Chapter VII

SPECIAL METHODS FOR STAINING MICRO-ORGANISMS AND BLOOD.

It is impossible, within the limits of this work, to attempt any adequate description of the modern methods of bacteriological investigation. Some of these are very lengthy and complicated, and require much skill and practice before good results can be relied on. But those who do not desire to make a special study of bacteriology may often require to examine for the presence of organisms in sections, or in various excretions, and it is hoped that they may find the following short description of special methods sufficient for their purpose. For more elaborate work they must consult one of the many excellent textbooks on the subject.

The student should provide himself with the following dyes in powder:--

*Methylene blue.*

*Gentian violet.*

*Methyl violet.*

*Fuchsine.*

*Bismarck brown.*

The following solutions of these dyes are used:--

1. Saturated alcoholic solutions which may be kept in stoppered bottles.

2. One per cent. aqueous solutions. These must be freshly made each time of using.

In filtering either alcoholic or aqueous solutions it is well to moisten the filter paper beforehand with alcohol or water as the case may be.

The following special solutions will also be wanted:--

*Löffler’s methylene blue.*--In this solution a weak solution of caustic potash is employed as a mordant:--

Saturated alcoholic solution of methylene blue 3 volumes.
Caustic potash, aqueous solution 1 : 10,000 10 volumes.

Filter.

This solution is perhaps the most generally useful stain. It colours most bacilli and micrococci, and while rapid in its action rarely overstains. It must be made up fresh on each occasion. It is the best counterstain after staining tubercle bacilli, &c., with fuchsine.

*Ziehl’s carbol-fuchsine.*

_Carbolic acid_ (5 per cent. aqueous solution) 100 volumes.
_Fuchsine_ (saturated alcoholic solution) 11 volumes.

The solution must be filtered immediately before being used.

*Gram’s iodine solution.*--Sections are placed in this solution after being stained with aniline dyes. The iodine in some way fixes the dye in the organisms, so that they are not decolourised along with the rest of the tissues.

It is made thus:--

Iodine 1 grm. 1-1/2 grains.
Iodide of potassium 2 grms. 3 grains.
Distilled water 300 c.c. 1 ounce.

Ten per cent. aqueous solutions of *nitric* and *sulphuric* acids should be prepared and may be kept indefinitely.

The following are the general methods of employing these reagents for the purpose of staining organisms in sections. Special methods are required for special organisms, but one or two only can be given.

*Weigert’s method.*--The sections must be placed in a freshly made one per cent. aqueous solution of methyl violet, gentian violet, fuchsine, &c. The solution may be kept at the temperature of the body in an incubator. The organisms will often stain more readily if the section be passed through a 1 in 2000 solution of corrosive sublimate before putting it into the staining fluid. After staining the section is washed in distilled water and then in methylated spirit until it appears almost decolourised. Some prefer to decolourise the tissues by washing in a half per cent. solution of acetic acid instead of methylated spirit. Practice is required before the correct time for decolourising is accurately estimated. The beginner should float a section rapidly on the slide now and then, put on a cover-glass and examine it under a low power to see if the decoloration has been carried far enough. A contrast stain may then be used, such as picrocarmine, after which the section may be mounted in Farrant’s medium: or a weak solution of another aniline colour may be used as a counter stain, after which the section is clarified in xylol, and mounted in balsam dissolved in xylol.

*Gram’s method.*--Place some aniline oil in a test tube and add ten times its volume of distilled water. Close the end with the thumb and shake very thoroughly. Filter ninety drops into another clean test tube, and add ten drops of a saturated solution of gentian violet or some similar dye. Filter the mixture into a watch glass. Stain sections in it for from three minutes to half-an-hour according to the temperature,--the shorter time for the incubator at 100°, the longer when the sections are stained at the ordinary temperature of the room. Wash in distilled water, and transfer to Gram’s iodine solution until they become black, usually in a few minutes. They are then decolourised in absolute alcohol. This often takes some time. It may be hastened, as Crookshank suggests, by placing the section in clove oil, returning to alcohol, and so on.

*Ehrlich’s modification* of Gram’s method. The contrast stain is here used first.

Stain the section (_e.g._, that of a mitral valve in a case of ulcerative endocarditis), in an alcoholic solution of eosine (1 in 1500). Transfer to a solution of some aniline dye, such as gentian violet, dissolved in aniline oil water, exactly as in Gram’s method. The section floats on the surface and spreads out, owing to the alcohol diffusing out. Stain for about twenty minutes. Wash the section in water, and float out (p. 55) on a glass slide. Allow the water to drain off and add Gram’s iodine slowly from a pipette so as not to disarrange the section. When the section has become quite black pour off the Gram’s solution. Remove all superfluous fluid from the slide with blotting paper, and dry the section by carefully and firmly pressing on it a folded piece of blotting paper. If this is done with care the section need not be injured in the least. Decoloration is effected on the slide with aniline oil, instead of alcohol as in the preceding method. The slide is rocked about so that the colour may be evenly discharged by the aniline. When no more colour comes away, the aniline oil is poured off, the section clarified in xylol, and mounted in Canada balsam.

