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.
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°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 hæmatoxylin (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° 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° C. For all ordinary purposes 54°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
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. Friedländer'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 hæmatin 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°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 Spirochætes 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° 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 spirochætes 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:
2. Hyphomycetes (moulds).
3. Blastomycetes (yeasts and torulæ).
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 "hyphæ" springing from an axis or germinal tube which develops from the spore. Hyphæ 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.
Mucorinæ.—Mucor (Fig. 77).—Note the branching filaments—"mycelium" (a), "hyphæ" (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.
Perisporaceæ.—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.
Ascomycetæ.—Oïdium (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, "oïdia," and behave as spores.
Note the asexual reproduction. From the pseudomycelium arise true hyphæ (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 9.5µ 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 hæmatoxylin (see page 105).
During their growth and multiplication the blastomycetes split up solutions containing sugar into alcohol and CO2.
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.