 |
7. Repeat this process for each of the different optical combinations and record the results.
To measure an object by this method, simply note the number of revolutions and fractions of a revolution of the screw-head required to traverse such object from edge to edge, and express the result as micra by reference to the recorded values for that particular optical combination.
Microscope Illuminant.—In tropical and subtropical regions diffuse daylight is the best illuminant. In temperate climes however daylight of the desirable quantity is not always available, and recourse must be had to oil lamps, gas lamps—preferably those with incandescent mantles—and electricity; and of these the last is undoubtedly the best. A handy lamp holder which can be manufactured in the laboratory is shown in Fig. 60. It consists of a base board weighted with lead to which is attached the ordinary domestic lamp holder, and behind this is fastened a curved sheet-iron reflector. An obscured metal filament lamp of about 16 candle power gives the most suitable light, and if monochromatic light is needed, the blue grease pencil is streaked over the side of the lamp nearest the microscope; the current is switched on and when the glass bulb is warm, rubbing with a wad of cotton-wool will readily distribute the blue greasy material in an even film over the ground glass.
FOOTNOTES:
[1] Its importance will be realised, however, when it is stated in the words of the late Professor Abbe: "The numerical aperture of a lens determines all its essential qualities; the brightness of the image increases with a given magnification and other things being equal, as the square of the aperture; the resolving and defining powers are directly related to it, the focal depth of differentiation of depths varies inversely as the aperture, and so forth."
[2] Made by Mr. Otto Baumbach, 10, Lime Grove, Manchester.
V. MICROSCOPICAL EXAMINATION OF BACTERIA AND OTHER MICRO-FUNGI.
APPARATUS AND REAGENTS USED IN ORDINARY MICROSCOPICAL EXAMINATION.
The following comprises the essential apparatus and reagents for routine work with which each student should be provided.
1. India-rubber "change-mat" upon which cover-glasses may be rested during the process of staining.
2. Squares of blotting paper about 10 cm., for drying cover-slips and slides.
(The filter paper known as "German lined"—a highly absorbent, closely woven paper, having an even surface and no loose "fluff" to adhere to the specimens—is the most useful for this purpose.)
3. Glass jar filled with 2 per cent. lysol solution for the reception of infected cover-glasses and infected pipettes, etc.
4. A square glazed earthenware box with a loose lining containing 2 per cent. lysol solution for the reception of infected material and used slides. The bottom of the lining is perforated so that when full the lining and its contents can be lifted bodily out of the box, when the disinfectant solution drains away and the slides, etc., can easily be emptied out. The empty lining is then returned to the box with its disinfectant solution (Fig. 61).
5. Bunsen burner provided with "peep-flame" by-pass.
6. Porcelain trough holding five or six hanging-drop slides (Fig. 62).
The best form of hanging-drop slide is a modification of Boettcher's glass ring slide, and is prepared by cementing a circular cell of tin, 13 to 15 mm. diameter, and 1 to 2 mm. in height, to the centre of a 3 by 1 slip by means of Canada balsam. It is often extremely convenient to have two of these cells cemented close together on one slide (Fig. 62, a).
Another form of hanging-drop slide is made in which a circular or oval concavity or "cell" is ground out of the centre of a 3 by 1 slip. These are more expensive, less convenient to work with, and are more easily contaminated by drops of material under examination, and should be carefully avoided.
7. Three aluminium rods (Fig. 63), each about 25 cm. long and carrying a piece of 0.015 gauge platino-iridium wire 7.5 cm. in length. The end of one of the wires is bent round to form an oval loop, of about 1 mm. in its short diameter, and is termed a loop or an oese; the terminal 3 or 4 mm. of another wire is flattened out by hammering it on a smooth iron surface to form a "spatula"; the third is left untouched or is pointed by the aid of a file. These instruments are used for inoculating culture tubes and preparing specimens for microscopical examination.