As soon as the section is decolourised it may be treated with a contrast stain, the most suitable being alcoholic solutions of eosine or Bismarck brown if a blue stain has been employed, or methylene blue if fuchsine has been the first stain used.

The following will be found the most useful stains and contrast stains:--

STAINS. CONTRAST STAINS.

{ Picrocarmine.
Gentian violet. } { Eosine.
Methyl violet. } { Bismarck brown.
Methylene blue. } { Safranine.

Magenta. } Methylene blue.
Fuchsine. }

And _vice versâ_.

Much practice is required in using either of the methods before one can judge accurately how long to leave sections in the staining reagents or decolourising agents, and the beginner must not be discouraged if at first he is unable to obtain good results although he follows the book directions most minutely.

*Ehrlich method for tubercle bacilli.*--Sections are stained for six to twenty-four hours in a one per cent. solution of gentian violet, methyl violet, methyl blue or fuchsine. They will stain more rapidly if the staining fluid be kept in an incubator at the body temperature. They should be removed from the staining fluid, and washed in distilled water, and then transferred (preferably on a glass section lifter) to a ten per cent. solution of nitric acid in distilled water until they are nearly decolourised. They should then be very thoroughly washed in distilled water. They may then be treated with some suitable contrast stain and mounted in Canada balsam.

*Neelsen’s stain for tubercle bacilli.*--Sections are placed in Ziehl’s carbol-fuchsine solution (p. 103) which should be warmed for ten minutes to half-an-hour. They are then decolourised in a solution of sulphuric acid. Twenty-five per cent. is the strength originally recommended, but a ten per cent. solution does equally well and injures the section less. They are then very thoroughly washed in a large quantity of water, and afterwards may be treated with a contrast stain.

*Gibbes’ double stain for tubercle bacilli.*--

(1) Rosaniline hydrochlorate 2 grms. 25 grs.
Methyl blue 1 grm. 12·5 grs.

Triturate in a glass mortar,

(2) Aniline oil 3 c.c. 37·5 grs.
Rectified spirit 15 c.c. 3-1/2 drms.

Dissolve and add slowly to (1).

(3) Lastly add slowly to the mixture
Distilled water 15 c.c. 3-1/2 drms.

Some of the solution is filtered into a watch glass and warmed. The sections are placed in it and left for some hours. They are then washed in methylated spirit till they are sufficiently decolourised, and then rapidly passed through absolute alcohol and oil of cloves and mounted in balsam and xylol. It is a very useful stain for examining the sputum for tubercle bacilli.

In order to stain fluids, such as blood, pus, or sputum, for organisms, a very thin layer should be obtained by placing a little of the fluid between two clean cover-glasses and pressing them together. They are then separated and allowed to dry. The film is fixed by holding the cover-glass in a pair of forceps, and passing it slowly through the flame of a spirit lamp two or three times. Films of pus should be ‘cleared’ after fixing by placing them in a twenty per cent. solution of acetic acid for three minutes.

For clinical purposes it is often necessary to examine urine, fæces or vomited matter for bacilli. Films are prepared in the usual way and allowed to evaporate slowly, and then fixed by passing through the flame, and then washed in distilled water before staining. In the case of vomited matter and fæces this is usually done without difficulty. In the case of urine however it is often difficult to get the urine to evaporate completely. A syrupy layer remains, and if more heat be applied it decomposes and chars, and the products cause precipitation of aniline during subsequent staining processes. This may be partly avoided by gently washing the film in distilled water before staining.

Another plan is to mix the urinary deposit with a little gelatine free from organisms, such as that in unused culture tubes. The gelatin is liquefied by heat, and mixed with the deposit. Films are made from this mixture, and allowed to set, and then thoroughly washed in distilled water. The film is then dried thoroughly, and the cover-glass laid flat with the film uppermost, and a few drops of the staining fluid filtered on to it. After it has been stained sufficiently the stain is drained off, and the slip gently washed. The film may then be stained with some contrast stain in exactly the same way as sections, again washed, dried between folds of blotting paper, and mounted in balsam.

It is sometimes difficult to tell which is the side of the cover-glass which bears the film. This is readily done by holding the glass obliquely so that light from a window is reflected from its surface. The side which is coated appears dull; while the other is smooth and bright.

METHODS OF EXAMINING BLOOD.

In all these methods blood is obtained by pricking the skin of one of the fingers, or the lobule of the ear, preferably the latter. The skin must previously be washed with soap and water or ether, to remove any grease or epithelial scales. The puncture should be made firmly so that blood may escape freely. The finger or ear must not be squeezed. Specimens must be made rapidly before red corpuscles have run into rouleaux. The slides and coverslips employed must be scrupulously clean, or it is impossible to get really good films. They should be cleaned with nitric acid and alcohol according to the directions on page 57.

