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THE ELEMENTS

OF

BACTERIOLOGICAL TECHNIQUE

A LABORATORY GUIDE FOR MEDICAL, DENTAL, AND TECHNICAL STUDENTS

BY

J. W. H. EYRE, M.D., M.S., F.R.S. (Edin.)

Director of the Bacteriological Department of Guy's Hospital, London, and Lecturer on Bacteriology in the Medical and Dental Schools; formerly Lecturer on Bacteriology at Charing Cross Hospital Medical School, and Bacteriologist to Charing Cross Hospital; sometime Hunterian Professor, Royal College of Surgeons, England

SECOND EDITION REWRITTEN AND ENLARGED

PHILADELPHIA AND LONDON
W. B. SAUNDERS COMPANY
1913

Copyright, 1902, by W. B. Saunders and Company Revised, entirely
reset, reprinted, and recopyrighted July, 1913

Copyright, 1913, by W. B. Saunders Company

Registered at Stationers' Hall, London, England

PRINTED IN AMERICA

PRESS OF
W. B. SAUNDERS COMPANY
PHILADELPHIA

TO THE MEMORY OF

JOHN WICHENFORD WASHBOURN, C.M.G., M.D., F.R.C.P.

Physician to Guy's Hospital and Lecturer on Bacteriology in the
Medical School, and Physician to the London Fever Hospital

MY TEACHER, FRIEND, AND CO-WORKER

PREFACE TO THE SECOND EDITION

Bacteriology is essentially a practical study, and even the elements of its technique can only be taught by personal instruction in the laboratory. This is a self-evident proposition that needs no emphasis, yet I venture to believe that the former collection of tried and proved methods has already been of some utility, not only to the student in the absence of his teacher, but also to isolated workers in laboratories far removed from centres of instruction, reminding them of forgotten details in methods already acquired. If this assumption is based on fact no further apology is needed for the present revised edition in which the changes are chiefly in the nature of additions—rendered necessary by the introduction of new methods during recent years.

I take this opportunity of expressing my deep sense of obligation to my confrère in the Physiological Department of our medical school—Mr. J. H. Ryffel, B. C., B. Sc.—who has revised those pages dealing with the analysis of the metabolic products of bacterial life; to successive colleagues in the Bacteriological Department of Guy's Hospital, for their ready co-operation in working out or in testing new methods; and finally to my Chief Laboratory Assistant, Mr. J. C. Turner whose assistance and experience have been of the utmost value to me in the preparation of this volume. I have also to thank Mrs. Constant Ponder for many of the new line drawings and for redrawing a number of the original cuts.

PREFACE TO THE FIRST EDITION

In the following pages I have endeavoured to arrange briefly and concisely the various methods at present in use for the study of bacteria, and the elucidation of such points in their life-histories as are debatable or still undetermined.

Of these methods, some are new, others are not; but all are reliable, only such having been included as are capable of giving satisfactory results even in the hands of beginners. In fact, the bulk of the matter is simply an elaboration of the typewritten notes distributed to some of my laboratory classes in practical and applied bacteriology; consequently an attempt has been made to present the elements of bacteriological technique in their logical sequence.

I make no apology for the space devoted to illustrations, nearly all of which have been prepared especially for this volume; for a picture, if good, possesses a higher educational value and conveys a more accurate impression than a page of print; and even sketches of apparatus serve a distinct purpose in suggesting to the student those alterations and modifications which may be rendered necessary or advisable by the character of his laboratory equipment.

The excellent and appropriate terminology introduced by Chester in his recent work on "Determinative Bacteriology" I have adopted in its entirety, for I consider it only needs to be used to convince one of its extreme utility, whilst its inclusion in an elementary manual is calculated to induce in the student habits of accurate observation and concise description.

With the exception of Section XVII—"Outlines for the Study of Pathogenic Bacteria"—introduced with the idea of completing the volume from the point of view of the medical and dental student, the work has been arranged to allow of its use as a laboratory guide by the technical student generally, whether of brewing, dairying, or agriculture.

So alive am I to its many inperfections that it appears almost superfluous to state that the book is in no sense intended as a rival to the many and excellent manuals of bacteriology at present in use, but aims only at supplementing the usually scanty details of technique, and at instructing the student how to fit up and adapt apparatus for his daily work, and how to carry out thoroughly and systematically the various bacterioscopical analyses that are daily demanded of the bacteriologist by the hygienist.

Finally, it is with much pleasure that I acknowledge the valuable assistance received from my late assistant, Mr. J. B. Gall, A. I. C., in the preparation of the section dealing with the chemical products of bacterial life, and which has been based upon the work of Lehmann.

[Pg ix]

CONTENTS

Page

I. Laboratory Regulations 1


II. Glass Apparatus in Common Use 3

The Selection, Preparation, and Care of
Glassware, 8—Cleaning of Glass
Apparatus, 18—Plugging Test-tubes and
Flasks, 24.


III. Methods of Sterilisation 26

Sterilising Agents, 26—Methods of
Application, 27—Electric Signal Timing
Clock, 38.


IV. The Microscope 49

Essentials, 49—Accessories, 57—Methods
of Micrometry, 61.


V. Microscopical Examination of Bacteria and Other
Micro-fungi 69

Apparatus and Reagents used in Ordinary
Microscopical Examination, 69—Methods of
Examination, 74.


VI. Staining Methods 90

Bacteria Stains, 90—Contrast Stains,
93—Tissue Stains, 95—Blood Stains,
97—Methods of Demonstrating Structure of
Bacteria, 99—Differential Methods of
Staining, 108.


VII. Methods of Demonstrating Bacteria in Tissues 114

Freezing Method, 115—Paraffin Method,
117—Special Staining Methods for
Sections, 121.


VIII. Classification of Fungi 126

Morphology of the Hyphomycetes,
126—Morphology of the Blastomycetes,
129.


IX. Schizomycetes 131

Anatomy, 134—Physiology,
136—Biochemistry, 144.


X. Nutrient Media 146

Meat Extract, 148—Standardisation of
Media, 154—The Filtration of Media,
156—Storing Media in Bulk, 159—Tubing
Nutrient Media, 160.


[Pg x]XI. Ordinary or Stock Culture Media 163


XII. Special Media 182


XIII. Incubators 216


XIV. Methods of Cultivation 221

Aerobic, 222—Anaerobic, 236.


XV. Methods of Isolation 248


XVI. Methods of Identification and Study 259

Scheme of Study, 259—Macroscopical
Examination of Cultivations,
261—Microscopical Methods,
272—Biochemical Methods, 276—Physical
Methods, 295—Inoculation Methods,
315—Immunisation, 321—Active
Immunisation, 322—The Preparation of
Hæmolytic Serum, 327—The Titration of
Hæmolytic Serum, 328—Storage of
Hæmolysin, 331.


XVII. Experimental Inoculation of Animals 332

Selection and Care of Animals,
335 —Methods of Inoculation, 352.


XVIII. The Study of Experimental Infections During Life 370

General Observations, 371—Blood
Examinations, 373—Serological
Investigations, 378—Agglutinin,
381—Opsonin, 387—Immune Body, 393.


XIX. Post-mortem Examination of Experimental Animals 396


XX. The Study of the Pathogenic Bacteria 408


XXI. Bacteriological Analyses 415

Bacteriological Examination of Water,
416—Examination of Milk, 441—Ice Cream,
457—Examination of Cream and Butter,
457—Examination of Unsound Meats,
460—Examination of Oysters and Other
Shellfish, 463—Examination of Sewage and
Sewage Effluents, 466—Examination of
Air, 468—Examination of Soil,
470—Testing Filters, 478—Testing of
Disinfectants, 480.


Appendix 492


Index 505

[Pg 1]

BACTERIOLOGICAL TECHNIQUE.

I. LABORATORY REGULATIONS.

The following regulations are laid down for observance in the Bacteriological Laboratories under the direction of the author. Similar regulations should be enforced in all laboratories where pathogenic bacteria are studied.

Guy's Hospital.

[Pg 3]

II. GLASS APPARATUS IN COMMON USE.

The equipment of the bacteriological laboratory, so far as the glass apparatus is concerned, differs but little from that of a chemical laboratory, and the cleanliness of the apparatus is equally important. The glassware comprised in the following list, in addition to being clean, must be stored in a sterile or germ-free condition.

Test-tubes.—It is convenient to keep several sizes of test-tubes in stock, to meet special requirements, viz.:

1. 18 × 1.5 cm., to contain media for ordinary tube cultivations.

2. 18 × 1.3 cm., to contain media used for pouring plate cultivations, and also for holding sterile "swabs."

3. 18 × 2 cm., to contain wedges of potato, beetroot, or other vegetable media.

4. 13 × 1.5 cm., to contain inspissated blood-serum.

The tubes should be made from the best German potash glass, "blue-lined," stout and heavy, with the edge of the mouth of the tube slightly turned over, but not to such an extent as to form a definite rim. (Cost about $1.50, or 6 shillings per gross.) Such tubes are expensive it is true, but they are sufficiently stout to resist rough handling, do not usually break if accidentally allowed to drop (a point of some moment when dealing with cultures of pathogenic bacteria), can be cleaned, sterilised, and used over and over again, and by their length of life fully justify their initial expense.

A point be noted is that the manufacturers rarely turn out such tubes as these absolutely uniform in[Pg 4] calibre, and a batch of 18 by 1.5 cm. tubes usually contains such extreme sizes as 18 by 2 cm. and 18 by 1.3 cm. Consequently, if a set of standard tubes is kept for comparison or callipers are used each new supply of so-called 18 by 1.5 cm. tubes may be easily sorted out into these three sizes, and so simplify ordering.