The method of mounting these wires may be described as follows:
Take a piece of aluminium wire 25 cm. long and about 0.25 cm. in diameter, and drill a fine hole completely through the wire about a centimetre from one end. Sink a straight narrow channel along one side of the wire, in its long axis, from the hole to the nearest end, shallow at first, but gradually becoming deeper.
On the opposite side of the wire make a short cut, 2 mm. in length, leading from the hole in the same direction. [The use of a fine dental drill and small circular saw, worked by a dental motor facilitates the manufacture of these aluminium handled instruments.]
Now pass one end of the platinum wire through the hole, turn up about 2 mm. at right angles and press the short piece into the short cut. Turn the long end of the wire sharply, also at right angles, and sink it into the long channel so that it emerges from about the centre of the cut end of the aluminium wire (Fig. 63). A few sharp taps with a watch maker's hammer will now close in the sides of the two channels over the wire and hold it securely.
8. Two pairs of sharp-pointed spring forceps (10 cm. long), one of which must be kept perfectly clean and reserved for handling clean cover-slips, the other being for use during staining operations.
9. A box of clean 3 by 1 glass slips.
10. A glass capsule with tightly fitting (ground on) glass lid, containing clean cover-slips in absolute alcohol.
11. One of Faber's "grease pencils" (yellow, red, or blue) for writing on glass.
12. A wooden rack (Fig. 65) with twelve drop-bottles (Fig. 66) each 60 c.c. capacity, containing
Aniline water.
Gentian violet, saturated alcoholic solution.
Lugol's (Gram's) iodine.
Absolute alcohol.
Methylene-blue, } Fuchsin, basic, } saturated alcoholic solution.
Neutral red, 1 per cent. aqueous solution.
Leishman's modified Romanowsky stain.
Carbolic acid, 5 per cent. aqueous solution.
Acetic acid, 1 per cent. solution.
Sulphuric acid, 25 per cent. solution.
Xylol.
And two pots with air-tight glass caps (Fig. 67), each provided with a piece of glass rod and filled respectively with Canada balsam dissolved in xylol, and sterile vaseline.
METHODS OF EXAMINATION.
Bacteria, etc., are examined microscopically.
1. In the living state, unstained, or stained. 2. In the "fixed" condition (i. e., fixed, killed, and stained by suitable methods).
The preparation of a specimen from a tube cultivation for examination by these methods may be described as follows:
1. Living, Unstained.—(a) "Fresh" Preparation.—
1. Clean and dry a 3 by 1 glass slip and place it on one of the squares of filter paper. Deposit a drop of water (preferably distilled) or a drop of 1 per cent. solution of caustic potash, on the centre of the slip, by means of the platinum loop.
TECHNIQUE OF OPENING AND CLOSING A CULTURE TUBE.
2. Remove the tube cultivation from its rack or jar with the left hand and ignite the cotton-wool plug by holding it to the flame of the Bunsen burner. Extinguish the flame by blowing on the plug, whilst rotating the tube on its long axis, its mouth directed vertically upward, between the thumb and fingers. (This operation is termed "flaming the plug," and is intended to destroy any micro-organisms that may have become entangled in the loose fibres of the cotton-wool, and which, if not thus destroyed, might fall into the tube when the plug is removed and so accidentally contaminate the cultivation.)
3. Hold the tube at or near its centre between the ends of the thumb and first two fingers of the left hand, and allow the sealed end to rest upon the back of the hand between the thumb and forefinger, the plug pointing to the right. Keep the tube as nearly in the horizontal position as is consistent with safety, to diminish the risk of the accidental entry of organisms (Fig. 68).
4. Take the handle of the loop between the thumb and forefinger of the right hand, holding the instrument in a position similar to that occupied by a pen or a paint-brush, and sterilise the platinum portion by holding it in the flame of a Bunsen burner until it is red hot. Sterilise the adjacent portion of the aluminium handle by passing it rapidly twice or thrice through the flame. After sterilising it, the loop must not be allowed to leave the hand or to touch against anything but the material it is intended to examine, until it is finished with and has been again sterilised.