Fresh specimens should be examined. The coverslip is made just to touch the drop of blood at one edge, so as to transfer a small quantity only, and is at once lowered on to the slide with the aid of a mounted needle. If slide and coverslip be perfectly clean the blood will spread out into a thin film, the corpuscles lying quite flat. If there be any delay, or if the cover-glass be not quite clean the red corpuscles will run into masses and the specimen will be useless for minute examination. Another specimen may be mixed with a little of Ferrier’s solution (p. 129) before mounting. Permanent coverslip films may also be prepared.

Here again the use of absolutely clean coverslips is essential, and the blood must be taken immediately it escapes from the puncture. A little blood is taken on a cover-glass which is held horizontally. Another cover-glass is lowered on to this and by its weight and by capillary attraction, the drop of blood quickly becomes transformed into a thin film. The two covers are separated as soon as the film is formed by rapidly sliding them off one another. This manœuvre requires a little practice and dexterity. The movement of the slips must be in an exactly parallel direction otherwise the coating left will be uneven, just as when two pieces of bread and butter are pulled apart. Even with practice it is difficult to get more than one good film, the lower being usually best. There are four ways of fixing the film.

1. Exposure to *osmic acid vapour*.

The film while still moist is held over the mouth of a bottle containing at least one per cent. solution of osmic acid. In a minute or two the fixation will be complete, and the film becomes of a dirty brown colour. It is then left exposed to the air to get rid of all traces of osmic acid, and may afterwards be stained as described below.

2. Treatment with *saturated aqueous solution of corrosive sublimate* (Muir’s method).

The cover-glass on which the film has been spread, is floated before the latter has time to dry, film downwards on a saturated solution of corrosive sublimate in a watch glass for half an hour. The cover-glass is placed in distilled water and then in alcohol to remove excess of corrosive sublimate, and then stained. A little care is required when washing the film to prevent it sliding bodily off the cover-glass.

3. By drying and passing rapidly through the flame of a Bunsen burner, exactly as in preparing specimens of sputum, &c. (p. 111). This method is handy for ordinary clinical purposes.

4. By keeping the coverslips at a temperature of about 200° F. (Ehrlich’s method).

Ehrlich uses for this purpose a strip of copper about two inches wide and a foot long which is supported on a retort stand in a horizontal position. One end is heated by a Bunsen’s burner beneath. The point in the copper strip at which the temperature is at boiling point is readily ascertained by dropping a little water on. The point at which a drop of water assumes the spherical state indicates a temperature there of 212° F. The coverslips are placed an inch or two further than this point, and kept there at a temperature of about 200° F. for some hours.

STAINING METHODS.

Fresh blood may be stained by mixing with Ferrier’s fuchsine solution:--

Fuchsine 1 grm.
Distilled water 150 c.c.

Dissolve and add

Alcohol (80 per cent.) 50 c.c.
Neutral glycerine 200 c.c.

A spot of this solution is mixed with the blood on a slide by means of a mounted needle, and covered with a clean cover-glass. The red corpuscles are slightly stained, while the nuclei of the white corpuscles are stained a bright crimson, and the “blood plates” a deep pink colour.

Stained preparations may also be obtained by using *Toison’s fluid*, which serves also for diluting the blood in order to determine the exact number of red and white corpuscles present by means of Gowers’ or the Thoma-Zeiss hæmocytometer. It is prepared thus:--

Glycerine 30 c.c. 1 oz.
Sodium sulphate 8 grms. 2 drms.
Sodium chloride 1 grm. 15 grs.
Methyl violet ·25 grm. 4 grs.
Distilled water 160 c.c. 5 oz.

It stains the nuclei and blood plates, but does not alter the shape of the red cells. It requires to be made up fresh occasionally as torulæ are apt to form and multiply in it.

Dried films may be stained with hæmatoxyline, picrocarmine, or any of the general stains. The nuclei of the leucocytes may be stained rapidly in a couple of minutes in a one per cent. solution of methyl violet, washing in water, drying between blotting paper and mounting in balsam. The best method for general purposes is to stain with a saturated aqueous solution of methyl blue for half an hour or longer. Wash in water, and then stain for ten minutes in a half saturated aqueous solution of eosine. In this way the eosinophile granules of the leucocytes and the red corpuscles, are stained by the eosine, while the nuclei of the leucocytes are stained by the methyl blue.

Kanthack and Drysdale recommend that the film should first be stained with a half per cent. solution of eosine in 50 per cent. alcohol, then washed, dried and fixed in the flame, and stained for a short time in Löffler’s solution of methylene blue (p. 104).

These films may be stained for micro-organisms in the way described for cover-glass preparations (p. 112).