5. 5 × 0.7 cm., for use in the inverted position inside the tubes containing carbohydrate media, as gas-collecting tubes.

These tubes, "unrimmed," may be of common thin glass as less than two per cent. are fit for use a second time.

Fig. 1.—Bohemian flask.

Fig. 2.—Pear-shaped flask.

Fig. 3.—Erlenmeyer flask (narrow neck).

Bohemian Flasks (Fig. 1).—These are the ordinary flasks of the chemical laboratory. A good variety, ranging in capacity from 250 to 3000 c.c., should be kept on hand. A modified form, known as the "pear-shaped" (Fig. 2), is preferable for the smaller sizes—i. e., 250 and 500 c.c.

Erlenmeyer's Flasks (Fig. 3).—Erlenmeyer's flasks of 75, 100, and 250 c.c. capacity are extremely useful. For use as culture flasks care should be taken to select only such as have a narrow neck of about 2 cm. in length.

Kolle's Culture Flasks (Fig. 4).—These thin, flat flasks (to contain agar or gelatine, which is allowed to solidify in a layer on one side) are extremely useful[Pg 5] on account of the large nutrient surface available for growth. A surface cultivation in one of these will yield as much growth as ten or twelve "oblique" tube cultures. The wide mouth, however, is a disadvantage, and for many purposes thin, flat culture bottles known as Roux's bottles (Fig. 5) are to be preferred.

Fig. 4.—Kolle's culture flask.

Fig. 5.—Roux's culture bottle.

Fig. 6.—Guy's culture bottle.

Fig. 7.—Filter flask.

An even more convenient pattern is that used in the author's laboratory (Fig. 6), as owing to the greater depth of medium which it is possible to obtain in these flasks an exceedingly luxuriant growth is possible; the narrow neck reduces the chance of accidental contamination to a minimum and the general shape permits the flasks to be stacked one upon the other.[Pg 6]

Filter Flasks or Kitasato's Serum Flasks (Fig. 7).—Various sizes, from 250 to 2000 c.c. capacity. These must be of stout glass, to resist the pressure to which they are subjected, but at the same time must be thoroughly well annealed, in order to withstand the temperature necessary for sterilisation.

All flasks should be either of Jena glass or the almost equally well-known Resistance or R glass, the extra initial expense being justified by the comparative immunity of the glass from breakage.

Petri's Dishes or "Plates" (Fig. 8, a).—These have now completely replaced the rectangular sheets of glass introduced by Koch for the plate method of cultivation. Each "plate" consists of a pair of circular discs of glass with sharply upturned edges, thus forming shallow dishes, one of slightly greater diameter than the other, and so, when inverted, forming a cover or cap for the smaller. Plates having an outside diameter of 10 cm. and a height of 1.5 cm. are the most generally useful. A batch of eighteen such plates is sterilised and stored in a cylindrical copper box (30 cm. high by 12 cm. diameter) provided with a "pull-off" lid. Inside each box is a copper stirrup with a circular bottom, upon which the plates rest, and by means of which each can be raised in turn to the mouth of the box (Fig. 9) for removal.

Capsules (Fig. 8, b and c).—These are Petri's dishes of smaller diameter but greater depth than those termed plates. Two sizes will be found especially useful—viz., 4 cm. diameter by 2 cm. high, capacity about 14 c.c.; and 5 cm. diameter by 2 cm. high, capacity about 25 c.c. These are stored in copper cylinders of similar construction to those used for plates, but measuring 20 by 6 cm. and 20 by 7 cm., respectively.

Graduated Pipettes.—Several varieties of these are required, viz.:

1. Pipettes of 1 c.c. capacity graduated in 0.1 c.c.[Pg 7]

2. Pipettes of 1 c.c. capacity graduated in 0.01 c.c. (Fig. 10, a).

Fig. 8.—Petri dish (a), and capsules (b, c).

Fig. 9.—Plate box with stirrup.

3. Pipettes of 10 c.c. capacity graduated in 0.1 c.c. (Fig. 10, b).

These should be about 30 cm. in length (1 and 2 of fairly narrow bore), graduated to the extreme point, and having at least a 10 cm. length of clear space between the first graduation and the upper end; the open mouth should be plugged with cotton-wool. Each variety should be sterilised and stored in a separate cylindrical copper case some 36 by 6 cm., with "pull-off" lid, upon which is stamped, in plain figures, the capacity of the contained pipettes.

Fig. 10.—Measuring pipettes, a and b.

The laboratory should also be provided with a complete set of "Standard" graduated pipettes, each pipette in the set being stamped and authenticated by a certificate from one of the recognised Physical Measurement Laboratories, such as Charlottenburg.[Pg 8] These instruments are expensive and should be reserved solely for standardising the pipettes in ordinary use, and for calibrating small pipettes manufactured in the laboratory. Such a set should comprise, at least, pipettes delivering 10 c.c., 5 c.c., 2.5 c.c., 2 c.c., 1 c.c., 0.5 c.c., 0.25 c.c., 0.2 c.c., 0.1 c.c., 0.05 c.c., and 0.01 c.c., respectively.

In the immediately following sections are described small pieces of glass apparatus which should be prepared in the laboratory from glass tubing of various sizes. In their preparation three articles are essential; first a three-square hard-steel file or preferably a glass-worker's knife of hard Thuringian steel for cutting glass tubes etc.; next a blowpipe flame, for although much can be done with the ordinary Bunsen burner, a blowpipe flame makes for rapid work; and lastly a bat's-wing burner.

Fig. 11.—Glass-cutting knife. a. handle. b. double edged blade. c. shaft. d. locking nut. e. spanner for nut.

1. The glass-cutting knife. This article is sold in two forms, a bench knife (Fig. 11) and a pocket knife. The former is provided with a blade some 8 cm. in length and having two cutting edges. The cutting edge when examined in a strong light is seen to be composed of small closely set teeth, similar to those in a saw. The knife should be kept sharp by frequent stroppings on a sandstone hone. The pocket form, about 6-cm. long[Pg 9] over all, consists of a small spring blade with one cutting edge mounted in scales like an ordinary pocket knife.

2. For real convenience of work the blowpipe should be mounted on a special table connected up with cylindrical bellows operated by a pedal. That figured (Fig. 12) is made by mounting a teak top 60 cm. square upon the uprights of an enclosed double-action concertina bellows (Enfer's) and provided with a Fletcher's Universal gas blowpipe.

3. An ordinary bat's-wing gas-burner mounted at the far corner of the table top is invaluable in the preparation of tubular apparatus with sharp curves, and for coating newly-made glass apparatus with a layer of soot to prevent too rapid cooling, and its usually associated result—cracking.

Fig. 12.—Glass blower's table with Enfer's foot bellows.

6. Sedimentation tubes 5×0.5 cm., for sedimentation reactions, etc., and for containing small quantities of fluid to be centrifugalised in the hæmatocrit. These are made by taking 14-cm. lengths of stout glass tubing of the requisite diameter and heating the centre in the Bunsen or blowpipe flame. When the central portion is quite soft draw the ends quickly apart and then round off the pointed ends of the two test-tubes thus[Pg 10] formed. With the glass-cutting knife cut off whatever may be necessary from the open ends to make the tubes the required length.

A rectangular block of "plasticine" (modelling clay) into which the conical ends can be thrust makes a very convenient stand for these small tubes.

Capillary Pipettes or Pasteur's Pipettes (Fig. 13 a).—These little instruments are invaluable, and a goodly supply should be kept on hand. They are prepared from soft-glass tubing of various-sized calibre (the most generally useful size being 8 mm. diameter) in the following manner: Hold a 10 cm. length of glass tube by each end, and whilst rotating it heat the central portion in the Bunsen flame or the blowpipe blast-flame until the glass is red hot and soft. Now remove it from the flame and steadily pull the ends apart, so drawing the heated portion out into a roomy capillary tube; break the capillary portion at its centre, seal the broken ends in the flame, and round off the edges of the open end of each pipette. A loose plug of cotton-wool in the open mouth completes the capillary pipette. After a number have been prepared, they are sterilised and stored in batches, either in metal cases similar to those used for the graduated pipettes or in large-sized test-tubes—sealed ends downward and plugged ends toward the mouth of the case.

Fig. 13.—Capillary pipettes. a, b, c.

The filling and emptying of the capillary pipette is most satisfactorily accomplished by slipping a small rubber teat (similar to that on a baby's feeding bottle but not perforated) on the upper end, after cutting or[Pg 11] snapping off the sealed point of the capillary portion. If pressure is now exerted upon the elastic bulb by a finger and thumb whilst the capillary end is below the surface of the fluid to be taken up, some of the contained air will be driven out, and subsequent relaxation of that pressure (resulting in the formation of a partial vacuum) will cause the fluid to ascend the capillary tube. Subsequent compression of the bulb will naturally result in the complete expulsion of the fluid from the pipette (Fig. 14).

Fig. 14.—Filling the capillary teat-pipette.

A modification of this pipette, in which a constriction or short length of capillary tube is introduced just below the plugged mouth (Fig. 13, b), will also be found extremely useful in the collection and storage of morbid exudations.

A third form, where the capillary portion is about 4 or 5 cm. long and only forms a small fraction of the entire length of the pipette (Fig. 13, c), will also be found useful.