5. Grasp the cotton-wool plug of the test-tube between the little finger and the palm of the right hand (whilst still holding the loop as directed in step 4), and remove it from the mouth of the tube by a "screwing" motion of the right hand.
6. Introduce the platinum loop into the tube and hold it in this position until satisfied that it is quite cool. (The cooling may be hastened by touching the loop on one of the drops of moisture which are usually to be found condensed on the interior of the glass tube, or by dipping it into the condensation water at the bottom; at the same time care must be taken in the case of cultures on solid media to avoid touching either the medium or the growth.)
7. Remove a small portion of the growth by taking up a drop of liquid, in the case of a fluid culture, in the loop; or by touching the loop on the surface of the growth when the culture is on solid medium; and withdraw the loop from the tube without again touching the medium or the glass sides of the tube.
8. Replace the cotton-wool plug in the mouth of the tube.
9. Replace the tube cultivation in its rack or jar.
10. Mix the contents of the loop thoroughly with the drop of water on the 3 by 1 slide.
11. Again sterilise the loop as directed in step 4, and replace it in its stand.
12. Remove a cover-slip from the glass capsule by means of the cover-slip forceps, rest it for a moment on its edge, on a piece of filter paper to remove the excess of alcohol, then pass it through the flame of the Bunsen burner. This burns off the remainder of the alcohol, and the cover-slip so "flamed" is now clean, dry, and sterile.
13. Lower the cover-slip, still held in the forceps, on to the surface of the drop of fluid on the 3 by 1 slip, carefully and gently, to avoid the inclusion of air bubbles.
14. Examine microscopically (vide infra).
During the microscopical examination, stains and other reagents may be run in under a cover-slip by the simple method of placing a drop of the reagent in contact with one edge of the cover-glass and applying the torn edge of a piece of blotting paper to the opposite side. The reagent may then be observed to flow across the field and come into contact with such of the micro-organisms as lie in its path.
The non-toxic basic dyes most generally employed for the intra-vitam staining of bacteria are
Neutral red, } Quinoleine blue } Methylene green } in 0.5 per cent. aqueous solutions. Vesuvin, }
Negative Stain (Burri).—By this method of demonstration the appearances presented by dark ground illumination (by means of a paraboloid condenser) are closely simulated, since minute particles, bacteria, blood or pus cells etc. stand out as brilliantly white or colourless bodies on a dark grey-brown background.
Reagent required:
Any one of the liquid waterproof black drawing inks (Chin-chin, Pelican, etc.). This is prepared for use as follows:
Measure out and mix:
Liquid black ink, 25 c.c. Tincture of iodine 1 c.c.
Allow the mixture to stand 24 hours, centrifugalise thoroughly, pipette off the supernatant liquid to a clean bottle and then add a crystal of thymol or one drop of formalin as a preservative.
METHOD.—
1. With the sterilised loop deposit one drop of the liquid ink close to one end of a 3 by 1 slide.
2. With the sterilised loop deposit a drop of the fluid culture (or of an emulsion from a solid culture) by the side of the drop of ink (Fig. 69, a); mix the two drops thoroughly by the aid of the loop.
3. Sterilise the loop.
4. Hold the slide firmly on the bench with the thumb and forefinger of the left hand applied to the end nearest the drop of fluid.
5. Take another clean 3 by 1 slide in the right hand and lower its short end obliquely (at an angle of about 60 deg.) transversely on to the mixed ink and culture on the first slide, and allow the fluid to spread across the slide and fill the angle of incidence.
6. Maintaining the original angle, draw the second slide firmly and evenly along the first toward the end farthest from the left hand (Fig. 69, b).
7. Throw the second slide into a pot of disinfectant; allow the first slide to dry in the air.
8. Place a drop of immersion oil on the centre of the film, lower the 1/12-inch objective into the oil and examine microscopically without the intervention of a cover-slip.
(The film of ink may be covered with a long cover-glass and xylol balsam as a permanent preparation.)
(<b) Hanging-drop Preparation.—
1. Smear a layer of sterile vaseline on the upper surface of the ring cell of a hanging-drop slide by means of the glass rod provided with the vaseline bottle, and place the slide on a piece of filter paper.
