The Filar or cobweb Micrometer (Ramsden's micrometer) eyepiece (Fig. 58) consists of an ocular having a fine "fixed" wire stretching horizontally across the field (Fig. 59), a vertical reference wire—fixed—adjusted at right angles to the first; and a fine wire, parallel to the reference wire, which can be moved across the field by the action of a micrometer screw; the drum head is divided into one hundred parts, which successively pass a fixed index as the head is turned. In the lower part of the field is a comb with the intervals between its teeth corresponding to one complete revolution of this screw-head.
As in the previous method, the value of each division of the micrometer scale (i. e., the comb) must first be determined for each optical combination. This is effected as follows:
1. Place the filar micrometer and the stage micrometer in their respective positions.
2. Rotate the screw of the filar micrometer until the movable wire coincides with the fixed one, and the index marks zero on the drum head. (If when the drum head is at zero the two wires do not exactly coincide they must be adjusted by loosening the drum screw and resetting the drum.)
3. Focus the scale of each micrometer accurately, and make the lines on them parallel.
4. Rotate the head of the micrometer screw until the movable line has transversed one division of the stage micrometer. Note the number of complete revolutions (by means of the recording comb) and the fractions of a revolution (by means of scale on the head of the micrometer screw), which are required to measure the 0.01 mm.
5. Make several such estimations and average the results.
6. Note the optical combination employed in this experiment and record it carefully, together with the micrometer value in terms of µ.
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.
[1] Its importance will be realised, however, when it is stated in the words of the late Professor Abbé: "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.
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
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.
Bacteria, etc., are examined microscopically.
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.
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, | } | in 0.5 per cent. aqueous solutions. |
| Quinoleine blue | } | |
| Methylene green | } | |
| 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.
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°) 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 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° 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.
2. Absolute alcohol, Ether, equal parts, for five to thirty minutes (e. g., for blood or 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 hæmoglobin 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.
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.
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, hæmatoxylin) 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 Gram's method. |
| Eosin, | } |
| 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.
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.
Methylene-blue.—
1. Saturated Aqueous Solution.
Weigh out