"Blood" Pipettes (Fig 15).—Special pipettes for the collection of fairly large quantities of blood (as suggested by Pakes) should also be prepared. These are made from soft glass tubing of 1 cm. bore, in a similar manner to the Pasteur pipettes, except that[Pg 12] the point of the blowpipe flame must be used in order to obtain the sharp shoulder at either end of the central bulb. The terminal tubes must retain a diameter of at least 1 mm., in order to avoid capillary action during the collection of the fluid.

Fig. 15.—Blood pipettes and hair-lip pin in a test-tube.

Fig. 16.—Blood-pipette in metal thermometer case.

For sterilisation and storage each pipette is placed inside a test-tube, resting on a wad of cotton-wool, and the tube plugged in the ordinary manner. As these tubes are used almost exclusively for blood work, it is usual to place a lance-headed hare-lip pin or a No. 9 flat Hagedorn needle inside the tube so that the entire outfit may be sterilised at one time.

For the collection of small quantities of blood for agglutination reactions and the like, many prefer a short straight piece of narrow glass tubing drawn out at either extremity to almost capillary dimensions. Such pipettes, about 8 cm. in length over all, are most[Pg 13] conveniently sterilized in ordinary metal thermometer cases (Fig. 16).

Graduated Capillary Pipettes (Fig. 17).—These should also be made in the laboratory—from manometer tubing—of simple, convenient shape, and graduated by the aid of "standard" pipettes (in hundredths) to contain such quantities as 10, 50, and 90 c. mm., and carefully marked with a writing diamond. These, previously sterilised in large test-tubes, will be found extremely useful in preparing accurate percentage solutions, when only minute quantities of fluid are available.

Fig. 17.—Capillary graduated pipettes.

Automatic ("Throttle") Pipettes.—These ingenious pipettes, introduced by Wright, can easily be calibrated in the laboratory and are exceedingly useful for graduating small pipettes, for measuring small quantities of fluids, in preparing dilutions of serum for agglutination reactions, etc. They are usually made from the Capillary Pasteur pipettes (Fig. 13, a). The following description of the manufacture of a 5 c. mm. pipette will serve to show how the small automatic pipettes are calibrated.

1. Select a pipette the capillary portion of which is fairly roomy in bore and possesses regular even walls, and remove the cotton-wool plug from the open end.

2. Heat the capillary portion near the free extremity in the by-pass flame of the bunsen burner and draw it out into a very fine hair-like tube and break this across. This hair-like extremity will permit the passage of air but is too fine for metallic mercury to pass.

3. From a standard graduated pipette deliver 5 c. mm. clean mercury into the upper wide portion of the pipette.[Pg 14]

4. Adjust a rubber teat to the pipette and by pressure on the bulb gradually drive the mercury in an unbroken column down the capillary tube until it is stopped by the filiform extremity.

5. Cut off the capillary tube exactly at the upper level of the column of mercury, invert it and allow the mercury to run out.

6. Snap off the remainder of the capillary tube from the broad upper portion of the pipette which is now destined to form the covering tube or air chamber, or what we may term the "barrel." This barrel now has the lower end in the form of a truncated cone, the upper end being cut square. Remove the teat.

7. Introduce the capillary tube into this barrel with the filiform extremity uppermost, and the square cut end projecting about 0.5 cm. beyond the tapering end of the barrel.

Fig. 18.—Throttle pipette—small capacity.

8. Drop a small pellet of sealing wax into the barrel by the side of the capillary tube and then warm the tube at the gas flame until the wax becomes softened and makes an air-tight joint between the capillary tube and the end of the barrel.

9. Fit a rubber teat to the open end of the barrel, and so complete a pipette which can be depended upon to always aspirate and deliver exactly 5 cm. of fluid.

Slight modification of this procedure is necessary in making tubes to measure larger volumes than say 75 c. mm. Thus to make a throttle pipette to measure 100 c. mm.:

1. Take a short length of quill tubing and draw out one end into a roomy capillary stem, and again draw out the extremity into a fine hair point, thus forming[Pg 15] a small Pasteur pipette with a hair-like capillary extremity.

2. With a standard pipette fill 100 c. mm. into the neck of this pipette, and make a scratch with a writing diamond at the upper level (a) of the mercury meniscus (Fig. 19, A).

Fig. 19.—Making throttle pipettes—large capacity

Now force the mercury down into the capillary stem as far as it will go, so as to leave the upper part of the tube in the region of the diamond scratch empty (Fig. 19, B).

3. Heat the tube in the region of the diamond scratch in the blowpipe flame, and removing the tube from the flame draw it out so that the diamond scratch now occupies a position somewhere near the centre of this new capillary portion (Fig. 19, C).[Pg 16]

4. Heat the tube in this position in the peep flame of the Bunsen burner, and draw it out into a hair-like extremity. Snap off the glass tube, leaving about 5 mm. of hair-like extremity attached to the upper capillary portion (Fig. 19, D). Allow the glass to cool.

5. Lift up the bulb by the long capillary stem and allow the mercury to return to its original position—an operation which will be facilitated by snapping off the hair-like extremity from the long piece of capillary tubing.

6. Mark on the capillary stem with a grease pencil the position of the end of the column of mercury (Fig. 19, E.)

7. Warm the capillary tubing at this spot in the peep flame of the Bunsen burner, and draw it out very slightly so that when cut at this position a pointed extremity will be obtained.

8. With a glass-cutting knife cut the capillary tube through at the point "b," and allow the mercury to run out.

9. Now apply a thick layer of sealing wax to the neck of the bulb.

10. Take a piece of 5 mm. bore glass tubing and draw it out as if making an ordinary Pasteur pipette.

11. Break the capillary portion off so as to leave a covering tube similar to that already used for the smaller graduated pipettes. Into this covering tube drop the graduated bulb and draw the capillary stem down through the conical extremity until further progress is stopped by the layer of sealing wax.

12. Warm the pipette in the gas flame so as to melt the sealing wax and make an air-tight joint.

13. Fit an india-rubber teat over the open end of the covering tube, and the automatic pipette is ready for use (Fig. 19, F).

Sedimentation Pipettes (Fig. 20).—These are prepared from 10 cm. lengths of narrow glass tubing by sealing[Pg 17] one extremity, blowing a small bulb at the centre, and plugging the open end with cotton-wool; after sterilisation the open end is provided with a short piece of rubber tubing and a glass mouthpiece. When it is necessary to observe sedimentation reactions in very small quantities of fluid, these tubes will be found much more convenient than the 5 by 0.5 cm. test-tubes previously mentioned.

Fig. 20.—Sedimentation pipette.

Pasteur pipettes fitted with india-rubber teats will also be found useful for sedimentation tests when dealing with minute quantities of serum, etc.

Fig. 21.—Fermentation tubes.

Fermentation Tubes (Fig. 21).—These are used for the collection and analysis of the gases liberated from the media during the growth of some varieties of bacteria and may be either plain (a) or graduated (b). A simple form (Fig. 21, c) may be made from 14 cm. lengths of soft glass tubing of 1.5 cm. diameter. The Bunsen flame is applied to a spot some 5 cm. from one end of such a piece of tubing and the tube slightly drawn out to form a constriction, the constricted part[Pg 18] is bent in the bat's-wing flame, to an acute angle, and the open extremity of the long arm sealed off in the blowpipe flame. The open end of the short arm is rounded off and then plugged with cotton-wool, and the tube is ready for sterilisation.

CLEANING OF GLASS APPARATUS.

All glassware used in the bacteriological laboratory must be thoroughly cleaned before use, and this rule applies as forcibly to new as to old apparatus, although the methods employed may vary slightly.

To Clean New Test-tubes.—

1. Place the tubes in a bucket or other convenient receptacle, fill with water and add a handful of "Sapon" or other soap powder. See that the tubes are full and submerged.

2. Fix the bucket over a large Bunsen flame and boil for thirty minutes—or boil in the autoclave for a similar period.

3. Cleanse the interior of the tubes with the aid of test-tube brushes, and rinse thoroughly in cold water.

4. Invert the tubes and allow them to drain completely.

5. Dry the tubes and polish the glass inside and out with a soft cloth, such as selvyt.

New flasks, plates, and capsules must be cleaned in a similar manner.

To Clean New Graduated Pipettes.—

1. Place the pipettes in a convenient receptacle, filled with water to which soap powder has been added.

2. Boil the water vigorously for twenty minutes over a Bunsen flame.

3. Rinse the pipettes in running water and drain.

4. Run distilled water through the pipettes and drain.[Pg 19]

5. Run rectified spirits through the pipette and drain as completely as possible.

6. Place the pipettes in the hot-air oven (vide page 31), close the door, open the ventilating slide, and run the temperature slowly up to about 80° C. Turn off the gas and allow the oven to cool.

Or 6a. Attach each pipette in turn to the rubber tube of the foot bellows, or blowpipe air-blast, and blow air through the pipette until the interior is dry.

Glassware that has already been used is regarded as infected, and is treated in a slightly different manner.

Infected Test-tubes.—

1. Pack the tubes in the wire basket of the autoclave (having previously removed the cotton-wool plugs, caps, etc.), in the vertical position, and before replacing the basket see that there is a sufficiency of water in the bottom of the boiler. Now attach a piece of rubber tubing to the nearest water tap, and by means of this fill each tube with water.