2. "Flame" a cover-slip and place it on the filter paper by the side of the hanging-drop slide.
3. Place a drop of water on the centre of the cover-slip by means of the platinum loop.
4. Obtain a small quantity of the material it is desired to examine, in the manner detailed above (pages 74-76, steps 2 to 11 must be followed in their entirety and with the strictest exactitude whenever tube contents are being handled), and mix it with the drop of water on the cover-slip.
5. Raise the cover-slip in the points of the forceps and rapidly invert it on to the ring cell of the hanging-drop slide, so that the drop of fluid occupies the centre of the ring. (Carefully avoid contact between the drop of fluid and either the ring cell or the layer of vaseline. Should this happen, the now infected hanging-drop slide and its cover-slip must be dropped into the pot of lysol and a new preparation made.)
6. Press the cover-slip firmly down into the vaseline on to the top of the ring cell. (This spreads out the vaseline into a thin layer, and besides ensuring the adhesion of the cover-slip, seals the cells and so retards evaporation.)
7. Examine microscopically.
The examination of a "fresh" specimen or a "hanging-drop" preparation is directed to the determination of the following data:
1. The nature of the bacteria present—e. g., cocci, bacilli, etc.
2. The purity of the cultivation; this can only be determined when gross morphological differences exist between the organisms present.
3. The presence or absence of spores; when present, spores show their typical refrangibility exceedingly well by this method.
4. The presence or absence of mobility. In a hanging-drop specimen some form of movement can practically always be observed, and its character must be carefully determined by noting the relative positions of adjacent micro-organisms.
(a) Brownian or molecular movement. Minute particles of solid matter (including bacteria), when suspended in a fluid, will always show a vibratory movement affecting the entire field, but never altering the relative positions of the bacteria. (Cocci exhibit this movement, but with the exception of the Micrococcus agilis, the cocci are non-motile.)
(b) Streaming movement. This is due to currents set up in the hanging drop as a result of jarring of the specimen or of evaporation, or to the fact that the cover-slip is not perfectly level, and although the relative positions of the bacteria may vary, still the flowing movement of large numbers of organisms in some one direction will usually be sufficient to demonstrate the nature of this motion.
(c) Locomotive movement, or true motility, is determined by observing some one particular bacillus changing its position in the field independently of, and in a direction contrary to, other organisms present.
When the examination is completed and the specimen finished with, the "fresh specimen"—i. e., the slide with the cover-slip attached—must be dropped into the lysol pot. In the hanging-drop specimen, however, the cover-slip only is infected, and this may be raised from the ring cell by means of forceps and dropped into the disinfectant.
Permanent Staining of the Hanging-drop Specimen.—Occasionally it is necessary to fix and stain a hanging-drop preparation. This may be done as follows:
1. Remove the cover-slip from the cell by the aid of the forceps.
2. If the drop is small, fix it by dropping it face downward, whilst still wet, on to the surface of some Gulland's solution or corrosive sublimate solution (vide page 82) in a watch-glass. If the drop is large, place it face upward on the rubber mat, cover it with an inverted watch-glass, and allow it to dry. Then fix it in the alcohol and ether solution (vide, page 82).
3. Dip the cover-glass into a beaker containing hot water in order to remove some of the vaseline adhering to it.
4. Wash successively in alcohol, xylol, ether, and alcohol, to remove the last traces of grease.
5. Wash in water.
6. Stain, wash, dry, and mount as for an ordinary cover-slip film preparation (vide pages 83-85).
2. Killed, Stained.—In this method three distinct processes are necessary:
"Preparing" and "fixing" the film. Staining. Mounting.
Preparing the Film.—
1. Flame a cover-slip and place it on a piece of filter paper.
2. Place a drop of water on the centre of the cover-slip by means of platinum loop.
3. Obtain a small quantity of the material to be examined upon a sterilised platinum loop (see pages 74-76, steps 2 to 11) and mix it with the drops of water on the cover-slip.
