glacial acetic
acid (3 c.c.) into the mixing flask before adding the
solutions A and B.
~Alum Carmine~ (Mayer).--
Weigh out
Alum 2.5 grammes
Carmine 1.0 gramme
and place in a glass beaker.
Measure out in a measuring cylinder,
Distilled water 100.0 c.c.
Place the beaker on a sand-bath, add the water in successive small
quantities, and keep the mixture boiling for twenty minutes. Measure the
solution and make up to 100 c.c. by the addition of distilled water.
Filter.
~Lithium Carmine~ (Orth).--
Weigh out
Carmine 2.5 grammes
and dissolve in
Lithium carbonate, cold saturated solution 100.0 c.c.
Filter.
~Picrocarmine.~--
Weigh out
Picrocarmine 2.0 grammes
and dissolve in
Distilled water 100.0 c.c.
BLOOD STAINS
When watery solutions of medicinal methylene blue and water soluble
eosins are mixed a precipitate is formed which is soluble only in
alcohol, and solutions of this precipitate impart a peculiar
reddish-purple colour to chromatin. This compound was first used by
Romanowsky to demonstrate malarial parasites, but various modifications
are now employed for staining blood films generally, and also for
bacteria and protozoa. The best modifications of the original Romanowsky
are those of Jenner and Leishman--Jenner being most suitable for the
histological study of the blood, and Leishman for the demonstration of
protozoa.
~Jenner's Stain.~--
A. Weigh out:
Eosin aqueous yellow 6.0 grammes
Dissolve in
Distilled water (non-alkaline) 250 c.c.
This will make a thick solution.
B. Weigh out:
Methylene blue (medicinally pure) Hoechst 5.0 grammes
Dissolve in
Distilled water (non-alkaline) 250 c.c.
1. Add B to A very slowly, stirring all the time. A viscous precipitate
forms which frequently loses its viscosity when heat is applied. (This
explains the necessity of mixing slowly).
2. Evaporate slowly in a porcelain basin, stirring occasionally, on a
water bath at 55 deg. C. When a paste begins to form scrape and break up
occasionally. (On no account must the paste be allowed to fuse.)
3. Grind the resulting mass into an amorphous powder.
4. Weigh out:
Amorphous powder 0.5 grammes
Dissolve in
Methylic alcohol (Merck's puriss, for analysis) 100 c.c.
Allow time for true solution. (About three days is sufficient.)
METHOD.--
1. Prepare film, dry, but _do not fix_.
2. Flood the unfixed film with the stain, allow it to act for 3 minutes
(the methylic alcohol of the stain fixes the film).
3. Pour off the stain and wash in distilled water until the film
presents a pink colour.
4. Dry and mount.
~Leishman's Stain.~--
_A._ Weigh out:
Methylene blue (medicinal) 1 gramme
Dissolve in
Sodium carbonate, 0.5 per cent. aqueous solution 100 c.c.
Keep at 65 deg. C. for 12 hours in either a hot incubator or a water-bath;
then stand in dark place at room temperature (20 deg. C.) for ten days.
_B._ Weigh out:
Eosin, extra B. A. 0.1 gramme
Dissolve in
Distilled water 100 c.c.
1. Mix the two solutions A and B in equal volumes, and allow the
mixture to stand for 12 hours with occasional stirring.
2. Filter, and collect precipitate on filter paper.
3. Wash precipitate thoroughly with distilled water, and dry.
4. Weigh out 0.15 gramme of the dried precipitate; rub up in a mortar
with 5 c.c. of methylic alcohol (Merck's puriss, for analysis).
Allow undissolved powder to settle, then decant the supernatant fluid to
a clean 100 c.c. measuring cylinder.
5. Add further 5 c.c. alcohol to sediment in mortar and repeat the
process, and so on until all the sediment has been dissolved.
6. Now make up the fluid in the measuring cylinder to 100 c.c. by the
addition of more methylic alcohol.
METHOD.--
1. Prepare film, dry, but _do not fix_.
2. Flood the unfixed film with stain, allow it to act 30 seconds.
3. Add double the volume of distilled water to the stain on the film,
and mix with glass rod or platinum loop.
4. Allow this diluted stain to act five minutes.
5. Wash off with distilled water.
6. Leave some water on film for thirty seconds to intensify the colour
contrasts.
7. Dry and mount.
METHODS OF DEMONSTRATING STRUCTURE OF BACTERIA, ETC.
~To Demonstrate Capsules.~
~1. MacConkey.~--
_Stain._--
Weigh out
Dahlia 0.5 gramme
Methyl green (00 crystals) 1.5 grammes
rub up in a mortar with
Distilled water 100.0 c.c.