2. Disinfect completely by exposing the tubes, etc., to a temperature of 120° C. for twenty minutes (vide page 37).

(If an autoclave is not available, the tubes must be placed in a digester, or even a large pan or pail with a tightly fitting cover, and boiled vigorously for some thirty to forty-five minutes to ensure disinfection.)

3. Whilst still hot, empty each tube in turn and roughly clean its interior with a stiff test-tube brush.

4. Place the tubes in a bucket or other convenient receptacle, fill with water and add a handful of Sapon or other soap powder. See that the tubes are full and submerged.

5. Fix the bucket over a large Bunsen flame and boil for thirty minutes.

6. Cleanse the interior of the tubes with the aid of test-tube brushes, and rinse thoroughly in cold water.[Pg 20]

7. Drain off the water and immerse tubes in a large jar containing water acidulated with 2 to 5 per cent. hydrochloric acid. Allow them to remain there for about fifteen minutes.

8. Remove from the acid jar, drain, rinse thoroughly in running water, then with distilled water.

9. Invert the tubes and allow them to drain completely.

Dry the tubes and polish the glass inside and out with a soft cloth, such as selvyt.

Infected flasks, plates, and capsules must be treated in a similar manner.

Flasks which have been used only in the preparation of media must be cleaned immediately they are finished with. Fill each flask with water to which some soap powder and a few crystals of potassium permanganate have been added, and let boil over the naked flame. The interior of the flask can then usually be perfectly cleaned with the aid of a flask brush, but in some cases water acidulated with 5 per cent. nitric acid, or a large wad of wet cotton-wool previously rolled in silver sand, must be shaken around the interior of the flask, after which rinse thoroughly with clean water, dry, and polish.

Infected Pipettes.—

1. Plunge infected pipettes immediately after use into tall glass cylinders containing a 2 per cent. solution of lysol, and allow them to remain therein for some days.

2. Remove from the jar and drain. Boil in water to which a little soap has been added, for thirty minutes.

3. Rinse thoroughly in cold water.

4. Immerse in 5 per cent. nitric acid for an hour or two.[Pg 21]

5. Rinse again in running water to remove all traces of acid.

6. Complete the cleaning as described under "new pipettes."

When dealing with graduated capillary pipettes employed for blood or serum work (whether new or infected), much time is consumed in the various steps from 5 onward, and the cleansing process can be materially hastened if the following device is adopted.

Fit up a large-sized Kitasato's filter flask to a Sprengel's suction pump or a Geryk air pump (see page 43). To the side tubulure of the filter flask attach a 20 cm. length of rubber pressure tubing having a calibre sufficiently large to admit the ends of the pipettes.

Next fill a small beaker with distilled water. Attach the first pipette to the free end of the rubber tubing, place the pipette point downward in the beaker of water and start the pump (Fig. 22).

Fig. 22.—Cleaning blood pipettes.

When all the water has been aspirated through the pipette into the filter flask, fill the beaker with rectified spirit and when this is exhausted refill with ether. Detach the pipette and dry in the hot-air oven.

Slides and cover-slips (Fig. 23), when first purchased,[Pg 22] have "greasy" surfaces, upon which water gathers in minute drops and effectually prevents the spreading of thin, even films.

Microscopical Slides.—The slides in general use are those known as "three by one" slips (measuring 3 inches by 1 inch, or 76 by 26 mm.), and should be of good white crown glass, with ground edges.

New slides should be allowed to remain in alcohol acidulated with 5 per cent. hydrochloric acid for some hours, rinsed in running water, roughly drained on a towel, dried, and finally polished with a selvyt cloth.

Fig. 23.—Slides and cover-slips, actual size.

If only a few slides are required for immediate use a good plan is to rub the surface with jeweler's emery paper (Hubert's 00). A piece of hard wood 76×26×26 mm. with a piece of this emery paper gummed tightly around it is an exceedingly useful article on the microscope bench.

Cover-slips.—The most useful sizes are the 19 mm. squares for ordinary cover-glass film preparations, and 38 by 19 mm. rectangles for blood films and serial sections; both varieties must be of "No. 1" thickness, which varies between 0.15 and 0.22 mm., that they may be available for use with the high-power immersion lenses.

Cover-slips should be cleaned in the following manner:

1. Drop the cover-slips one by one into an enamelled iron pot or tall glass beaker, containing a 10 per cent. solution of chromic acid.[Pg 23]

2. Heat over a Bunsen flame and allow the acid to boil gently for twenty minutes.

3. Turn the cover-slips out into a flat glass dish and wash in running water under the tap until all trace of yellow colour has disappeared. During the washing keep the cover-slips in motion by imparting a rotatory movement to the dish.

4. Wash in distilled water in a similar manner.

5. Wash in rectified spirit.

6. Transfer the cover-slips, by means of a pair of clean forceps, previously heated in the Bunsen flame to destroy any trace of grease, to a small beaker of absolute alcohol.

Drain off the alcohol and transfer the cover-slips, by means of the forceps, to a wide-mouthed glass pot, containing absolute alcohol, in which they are to be stored, and stopper tightly.

Used Slides and Cover-slips.—Used slides with the mounted cover-slip preparations, and cover-slips used for hanging-drop mounts, should, when discarded, be thrown into a pot containing a 2 per cent. solution of lysol.

After immersion therein for a week or so, even the cover-slips mounted with Canada balsam can be readily detached from their slides.

Slides.—

1. Wash the slides thoroughly in running water.

2. Boil the slides in water to which "sapon" has been added, for half an hour.

3. Rinse thoroughly in cold water.

4. Dry and polish with a dry cloth.[Pg 24]

Cover-slips.—

1. Wash the cover-slips thoroughly in running water.

2. Boil the cover-slips in 10 per cent. solution of chromic acid, as for new cover-slips.

3. Wash thoroughly in running water.

4. Pick out those cover-slips which show much adherent dirty matter, and rub them between thumb and forefinger under the water tap. The dirt usually rubs off easily, as it has become friable from contact with the chromic acid.

5. Return all the cover-slips to the beaker, fill in fresh chromic acid solution, and treat as new cover-slips.

PLUGGING TEST-TUBES AND FLASKS.

Before sterilisation all test-tubes and flasks must be carefully plugged with cotton-wool, and for this purpose best absorbent cotton-wool (preferably that put up in cylindrical one-pound packets and interleaved with tissue paper—known as surgeons' wool) should be employed.

1. For a test-tube or a small flask, tear a strip of cotton-wool some 10 cm. long by 2 cm. wide from the roll.

2. Turn in the ends neatly and roll the strip of wool lightly between the thumb and fingers of both hands to form a long cylinder.

3. Double this at the centre and introduce the now rounded end into the open mouth of the tube or flask.

4. Now, whilst supporting the wool between the thumb and fingers of the right hand, rotate the test-tube[Pg 25] between those of the left, and gradually screw the plug of wool into its mouth for a distance of about 2.5 cm., leaving about the same length of wool projecting.

Fig 24..—Plugging test-tubes: a, cylinder of wool being rolled; b, cylinder of wool being doubled; c, cylinder of wool being inserted in tube.

The plug must be firm and fit the tube or flask fairly tightly, sufficiently tightly in fact to bear the weight of the glass plus the amount of medium the vessel is intended to contain, but not so tightly as to prevent it from being easily removed by a screwing motion when grasped between the fourth, or third and fourth, fingers, and the palm of the hand.

For a large flask a similar but larger strip of wool must be taken; the method of making and inserting the plug is identical.

[Pg 26]

III. METHODS OF STERILISATION.

STERILISING AGENTS.

Sterilisation—i. e., the removal or the destruction of germ life—may be effected by the use of various agents. As applied to the practical requirements of the bacteriological laboratory, many of these agents, such as electricity, sunlight, etc., are of little value, others are limited in their applications; others again are so well suited to particular purposes that their use is almost entirely restricted to such.

The sterilising agents in common use are:

Chemical Reagents.—Disinfectants (for the disinfection of glass and metal apparatus and of morbid tissues).

Physical Agents. Heat.—(a) Dry Heat:

1. Naked flame (for the sterilisation of platinum needles, etc.).

2. Muffle furnace (for the sterilisation of filter candles, and for the destruction of morbid tissues).

3. Hot air (for the sterilisation of all glassware and of metal apparatus).

(b) Moist Heat:

1. Water at 56° C. (for the sterilisation of certain albuminous fluids).

2. Water at 100° C. (for the sterilisation of surgical instruments, rubber tubing, and stoppers, etc.).

3. Streaming steam at 100° C. (for the sterilisation of media).

4. Superheated steam at 115° C. or 120° C. (for the disinfection of contaminated articles and the destruction of old cultivations of bacteria).[Pg 27]

Filtration.—

1. Cotton-wool filters (for the sterilisation of air and gases).

2. Porcelain filters (for the sterilisation of various liquids).

METHODS OF APPLICATION.

Chemical Reagents, such as belong to the class known as antiseptics (i. e., substances which inhibit the growth of, but do not destroy, bacterial life), are obviously useless. Disinfectants or germicides (i. e., substances which destroy bacterial life), on the other hand, are of value in the disinfection of morbid material, and also of various pieces of apparatus, such as pipettes, pending their cleansing and complete sterilisation by other processes. To this class (in order of general utility) belong:

Formaldehyde is a powerful germicide, but its penetrating vapor restricts its use. These disinfectants are but little used in the final sterilisation of apparatus, chiefly on account of the difficulty of effecting their complete removal, for the presence of even traces of these chemicals is sufficient to so inhibit or alter the growth of bacteria as to vitiate subsequent experiments conducted by the aid of apparatus sterilised in this manner.[Pg 28]

Dry Heat.—The naked flame of the Bunsen burner is invariably used for sterilising the platinum needles (which are heated to redness) and may be employed for sterilising the points of forceps, or other small instruments, cover-glasses, pipettes, etc., a very short exposure to this heat being sufficient.