4. Spread the drop of emulsion evenly over the cover-slip in the form of a square film to within 1 mm. of each edge of the cover-slip.
5. Allow it to dry completely in the air.
Fixing.—Fix by passing the cover-slip, held in the fingers, three or four times through the flame of a Bunsen burner.
In some instances (e. g., when the films after staining are intended for micrometric observations) it is almost essential to fix by exposure to a uniform temperature of 115 deg. C., for twenty minutes. This is best done in a carefully regulated hot-air oven.
Fixation may also be effected by immersing in some fixative fluid, such as one of the following:
1. Absolute alcohol, for five to fifteen minutes.
{ equal parts, for five to thirty 2. Absolute alcohol, { minutes (e. g., for blood or Ether, { milk).
3. Osmic acid, 1 per cent. aqueous solution, for thirty seconds.
4. Corrosive sublimate, saturated aqueous solution, for five minutes.
5. Corrosive sublimate (Lang), for five minutes. This solution is prepared by dissolving:
Sodium chloride 0.75 gramme Hydrarg. perchloride 12.00 grammes Acetic acid 5.00 grammes In distilled water 100.00 c.c. Filter.
6. Gulland's solution, for five minutes. This solution is prepared by mixing:
Absolute alcohol 25.0 c.c. Ether 25.0 c.c. Corrosive sublimate, 20 per cent. alcoholic solution 0.4 c.c.
7. Formalin 10 per cent. aqueous solution (= 4 per cent. aqueous solution of formaldehyde since formalin is a 40 per cent. solution of the gas in water).
Either of these methods of fixation coagulates the albuminous material and ensures perfect adhesion of the film to the cover-slip.
Clearing.—Wash the cover-slip thoroughly in running water and proceed with the staining.
If the film has been prepared from broth, liquefied gelatine, or pus or other morbid exudations, saturate the film after fixation with acetic acid 2 per cent. and allow it to act for two minutes.
Wash with alcohol, then let the alcohol remain on the cover-slip for two minutes. (This will "clear" the groundwork and give a much sharper and cleaner film than would otherwise be obtained.)
If the film has been prepared from blood or bloodstained fluid, treat with acetic acid 2 per cent. for two minutes after fixation. Wash with water, dry, and proceed with the staining. (This will remove the haemoglobin and facilitate examination.)
Staining.—
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.
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.—
Tannic acid 1 gramme 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.
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.
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.
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.
NOTE.—If large, thick sections have to be manipulated, or if time is of importance or acids are used during the staining process, it is often advisable to add a trace of Mayer's albumin to the alcohol before floating out the section. If this substance is employed, a sojourn of twenty minutes to half an hour in the "hot" incubator will be found ample to ensure firm adhesion of the section to the slide. The albuminous fluid is prepared as follows:
Mayer's Albumin.—
Weigh out Salicylate of soda 1 gramme and dissolve in Glycerine 50 c.c. Add White of egg 50 c.c.
Mix thoroughly by means of an egg whisk.
Filter into a clean bottle.
As an alternative method paint a thin layer of Schallibaum's solution on the slide with a camel's hair pencil; lay the section carefully on this film and heat gently to fix the section.
Schallibaum's solution:
Clove oil 30 c.c. Collodion 10 c.c.
Keep in a dark blue bottle in a cool place.
Staining Paraffin Sections.—
1. Warm paraffin section over the Bunsen flame to soften (but not to melt) the paraffin, then dissolve out the wax with xylol poured on from a drop bottle.
2. Remove xylol by flushing the section with alcohol.
3. If the tissue was originally "fixed" in a corrosive sublimate solution, the section must now be treated with Lugol's iodine solution for two minutes and subsequently immersed in 90 per cent. alcohol to remove all traces of yellow staining.
4. Wash in water.
5. Stain deeply, if using a single stain, as the subsequent processes decolourise.
6. Wash in water, decolourise if necessary.
7. Flood with several changes of absolute alcohol to dehydrate the section.
8. Clear in xylol. (Oil of cloves is not usually employed, as it decolourises the section.)
9. Mount in xylol balsam.
SPECIAL STAINING METHODS FOR SECTIONS.