Add
Fuchsin, saturated alcoholic solution 10.0 c.c.
and make up to 200 c.c. by the addition of
Distilled water 90.0 c.c.
Filter.
Allow the stain to stand for two weeks before use; keep in a dark place
or in an amber glass bottle. Owing to the unstable character of the
methyl green, this stain deteriorates after about six months.
METHOD.--
1. Prepare and fix film in the usual manner.
2. Flood the cover-slip with the stain and allow it to act for five to
ten minutes.
3. Wash very thoroughly in water; if necessary, direct a powerful stream
of water on the film from a wash-bottle.
4. Dry and mount.
~2. Muir's Method.~--
1. Prepare, dry and fix film in the ordinary manner.
2. Flood the film with carbolic fuchsin, warm until steam
begins to rise. Allow the stain to act for thirty seconds.
3. Wash quickly with methylated spirit.
4. Wash thoroughly with water.
5. Subject the film to the action of the following mordant
for five seconds:
Corrosive sublimate, saturated aqueous solution 2 c.c.
Tannic acid, 20 per cent. aqueous solution 2 c.c.
Potash alum saturated aqueous solution 5 c.c.
6. Wash thoroughly in water.
7. Treat with methylated spirit for about sixty seconds.
(The preparation should now be pale red.)
8. Wash thoroughly in water.
9. Counterstain in methylene blue, aqueous solution thirty
seconds.
10. Wash in water.
11. Dehydrate in alcohol.
12. Clear in xylol and mount in xylol balsam.
~3. Welch's Method.~--
1. Prepare and fix film in the usual manner.
2. Flood the slide with acetic acid 2 per cent.; allow the
acid to remain in contact with the film for two minutes.
This swells up and fixes the capsule and enables it to take
the stain.
3. Blow off the acetic acid by the aid of a pipette.
4. Immerse in aniline gentian violet, five to thirty
seconds.
5. Wash in water.
6. Dry and mount.
~4. Ribbert's Method.~--
_Stain._--
Measure out and mix:
Acetic acid, glacial 12.5 c.c.
Alcohol, absolute 50.0 c.c.
Distilled water 100.0 c.c.
Warm to 36 deg. C. (e. g., in the "hot" incubator) and
saturate with dahlia. Filter.
METHOD.--
1. Prepare and fix films in the usual manner.
2. Cover the film with the stain and allow it to act for one
or two seconds only.
3. Wash thoroughly in water.
4. Dry and mount.
~To Demonstrate Flagella.~
~1. Muir's Modified Pitfield.~--This is the best method and gives the most
reliable results, for not only is the percentage of successful
preparations higher than with any other, but the bacilli and flagella
retain their relative proportions.
(a) ~Mordant.~--
Tannic acid, 10 per cent. aqueous solution 10 c.c.
Corrosive sublimate, saturated aqueous solution 5 c.c.
Alum, saturated aqueous solution 5 c.c.
Carbolic fuchsin (Ziehl) 5 c.c.
Mix thoroughly.
A precipitate forms which must be allowed to settle for a few hours.
Decant off the clear fluid into tubes and centrifugalise thoroughly.
This solution is at its best some four or five days after manufacture;
it keeps for about a couple of weeks, but must be re-centrifugalised
each time, before use.
(b) _Stain._--
Alum, saturated aqueous solution 25 c.c.
Gentian violet, saturated alcoholic solution 5 c.c.
Filter.
This stain must be freshly prepared.
METHOD.--The cultivations employed should be smear agar cultures, twelve
to eighteen hours old if incubated at 37 deg. C, twenty-four to thirty
hours if incubated at 22 deg. C.
1. Remove a very small quantity of the growth by means of the platinum
spatula.
2. Emulsify it with a few cubic centimetres of distilled water in a
watch-glass, by gently moving the spatula to and fro in the water. Do
not rub up the growth on the side of the watch-glass. Some workers
prefer to use tap water, others employ normal saline solution, but
distilled water gives the best emulsion.
3. Spread a thin film of the emulsion on a newly flamed cover-slip,
using no force, but rather _leading_ the drop over the cover-slip with
the platinum loop.
4. Allow the film to dry in the air, properly protected from falling
dust.
5. Fix by passing thrice through the Bunsen flame, holding the
cover-slip whilst doing so by one corner between the finger and thumb.
6. Pour on the film as much of the mordant as the cover-glass will hold.
Grasp the cover-slip with the forceps and hold it, high above the flame,
until steam rises. Allow the steaming mordant to remain in contact with
the film two minutes.
7. Wash well in water and dry carefully.
8. Pour on the film as much of the stain as the cover-glass will hold.
Steam over the flame as before for two minutes.
9. Wash well in water.
10. Dry and mount.
~2. "Pitfield" Original Method.~--
(a) _Mordant._--