Ether Flame.—In an emergency small instruments, needles, etc., may be sterilised by dipping them in ether and after removal lighting the adherent fluid and allowing it to burn off the surface of the instruments. Repeat the process twice. It may then be safely assumed that the apparatus so treated is sterile.

Fig. 25.—Muffle furnace.

Muffle Furnace (Fig. 25).—Although this form of heat is chiefly used for the destruction of the dead bodies of small infected animals, morbid tissues, etc., it is also employed for the sterilisation of porcelain filter candles (vide p. 42).

Filter candles are disinfected immediately after use by boiling in a beaker of water for some fifteen or twenty minutes. This treatment, however, leaves the dead bodies of the bacteria upon the surface and blocking the interstices of the filter.

To destroy the organic matter and prepare the filter candle for further use proceed as follows:[Pg 29]

1. Roll each bougie up in a piece of asbestos cloth, secure the ends of the cloth with a few turns of copper wire, and place inside the muffle (a small muffle 76×88×163 mm. will hold perhaps four small filter candles).

2. Light the gas and raise the contents of the muffle to a white heat; maintain this temperature for five minutes.

3. Extinguish the gas, and when the muffle has become quite cold remove the filter candles, and store them (without removing the asbestos wrappings) in sterile metal boxes.

Hot Air.—Hot air at 150° C. destroys all bacteria, spores, etc:, in about thirty minutes; a momentary exposure to a temperature of 175° to 180° C. will effect the same result and offers the more convenient method of sterilisation. This method is only applicable to glass and metallic substances, and the small bulk of cotton-wool comprised in the test-tube plugs, etc. Large masses of fabric are not effectually sterilised by dry heat—short of charring—as its power of penetration is not great.

Sterilisation by hot air is effected in the hot-air oven (Fig. 18). This is a rectangular, double-walled metal box, mounted on a stand and heated from below by a large Bunsen burner. The interior of the oven is provided with loose shelves upon which the articles to be sterilised are arranged, either singly or packed in square wire baskets or crates, kept specially for this purpose. One of the sides is hinged to form a door. The central portion of the metal bottom, on which the Bunsen flame would play, is cut away, and replaced by firebrick plates, which slide in metal grooves and[Pg 30] are easily replaced when broken or worn out. The top of the oven is provided with a perforated ventilator slide and two tubulures, the one for the reception of a centigrade thermometer graduated to 200° or 250°C., the other for a thermo-regulator. An ordinary mercurial thermo-regulator may be used but it is preferable to employ a regulating capsule of the Hearson type (see p. 219) with a spring arm adjusted to the lever so that when the boiling-point of the capsule (e. g., 175°C.) is reached the gas supply is absolutely cut off and the jet cannot again be lighted until the spring-arm has been readjusted by hand. The thermo-regulator is by no means a necessity, and may be replaced by a large bore thermometer with a sliding platinum point, connected with an electric bell, which can be easily adjusted to ring at any given temperature. Even if the steriliser is provided with the capsule regulator above described the contact thermometer should also be fitted.

Fig. 26.—Hot-air oven.

[Pg 31]

To Use the Hot-air Oven.—

1. Place the crates of test-tubes, metal cases containing plates and pipettes, loose apparatus, etc., inside the oven, taking particular care that none of the cotton-wool plugs are in contact with the walls, otherwise the heat transmitted by the metal will char or even flame them.

2. Close the oven door, and open the ventilating slide, in order that any moisture left in the tubes, etc., may escape; light the gas below; set the electric alarm to ring at 100°C.

3. When the temperature of the oven has reached 100°C., close the ventilating slide; reset the alarm to ring at 175°C.

4. Run the temperature up to 175°C.

5. Extinguish the gas at once, and allow the apparatus to cool.

6. When the temperature of the interior, as recorded by the thermometer, has fallen to 60°C.—but not before—the door may be opened and the sterile articles removed and stored away.

[Pg 32]

On removal from the oven, the cotton-wool plugs will probably be slightly brown in colour.

Metal instruments, such as knives, scissors, and forceps, may be sterilised in the hot-air oven as described above, but exposure to 175° C. is likely to seriously affect the temper of the steel and certainly blunts the cutting edges. If, however, it is desired to sterilise surgical instruments by hot air, they should be packed in a metal box, or boxes, and heated to 130° C. and retained at that temperature for about thirty minutes.

Moist Heat.—Water at 56° C.—This temperature, if maintained for thirty minutes, is sufficient to destroy the vegetative forms of bacteria, but has practically no effect on spores. Its use is limited to the sterilisation of such albuminous "fluid" media as would coagulate at a higher temperature.

Method.—

1. Fit up a water-bath, heated by a Bunsen flame which is controlled by a thermo-regulator, so that the temperature of the water remains at 56° C.

2. Immerse the tubes or flasks containing the albuminous fluid in the water-bath so that the upper level of such fluid is at least 2 cm. below the level of the water. (The temperature of the bath will now fall somewhat, but after a few minutes will again rise to 56° C).

3. After thirty minutes' exposure to 56° C, extinguish the gas, remove the tubes or flasks from the bath, and subject them to the action of running water so that their contents are rapidly cooled.

4. The vegetative forms of bacteria present in the liquid being killed, stand it for twenty-four hours in a cool, dark place; at the end of that time some at least of such spores as may be present will have germinated and assumed the vegetative form.[Pg 33]

5. Destroy these new vegetative forms by a similar exposure to 56° C. on the second day, whilst others, of slower germination, may be caught on the third day, and so on.

6. In order to ensure thorough sterilisation, repeat the process on each of six successive days.

This method of exposing liquids to a temperature of 56° C. in a water-bath for half an hour on each of six successive days is termed fractional sterilisation.

Water at 100°C. destroys the vegetative forms of bacteria almost instantaneously, and spores in from five to fifteen minutes. This method of sterilisation is applicable to the metal instruments, such as knives, forceps, etc., used in animal experiments; syringes, rubber corks, rubber and glass tubing, and other small apparatus, and is effected in what is usually spoken of as the "water steriliser" (Fig. 27).

Fig. 27.—Water sterilizer.

This is a rectangular copper box, 26 cm. long, 18 cm. wide, and 12 cm. deep, mounted on legs, heated from below by a Bunsen or radial gas burner, and containing a movable copper wire tray, 2 cm. smaller in every[Pg 34] dimension than the steriliser itself, and provided with handles. The top of the steriliser is hinged to form a lid.

Method.—

1. Place the instruments, etc., to be sterilised inside the copper basket, and replace the basket in the steriliser.

2. Pour a sufficient quantity of water into the steriliser, shut down the lid, and light the gas below.

Fig. 28.—Koch's steriliser.

Fig. 29.—Arnold's steriliser.

3. After the water has boiled and steam has been issuing from beneath the lid for at least ten minutes, extinguish the gas, open the lid, and lift out the wire basket by its handles and rest it diagonally on the walls of the steriliser; the contained instruments, etc., are now sterile and ready for use.

4. After use, or when accidentally contaminated, replace the instruments in the basket and return that to the steriliser; completely disinfect by a further boiling for fifteen minutes.

5. After disinfection, and whilst still hot, take out[Pg 35] the instruments, dry carefully and at once, and return them to their store cases.

Streaming steam—i. e., steam at 100°C.—destroys the vegetative forms of bacteria in from fifteen to twenty minutes, and the sporing forms in from one to two hours. This method is chiefly used for the sterilisation of the various nutrient media intended for the cultivation of bacteria, and is carried out in a steam kettle of special construction, known as Koch's steam steriliser (Fig. 28) or in one of its many modifications, the most efficient of which is Arnold's (Fig. 29).

The steam steriliser in its simplest form consists of a tall tinned-iron or copper cylindrical vessel, divided into two unequal parts by a movable perforated metal diaphragm, the lower, smaller portion serving for a water reservoir, and the upper part for the reception of wire baskets containing the articles to be sterilised. The vessel is closed by a loose conical lid, provided with handles, and perforated at its apex by a tubulure; it is mounted on a tripod stand and heated from below by a Bunsen burner. The more elaborate steriliser is cased with felt or asbestos board, and provided with a water gauge, also a tap for emptying the water compartment.

To Use the Steam Steriliser.—

1. Fill the water compartment to the level of the perforated diaphragm, place the lid in position, and light the Bunsen burner.

2. After the water has boiled, allow sufficient time to elapse for steam to replace the air in the sterilising compartment, as shown by the steam issuing in a steady, continuous stream from the tubulure in the lid.

3. Remove the lid, quickly lower the wire basket containing media tubes, etc., into the sterilising compartment until it rests on the diaphragm, and replace the lid.[Pg 36]

4. After an interval of twenty minutes in the case of fluid media, or thirty minutes in the case of solid media, take off the lid and remove the basket with its contents.

5. Now, but not before, extinguish the gas.

This treatment will destroy any vegetative forms of bacteria; during the hours of cooling any spores present will germinate, and the young organisms will be destroyed by repeating the process twenty-four hours later; a third sterilisation after a similar interval makes assurance doubly sure.