Double-staining Carmine and Gram-Weigert.—
1. Prepare the section for staining as above, sections 1 to 3.
2. Stain in lithium carmine (Orth's) or picrocarmine for ten to thirty minutes, in a porcelain staining pot (Fig. 76).
3. Wash in picric acid solution until yellow. At this stage cell nuclei are red, protoplasm is yellow, and bacteria are colourless.
Picric acid solution is prepared by mixing
Picric acid, saturated aqueous solution 40 c.c. Hydrochloric acid 1 c.c. Alcohol (90 per cent.) 160 c.c.
4. Wash in water.
5. Wash in alcohol.
6. Stain in aniline gentian violet.
7. Wash in iodine solution till dark brown or black.
8. Wash in water.
9. Dip in absolute alcohol for a second.
10. Decolourise with aniline oil till no more colour is discharged.
11. Wash with aniline oil, 2 parts, xylol, 1 part.
12. Clear with xylol.
13. Mount in xylol balsam.
Alternative Gram-Weigert Method for Sections.—
1. Fix paraffin section on slide and prepare for staining in the usual manner.
2. Stain in alum carmine for about fifteen minutes.
3. Wash thoroughly in water.
4. Filter aniline gentian violet solution on to the section on the slide and allow to stain about twenty-five minutes.
5. Wash thoroughly in water.
6. Treat with Lugol's iodine until section ceases to become any blacker.
7. Wash thoroughly in water.
8. Treat with a mixture of equal parts of aniline oil and xylol until no more colour comes away.
9. Wash thoroughly with xylol.
10. Decolourise and dehydrate rapidly with absolute alcohol until there remains only a very faint bluish tint.
11. Clear with xylol.
12. Mount in xylol balsam.
(Then fibrin and hyaline tissue are stained deep blue, whilst bacteria which "stain Gram" appear of a deep blue-violet colour.)
Unna-Pappenheim Method.—
Stain.—
Weigh out and mix
Methylene green 0.15 gramme Pyronin 0.25 gramme
and dissolve in
Carbolic acid 0.5 per cent. aqueous solution 78 c.c.
Measure out
Alcohol 2.5 c.c. } Glycerine 20.0 c.c. } and add to the stain.
Method.—
1. Place tissue in the above stain for ten minutes.
2. Differentiate and dehydrate with absolute alcohol.
3. Clear in xylol.
4. Mount in xylol balsam.
To Demonstrate Capsules.—
1. MacConkey's Method.—Stain precisely as for cover-slip films (vide page 100).
2. Friedlaender's Method.—
Stain.—
Gentian violet, saturated alcoholic solution 50 c.c. Acetic acid, glacial 10 c.c. Distilled water 100 c.c.
METHOD.—
1. Prepare the sections for staining, secundum artem.
2. Stain sections in the warm (e. g., in the hot incubator) for twenty-four hours.
3. Wash with water.
4. Decolourise lightly with acetic acid, 1 per cent.
5. Dehydrate rapidly with absolute alcohol.
6. Clear with xylol.
7. Mount in xylol balsam.
To Demonstrate Acid-fast Bacilli.—
1. Prepare the sections for staining in the usual way.
2. Stain with haematin solution ten to twenty seconds, to obtain a pure nuclear stain; then wash in water.
3. Stain with carbolic fuchsin twenty to thirty minutes at 47 deg. C.; then wash in water.
4. Treat with aniline hydrochlorate, 2 per cent. aqueous solution, for two to five seconds.
5. Decolourise in 75 per cent. alcohol till section appears free from stain—fifteen to thirty minutes.
6. Dehydrate with absolute alcohol.
7. Clear very rapidly with xylol.
8. Mount in xylol balsam.
To Demonstrate Spirochaetes in Tissues.
Piridin Method (Levaditi).—
1. Cut slices of tissue 1 mm. thick.
2. Fix in 10 per cent. formalin solution for twenty-four hours.
3. Wash in water for one hour.
4. Place in 96 per cent. alcohol for twenty-four hours.
5. Measure into a dark green or amber bottle 100 c.c. silver nitrate solution 1 per cent., and 10 grammes pyridin puriss. Transfer slices of tissue to this. Stopper and keep at room temperature three hours, then in thermostat at 50 deg. C. for four to six hours.
6. Wash quickly in 10 per cent. pyridin solution.
7. Reduce silver by transferring slices of tissue to following solution for forty-eight hours.
Pyrogallic acid 4 grammes Acetone 10 c.c. Pyridin puriss 15 grammes Distilled water 100 c.c.
8. Wash well in water.
Take through alcohols of increasing strength up to absolute, keeping in each strength for twenty-four hours.
9. Clear, embed, cut very thin sections, mount, remove paraffin, again clear and mount in xylol balsam.
The spirochaetes if present are black and show up against the pale yellow color of the background.
Weak carbol fuchsin, neutral red or toluidin blue can also be used to stain the background if desired, after the removal of the paraffin in step 9.
To Demonstrate Protozoa in Sections (Leishman).—
Reagents required:
Leishman's Polychrome stain. Acetic acid 1 in 1500 aqueous solution. Caustic soda 1 in 7000 aqueous solution. Distilled water.
1. Mount section, remove paraffin and take into distilled water as usual (vide page 121).
2. Drain off the excess of water.
3. Cover the section with diluted Leishman (1 part stain, 2 parts distilled water) and allow to act for five to ten minutes (until tissue appears a deep blue).
4. Decolourise with acetic acid solution until only the nuclei appear blue (examine the section wet, with low power objective).
5. If the eosin colour is too well marked treat with the caustic soda solution until the desired tint is obtained (as seen with the 1/6-inch objective).
6. Wash with distilled water.
7. Rapidly dehydrate with alcohol.
8. Clear with xylol.
9. Mount in xylol balsam.
VIII. CLASSIFICATION OF FUNGI.
For practical purposes FUNGI may be divided into:
1. Hymenomycetes (including the mushrooms, etc.). 2. Hyphomycetes (moulds). 3. Blastomycetes (yeasts and torulae). 4. Schizomycetes (bacteria).
NOTE.—Formerly myxomycetes were included in the fungi; they are now recognized as belonging to the animal kingdom, and are termed "mycetozoa."
MORPHOLOGY OF THE HYPHOMYCETES.
At the commencement of his studies, the attention of the student is directed to the various non-pathogenic moulds and yeasts, not only that he may gain the necessary technique whilst handling cultivations of harmless organisms, but also because these very species are amongst the commonest of those that may accidentally contaminate his future preparations.
The hyphomycetes are composed of a mycelium of short jointed rods or "hyphae" springing from an axis or germinal tube which develops from the spore. Hyphae are—
(a) Nutritive or submerged.
(b) Reproductive or aerial.
The protoplasm of these cells contains granules, pigment, oil globules, and sometimes crystals of calcium oxalate.
Reproduction.—Apical spore formation—asexual; zoospores—sexual.
Mucorinae.—Mucor (Fig. 77).—Note the branching filaments—"mycelium" (a), "hyphae" (b).
Note the asexual reproduction.
1. A filament grows upward. At its apex a septum forms, then a globular swelling appears—"sporagium" (d). This possesses a definite membrane.
2. From the septum grows a club-shaped mass of protoplasm—"columella" (c).
3. The rest of the contained protoplasm breaks up into "swarm spores" (e).
Finally the membrane ruptures and spores escape.
Perisporaceae.—Aspergillus (Fig. 78).—Note the branching filaments—"mycelium" (a).
Note the asexual reproduction.
1. A filament (b) grows upward, its termination becomes clubbed; on the clubbed extremity flask-shaped cells appear—"sterigmata" (c).
2. At free end of each sterigma is formed an oval body—a spore or "gonidium" (d), which, when ripe, is thrown off from the sterigma. Two or more gonidia may be supported upon each sterigma.