The method of sterilising by exposure to streaming steam at 100° C. for twenty minutes on each of three consecutive days is termed discontinuous or intermittent sterilisation.

Exposure to steam at 100° C. for a period of one or two hours, or continuous sterilisation, cannot always be depended upon and is therefore not to be recommended.

Superheated steam—i. e., steam under pressure (see Pressure-temperature table, Appendix, page 500) in sealed vessels at a temperature of 115° C.—will destroy both the vegetative and the sporing forms of bacteria within fifteen minutes; if the pressure is increased, and the temperature raised to 120° C., the same end is attained in ten minutes. This method was formerly employed for the sterilisation of media (and indeed is so used in some laboratories still), but most workers now realise that media subjected to this high temperature undergo hydrolytic changes which render them unsuitable for the cultivation of the more delicate micro-organisms. The use of superheated steam should be restricted almost entirely to the disinfection of such contaminated articles, old cultivations, etc.,[Pg 37] as cannot be dealt with by dry heat or the actual furnace. Sterilisation by means of superheated steam is carried out in a special boiler—Chamberland's autoclave (Fig. 30). The autoclave consists of a stout copper cylinder, provided with a copper or gun-metal lid, which is secured in place by means of bolts and thumbscrews, the joint between the cylinder and its lid being hermetically sealed by the interposition of a rubber washer. The cover is perforated for a branched tube carrying a vent cock, a manometer, and a safety valve. The copper boiler is mounted in the upper half of a cylindrical sheet-iron case—two concentric circular rows of Bunsen burners, each circle having an independent gas-supply, occupying the lower half. In the interior of the boiler is a large movable wire basket, mounted on legs, for the reception of the articles to be sterilised.

To Use the Autoclave.—

1. Pack the articles to be sterilised in the wire basket.

2. Run water into the boiler to the level of the bottom of the basket; also fill the contained flasks and tubes with water.

3. See that the rubber washer is in position, then replace the cover and fasten it tightly on to the autoclave by means of the thumbscrews.

4. Open the vent cock and light both rings of burners.

5. When steam is issuing in a steady, continuous stream from the vent tube, shut off the vent cock and extinguish the outer ring of gas burners.

6. Wait until the index of the manometer records a temperature of 120° C., then regulate the gas and the spring safety valve in such a manner that this temperature is just maintained, and leave it thus for twenty minutes. In the more expensive patterns of autoclave this regulation of the safety valve is carried[Pg 38] out automatically, the manometer being fitted with an adjustable pointer which can be set to any required pressure-temperature and so arranged that when the index of the manometer coincides with the adjustable hand the safety valve is opened.

7. Extinguish the gas and allow the manometer index to fall to zero.

Fig. 30.—Chamberland's Autoclave.

8. Now open the vent cock slowly, and allow the internal pressure to adjust itself to that of the atmosphere.

9. Remove the cover and take out the sterilised contents.

Sterilisation Periods.—An exceedingly useful device for the timing of sterilisation periods (and indeed for many other operations in the laboratory) is the

ELECTRIC SIGNAL TIMING CLOCK.

This is a clock of American type in which the face is surrounded by a metal plate having a series of 60[Pg 39] holes at equal distances apart, corresponding to the minutes on the dial. This plate is connected with one of the poles of a dry battery, the other pole of which is connected to the metal case of the clock for the purpose of actuating an ordinary magnet alarm bell. In the centre of each of the holes in the plate a metal rod is fixed, which then passes through an insulating ring and projects inside the clock face, where it makes contact with the hour hand. The clock is mounted on a heavy base, with a key-board containing 20 numbered plugs. If one of the plugs is inserted in a hole in the plate it makes contact with the rod, and when the hour hand of the clock touches the other end the circuit is completed and the bell starts ringing. The period of this friction contact is approximately 20 seconds. The clock can therefore be used for electrically noting the periods of time from one minute by multiples of one minute up to one hour.

Fig. 31.—Electric signal timing clock.

[Pg 40]

Filtration.—(a) Cotton-wool Filter.—Practically the only method in use in the laboratory for the sterilisation of air or of a gas is by filtration through dry cotton-wool or glass-wool, the fibres of which entangle the micro-organisms and prevent their passage.

Perhaps the best example of such a filter is the cotton-wool plug which closes the mouth of a culture tube. Not only does ordinary diffusion take place through it, but if a tube plugged in the usual manner with cotton-wool is removed from the hot incubator, the temperature of the contained air rapidly falls to that of the laboratory, and a partial vacuum is formed; air passes into the tube, through the cotton-wool plug, to restore the equilibrium, and, so long as the plug remains dry, in a germ-free condition. If, however, the plug becomes moist, either by absorption from the atmosphere, or from liquids coming into contact with it, micro-organisms (especially the mould fungi) commence to multiply, and the long thread forms rapidly penetrate the substance of the plug, and gain access to and contaminate the interior of the tube.

Fig. 32.—Cotton-wool air filter.

Method.—

If it is desired to sterilise gases before admission to a vessel containing a pure cultivation of a micro-organism, as, for instance, when forcing a current of oxygen over or through a broth cultivation of the diphtheria bacillus, this can be readily effected as follows:[Pg 41]

1. Take a length of glass tubing of, say, 1.5 cm. diameter, in the centre of which a bulb has been blown, fill the bulb with dry cotton-wool (Fig. 32), wrap a layer of cotton-wool around each end of the tube, and secure in position with a turn of thin copper wire or string; then sterilise the piece of apparatus in the hot-air oven.

2. Prepare the cultivation in a Ruffer or Woodhead flask (Fig. 33) the inlet tube of which has its free extremity enveloped in a layer of cotton-wool, secured by thread or wire, whilst the exit tube is plugged in the usual manner.

Fig. 33.—Ruffer's flask.

3. Sterilise a short length of rubber tubing by boiling. Transfer it from the boiling water to a beaker of absolute alcohol.

4. When all is ready remove the rubber tube from the alcohol by means of a pair of forceps, drain it thoroughly, and pass through the flame of a Bunsen burner to burn off the last traces of alcohol.

5. Remove the cotton-wool wraps from the entry tube of the flask and from one end of the filter tube and rapidly couple them up by means of the sterile rubber tubing.[Pg 42]

6. Connect the other end of the bulb tube with the delivery tube from the gas reservoir.

The gas in its passage through the dry sterile cotton-wool in the bulb of the filter tube will be freed from any contained micro-organisms and will enter the flask in a sterile condition.

(b) Porcelain Filter.—The sterilisation of liquids by filtration is effected by passing them through a cylindrical vessel, closed at one end like a test-tube, and made either of porous "biscuit" porcelain, hard-burnt and unglazed (Chamberland system), or of Kieselguhr, a fine diatomaceous earth (Berkefeld system), and termed a "bougie" or "candle" (Fig. 34).

In this method the bacteria are retained in the pores of the filter while the liquid passes through in a germ-free condition.

It is obvious that to be effective the pores of the filter must be extremely minute, and therefore the rate of filtration will usually be slow. Chamberland filter candles possess finer channels than Berkefeld candles and consequently filter much more slowly. To overcome this disadvantage, either aspiration or pressure, or a combination of these two forces, may be employed to hasten the process.

Doultons white porcelain filters it may be noted are as efficient as the Chamberland candles and filter rather more rapidly.

Apparatus Required.—

1. Separatory funnel containing the unfiltered fluid.

2. Sterile filter candle (Fig. 34), the open end fitted with a rubber stopper (Fig. 34, a) perforated to receive the delivery tube of the separatory funnel, and its neck passed through a large rubber washer (Fig. 34, b) which fits the mouth of the filter flask.

3. Sterile filter flask of suitable size, for the reception of the filtered fluid, its mouth closed by a cotton-wool plug.[Pg 43]

4. Water injector Sprengel (see Fig. 38, c) pump, or Geryk's pump (an air pump on the hydraulic principle, sealed by means of low vapor-tension oil, Fig. 35).

If this latter is employed, a Wulff's bottle, fitted as a wash-bottle and containing sulphuric acid, must be interposed between the filter flask and the pump, in order to prevent moist air reaching the oil in the pump.

5. Air filter (vide page 40) sterilised.

6. Pressure tubing.

7. Screw clamps (Fig. 36).

Method.—

1. Couple the exhaust pipe of the suction pump with the lateral tube of the filter flask (first removing the cotton-wool plug from this latter), by means of pressure tubing, interposing, if necessary, the wash-bottle of sulphuric acid.

Fig. 34.—Porcelain filter candle.

Fig. 35.—Geryk air pump.

[Pg 44]

2. Remove the cotton-wool plug from the neck of the filter flask and adjust the porcelain candle in its place.

Fig. 36.—Screw clamps.

3. Attach the nozzle of the separatory funnel to the filter candle by means of the perforated rubber stopper (Fig. 37).

Fig. 37.—Apparatus arranged for filtering—aspiration.

4. Open the tap of the funnel, and exhaust the air from the filter flask and wash-bottle; maintain the vacuum until the filtration is complete.

5. When the filtration is completed close the tap of[Pg 45] the funnel; adjust a screw clamp to the pressure tubing attached to the lateral branch of the filter flask; screw it up tightly, and disconnect the acid wash-bottle.

6. Attach the air filter to the open end of the pressure tubing; open the screw clamp gradually, and allow filtered air to enter the flask, to abolish the negative pressure.

7. Detach the rubber tubing from the lateral branch of the flask, flame the end of the branch in the Bunsen, and plug its orifice with sterile cotton-wool.