Penicillium (Fig. 79).—Note the branching filaments—"mycelium" (a) (frequently containing globules).
Note the asexual reproduction.
1. A filament grows upward—"goniodophore" (b)—and its apex divides up into several branches—"basidia" (c).
2. At the apex of each basidium a flask-shaped cell, "sterigma" (d), appears.
3. At the apex of each sterigma appears a row of oval cells—"spores" or "conidia" (e). These, when ripe, are cast off from the sterigmata.
Ascomycetae.—Oidium (Fig. 80).—(This family is perhaps as nearly related to the blastomycetes as it is to the hyphomycetes.)
Note the branching filaments—"pseudomycelium" (a). Here and there filaments are broken up at their ends into oval or rod-shaped segments, "oidia," and behave as spores.
Note the asexual reproduction. From the pseudomycelium arise true hyphae (b), each of which in turn ends in a chain of spores (c).
MORPHOLOGY OF THE BLASTOMYCETES.
The blastomycetes are composed of spherical or oval cells (8 to 9.5 mu in diameter), which, when rapidly multiplying by budding, may form a spurious mycelium. A thin cell-wall encloses the granular protoplasm, in which vacuoles and sometimes a nucleus may be noted. This latter is best seen when stained with haematoxylin (see page 105).
During their growth and multiplication the blastomycetes split up solutions containing sugar into alcohol and CO_{2}.
Saccharomyces (Fig. 81).—Note the round or oval cells of granular protoplasm (a) containing solid particles and vacuoles (c), and surrounded by a definite envelope.
Reproduction.—Budding; ascospores—asexual.
Note the asexual reproduction.
1. "Gemmation"—that is, the budding out of daughter cells (b) from various parts of the gradually enlarging mother cell. These are eventually cast off and in turn become mother cells and form fresh groups of buds.
2. Spore formation—"ascospores" (e). These are formed at definite temperatures and within well-defined periods; e. g., Saccharomyces cerevisiae, thirty hours at 25 deg. to 37 deg. C., or ten days at 12 deg. C.
Torulae (Fig. 82).—Torulae, whilst resembling yeasts in almost every other respect, never form endo-spores. Note the elongated, sausage-shaped cells (a) the larger oval cells (b) and the globular cells (c) the former two often interlacing and growing as a film.
Note the absence of ascospore formation.
IX. SCHIZOMYCETES.
Classification and Morphology.—Bacteria are often classified, in general terms, according to their life functions, into—
Saprogenic, or putrefactive bacteria; Zymogenic, or fermentative bacteria; Pathogenic, or disease-producing bacteria;
or according to their food requirements into—
Prototrophic, requiring no organic food (e. g., nitrifying bacteria); Metatrophic, requiring organic food (e. g., saprophytes and facultative parasites); Paratrophic, requiring living food (obligate parasites);
or according to their metabolic products into—
Chromogenic, or pigment-producing bacteria; Photogenic, or light-producing bacteria; Aerogenic, or gas-producing bacteria;
and so on.
Such broad groupings as these have, however, but little practical value when applied to the systematic study of the fission fungi.
On the other hand, no really scientific classification of the schizomycetes has yet been drawn up, and the varying morphological appearances of the members of the family are still utilised as a basis for classification, as under—
1. Cocci. (Fig. 83).—Rounded or oval cells, subdivided according to the arrangement of the individuals after fission, into—
Diplococci and Streptococci, where division takes place in one plane only, and the individuals remain attached (a) in pairs or (b) in chains.
Tetrads, Merismopedia, or Pediococci, where division takes place alternately in two planes at right angles to each other, and the individuals remain attached in flat tablets of four, or its multiples.
Sarcinae, where division takes place in three planes successively, and the individuals remain attached in cubical packets of eight and its multiples.
Micrococci or Staphylococci, where division takes place in three planes, but with no definite sequence; consequently the individuals remain attached in pairs, short chains, plates of four, cubical packets of eight, and irregular masses containing numerous cocci. |
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