8. Remove the filter candle from the mouth of the flask, flame the mouth, and plug the neck with sterile cotton-wool.

9. Disinfect the filter candle and separatory funnel by boiling.

If it is found necessary to employ pressure in addition to or in place of suction, insert a perforated rubber stopper into the mouth of the separatory funnel and secure in position with copper wire; next fit a piece of glass tubing through the stopper, and connect the external orifice with an air-pressure pump of some kind (an ordinary foot pump such as is employed for inflating bicycle tyres is one of the most generally useful, for this purpose) or with a cylinder of compressed air or other gas.

In order to filter a large bulk of fluid very rapidly it is necessary to use a higher pressure than glass would stand, and in these cases the metal receptacle designed by Pakes (Fig. 38, a), to hold the filter candle itself as well as the fluid to be filtered, should be employed. (A vacuum must also be maintained in the filter flask, by means of an exhaust pump, during the entire process.)

This piece of apparatus consists of a brass cylinder, capacity 2500 c.c., with two shoulders; and an opening in the neck at each end, provided with screw threads.

A nut carrying a pressure gauge fits into the top[Pg 46] screw; and into the bottom is fitted a brass cylinder carrying the filter candle and prolonged downwards into a delivery tube. Leakage is prevented by means of rubber washers.

Into the top shoulder a tube is inserted, bent at right angles and provided with a tap. All the brass-work is tinned inside (Fig. 38, a). In use the reservoir is generally mounted on a tripod stand.

To Sterilise.—

1. Insert the filter candle into its cylinder and screw this loosely on.

Fig. 38.—Pakes' filtering reservoir—pressure and aspiration.

2. Wrap a layer of cotton-wool around the delivery tube and fasten in position.

3. Remove the nut carrying the pressure gauge and plug the neck with cotton-wool.[Pg 47]

4. Heat the whole apparatus in the autoclave at 120° C. for twenty minutes.

Method.—

1. Remove the apparatus from the autoclave, and allow it to cool.

2. Screw home the box carrying the bougie.

3. Set the apparatus up in position, with its delivery tube (from which the cotton-wool wrapping has been removed) passing through a perforated rubber stopper in the neck of a filter flask.

Fig. 39.—Closed candle arranged for filtering.

4. Fill the fluid to be filtered into the cylinder and screw on the nut carrying the pressure gauge. (This nut should be immersed in boiling water for a few minutes previous to screwing on, in order to sterilise it.)

5. Connect the horizontal arm of the entry tube with a cylinder of compressed oxygen (or carbon dioxide, Fig. 38, b), by means of pressure tubing.

6. Connect the lateral arm of the filter flask with the exhaust pump (Fig. 38, c) and start the latter working.[Pg 48]

7. Open the tap of the gas cylinder; then open the tap on the entry tube of the filter cylinder and raise the pressure in its interior until the desired point is recorded on the manometer. Maintain this pressure, usually one or one and a half atmospheres, until filtration is completed, by regulating the tap on the entry tube.

Some forms of filter candle are made with the open end contracted into a delivery nozzle, which is glazed. In this case the apparatus is fitted up in a slightly different manner; the fluid to be filtered is contained in an open cylinder into which the candle is plunged, while its delivery nozzle is connected with the filter flask by means of a piece of flexible pressure tubing (previously sterilised by boiling), as in figure 39.

[Pg 49]

IV. THE MICROSCOPE.

The essentials of a microscope for bacteriological work may be briefly summed up as follows:

Fig. 40.—Microscope stand.

The instrument, of the monocular type, must be of good workmanship and well finished, rigid, firm, and free from vibration, not only when upright, but also when inclined to an angle or in the horizontal position. The various joints and movements must work smoothly and precisely, equally free from the defects of "loss of time" and "slipping." All screws, etc., should conform[Pg 50] to the Royal Microscopical Society's standard. It must also be provided with good lenses and a sufficiently large stage. The details of its component parts, to which attention must be specially directed, are as follows:

Fig. 41.—Foot, three types.

1. The Base or Foot (Fig. 40, a).—Two elementary forms—the tripod (Fig. 41, a) and the vertical column set into a plate known as the "horse-shoe" (Fig. 41, b)—serve as the patterns for countless modifications in shape and size of this portion of the stand. The chief desiderata—stability and ease of manipulation—are attained in the first by means of the "spread" of the three feet, which are usually shod with cork; in the second, by the dead weight of the foot-plate. The tripod is mechanically the more correct form, and for practical use is much to be preferred. Its chief rival, the Jackson foot (Fig. 41, c), is based upon the same principle, and on the score of appearance has much to recommend it.

2. The body tube (Fig. 40, b) may be either that known as the "long" or "English" (length 250 mm.), or the "short" or "Continental" (length 160 mm.). Neither length appears to possess any material advantage over the other, but it is absolutely necessary to secure objectives which have been manufactured for the particular tube length chosen. In the high-class microscope of the present day the body tube is usually[Pg 51] shorter than the Continental, but is provided with a draw tube which, when fully extended, gives a tube length greater than the English, thus permitting the use of either form of objective.

Fig. 42.—Coarse adjustment.

Fig. 43.—Fine adjustment.

3. The coarse adjustment (Fig. 40, c) should be a rack-and-pinion movement, steadiness and smoothness of action being secured by means of accurately fitting dovetailed bearings and perfect correspondence between the teeth of the rack and the leaves of the pinion (Fig. 42). Also provision should be made for taking up the "slack" (as by the screws AA, Fig. 42).

4. The fine adjustment (Fig. 40, d) should on no account depend upon the direct action of springs, but[Pg 52] should be of the lever pattern, preferably the Nelson (Fig. 43). In this form the unequal length of the arms of the lever secures very delicate movement, and, moreover, only a small portion of the weight of the body tube is transmitted to the thread of the vertical screw actuating the movement.

Fig. 44.—Spindle head to fine adjustment.

A spindle milled head (Fig. 44) will be found a very useful device to have fitted in place of the ordinary milled head controlling the fine adjustment. In this contrivance the axis of the milled head is prolonged upward in a short column, the diameter of which is one-sixth of that of the head. The spindle can be rapidly rotated between the fingers for medium power adjustments while the larger milled head can be slowly moved when focussing high powers.

5. The stage (Fig. 40, e) should be square in shape and large in area—at least 12 cm.—flat and rigid, in order to afford a safe support for the Petri dish used for plate cultivations; and should be supplied with spring clips (removable at will) to secure the 3 by 1 glass slides.

A mechanical stage must be classed as a necessity rather than a luxury so far as the bacteriologist is concerned, as when working with high powers, and especially when examining hanging-drop specimens, it is almost impossible to execute sufficiently delicate movements with the fingers. In selecting a mechanical stage, preference should be given to one which forms an integral part of the instrument (Fig. 45) rather than one which needs to be clamped on to an ordinary plain stage every time it is required, and its traversing movements should be controlled by stationary milled heads (Fig. 45, AA'). The shape of the aperture is a not unimportant point; it should be square to allow of free movement over the substage[Pg 53] condenser. The mechanical stage should be tapped for three (removable) screw studs to be used in place of the sliding bar, so that if desired the Vernier finder (Fig. 45, BB'), such as is usually fitted to this class of stage, or a Maltwood finder, may be employed.

Fig. 45.—Mechanical stage.

Fig. 46.—Iris diaphragm.

6. Diaphragm.—Separate single diaphragms must be avoided; a revolving plate pierced with different sized apertures and secured below the stage is preferable, but undoubtedly the best form is the "iris"[Pg 54] diaphragm (Fig. 46) which enters into the construction of the substage condenser.

7. The substage condenser is a necessary part of the optical outfit. Its purpose is to collect the beam of parallel rays of light reflected by the plane mirror, by virtue of a short focus system of lenses, into a cone of large aperture (reducible at will by means of an iris diaphragm mounted as a part of the condenser), which can be accurately focussed on the plane of the object. This focussing must be performed anew for each object, on account of the variation in the thickness of the slides.

The form in most general use is that known as the Abbé (Fig. 47) and consists of a plano-convex lens mounted above a biconvex lens. This combination is carried in a screw-centering holder known as the substage below the stage of the microscope (Fig. 40 f), and must be accurately adjusted so that its optical axis coincides with that of the objective. Vertical movement of the entire substage apparatus effected by means of a rack and pinion is a decided advantage, and some means should be provided for temporarily removing the condenser from the optical axis of the microscope.

Fig. 47—Optical part of Abbé illuminator.

With the oil immersion objective, however, an achromatic condenser, giving an illuminating cone of about 0.9, should be used if the full value of the lens is to be obtained. It is generally assumed that a good objective requires an illuminating cone equivalent to two-thirds of its numerical aperture. The best Abbé condenser transmits a cone of about .45 whilst the aperture of the 1/12 inch immersion lenses of different makers varies from 1.0 to 1.4, hence, the efficiency of these lenses is much curtailed if the condenser is merely[Pg 55] the Abbé. These improved condensers must be absolutely centered to the objective and capable of very accurate focussing otherwise much of their value is lost.

8. Mirrors.—Below the substage condenser is attached a gymbal carrying a reversible circular frame with a plane mirror on one side and a concave mirror on the other (Fig. 40, g). The plane mirror is that usually employed, but occasionally, as for example when using low powers and with the condenser racked down and thrown out of the optical axis, the concave mirror is used.

9. Oculars, or Eyepieces.—Those known as the Huyghenian oculars (Fig. 48) will be sufficient for all ordinary work without resorting to the more expensive "compensation" oculars. Two or three, magnifying the "real" image (formed by the objective) four, six, or eight times respectively, form a useful equipment.

As an accessory Ehrlich's Eyepiece is a very useful piece of apparatus when the enumeration of cells or bacteria has to be carried out. This is an ordinary eyepiece fitted with an adjustable square diaphragm operated by a lever projecting from the side of the mount. Three notches are made in one of the sides of the square and by moving the lever square aperture can be reduced to three-quarters, one-half or one-quarter of the original size.

10. Objectives.—Three objectives are necessary: one for low-power work—e. g., 1 inch, 2/3 inch, or 1/2 inch; one for high-power work—e. g., 1/12 inch oil immersion lens; and an intermediate "medium-power" lens—e. g., 1/6 inch or 1/8 inch (dry). These lenses must be carefully selected, especial attention being paid to the following points:

(a) Correction of Spherical Aberration.—Spherical aberration gives rise to an ill-defined image, due to the[Pg 56] central and peripheral rays focussing at different points.

(b) Correction of Chromatic Aberration.—Chromatic aberration gives rise to a coloured fringe around the edges of objects due to the fact that the different-coloured rays of the spectrum possess varying refrangibilities and that a simple lens acts toward them as a prism.

(c) Flatness of Field.—The ideal visual field would be large and, above all, flat; in other words, objects at the periphery of the field would be as distinctly "in focus" as those in the centre. Unfortunately, however, this is an optical impossibility and the field is always spherical in shape. Some makers succeed in giving a larger central area that is in focus at one time than others, and although this may theoretically cause an infinitesimal sacrifice of other qualities, it should always be sought for. Successive zones and the entire peripheral ring should come into focus with the alteration of the fine adjustment. This simultaneous sharpness of the entire circle is an indication of the perfect centering of the whole of the lenses in the objective.

Fig. 48.—Huyghenian eyepiece.

(d) Good Definition.—Actual magnification is, within limits, of course, of less value than clear definition and high resolving power, for it is upon these properties we depend for our knowledge of the detailed structure of the objects examined.

(e) Numerical Aperture (N. A.).—The numerical aperture may be defined, in general terms, as the ratio of the effective diameter of the back lens of the objective to its equivalent focal length. The determination of this point is a process requiring considerable technical[Pg 57] skill and mathematical ability, and is completely beyond the powers of the average microscopist.[1]

Although with the increase in power it is correspondingly difficult to combine all these corrections in one objective, they are brought to a high pitch of excellence in the present-day "achromatic" objectives, and so remove the necessity for the use of the higher priced and less durable apochromatic lenses.

In selecting objectives the best "test" objects to employ are:

Accessories.—Eye Shade (Fig. 49).—This piece of apparatus consists of a pear-shaped piece of blackened metal or ebonite, hinged to a collar which rotates on the upper part of the body tube of the microscope. It can be used to shut out the image of surrounding objects from the unoccupied eye, and when carrying out prolonged observations will be found of real service.

Nosepiece.—Perhaps the most useful accessory is a nosepiece to carry two of the objectives (Fig. 50), or, better still, all three (Fig. 51). This nosepiece, preferably constructed of aluminium, must be of the covered-in type, consisting of a curved plate attached to the lower end of the body tube—a circular aperture being cut to correspond to the lumen of that tube. To[Pg 58] the under surface of this plate is pivoted a similarly curved plate, fitted with three tubulures, each of which carries an objective. By rotating the lower plate each of the objectives can be brought successively in to the optical axis of the microscope.

Fig. 49.—Eye shade.

For critical work and particularly for photo-micrography, however, the interchangeable nosepiece is by no means perfect as it is next to impossible to secure accurate centreing of each lens in the optical axis. For special purposes, therefore, it is necessary to employ a special nosepiece such as that made by Zeiss or Leitz into which each objective slides on its own carrier and upon which it is accurately centred.

Fig. 50.—Double nosepiece.

Fig. 51.—Triple nosepiece.

Warm Stage (Fig. 52).—This is a flat metal case containing a system of tubes through the interior of which water of any required temperature can be circulated.[Pg 59] It is made to clamp on to the stage of the microscope by the screws A A', and is perforated with a large hole coinciding with the optical axis of the microscope; a short tube B, projecting from one end of the warm stage permits water of the desired temperature to be conducted from a reservoir through a length of rubber tubing to the interior of the stage and a similar tube at the other end B' of the stage allows exit to the waste water. By raising the temperature of hanging-drop preparations, etc., placed upon it, above that of the surrounding atmosphere, the warm stage renders possible exact observations on spore germination, hanging-drop cultivations, etc.

Fig. 52.—Warm stage.

A better form is the electrical hot stage designed by Lorrain Smith;[2] it requires the addition of a lamp resistance and sliding rheostat, also a delicate ammeter reading to .01 of an ampère. It consists of a wooden frame supporting a flat glass bulb with a long neck bent upward at an obtuse angle (Fig. 53). The bulb is filled with liquid paraffin, which rises in the open neck when expanded by heat. The neck also accommodates the thermometer. Two coils of manganin wire run in the paraffin at opposite sides of the bulb (outside the field of vision), coupled to brass terminals on the wooden frame by platinum wire fused into the glass. The resistance of the two coils in series is[Pg 60] about 10 ohms. A current of 2-1/2 ampères is needed, and is conducted to the coils in the stage through the rheostat. With the help of the ammeter any desired temperature can be obtained and maintained, up to about 200° C. If immersion oil contact is made between the top lens of the condenser and the lower surface of the bulb, this stage works very well indeed with the 1/12-inch oil immersion lens.

Fig. 53.—Lorrain Smith's warm stage.

Dark Ground or Paraboloid Condenser.—This is an immersion substage condenser of high aperture by means of which unstained objects such as bacteria can be shown as bright white particles upon a dense black background. The central rays of light are blocked out by means of an opaque stop while the peripheral rays are reflected from the paraboloidal sides of the condenser and refracted by the object viewed. To obtain the best results with this type of condenser a powerful illuminant—such as a small arc lamp or an incandescent gas lamp—is needed, together with picked slides of a certain thickness (specified for the[Pg 61] particular make of condenser but generally 1 mm.) and specially thin cover-glasses (not more than 0.17 mm.) The objective must not have a higher NA than 1.0, consequently immersion lenses must be fitted with an internal stop to cut down the aperture.

Micrometer.—Some form of micrometer for the purpose of measuring bacteria and other objects is also essential. Details of those in general use will be found in the following pages.

Fig. 54—Diamond Object marker.

Object Marker (Fig. 54).—This is an exceedingly useful piece of apparatus. Made in the form of an objective, the lenses are replaced by a diamond point, set slightly out of the centre, which can be rotated by means of a milled plate. Screwed on to the nosepiece in place of the objective, rotation of the diamond point will rule a small circle on the object slide to permanently record the position of an interesting portion of the specimen. The diamond is mounted on a spring which regulates the pressure, and the size of the circle can be adjusted by means of a lateral screw.

METHODS OF MICROMETRY.

The unit of length as applied to the measurement of microscopical objects is the one-thousandth part of a millimetre (0.001 mm.), denominated a micron (sometimes, and erroneously, referred to as a micro-millimetre), and indicated in writing by the Greek letter µ. Of the many methods in use for the measurement of bacteria, three only will be here described, viz.:

(a) By means of the Camera Lucida.

(b) By means of the ocular or Eyepiece Micrometer.

(c) By means of the Filar Micrometer (Ramsden's micrometer eyepiece).[Pg 62]

For each of these methods a stage micrometer is necessary. This is a 3 by 1 inch glass slip having engraved on it a scale divided to hundredths of a millimetre (0.01 mm.), every tenth line being made longer than the intervening ones, to facilitate counting; and from these engraved lines the measurement in every case is evaluated. A cover-glass is cemented over the scale to protect it from injury.

Fig. 55.—Camera lucida, Abbé pattern.

(a) By means of the Camera Lucida.

1. Attach a camera lucida (of the Wollaston, Beale, or Abbé pattern) (Fig. 55) to the eyepiece of the microscope.

2. Adjust the micrometer on the stage of the microscope and accurately focus the divisions.

3. Project the scale of the stage micrometer on to a piece of paper and with pen or pencil sketch in the magnified image, each division of which corresponds to 10µ. Mark on the paper the optical combination (ocular objective and tube length) employed to produce this particular magnification.[Pg 63]

4. Repeat this procedure for each of the possible combinations of oculars and objectives fitted to the microscope supplied, and carefully preserve the scales thus obtained.

To measure an object by this method simply project the image on to the scale corresponding to the particular optical combination in use at the moment. Read off the number of divisions it occupies and express them as micra.

In place of preserving a scale for each optical combination, the object to be measured and the micrometer scale may be projected and sketched, in turn, on the same piece of paper, taking particular care that the centre of the eyepiece is 25 cm. from the paper on which the divisions are drawn.

Fig. 56.—Eyepiece micrometer, ordinary.

Fig. 57.—Eyepiece micrometer, net.

(b) By means of the Eyepiece Micrometer.

The eyepiece micrometer is a circular glass disc having engraved on it a scale divided to tenths of a millimetre (0.1 mm.) (Fig. 56), or the entire surface ruled in 0.1 mm. squares (the net micrometer) (Fig. 57). It can be fitted inside the mount of any ocular just above the aperture of the diaphragm and must be adjusted exactly in the focus of the eye lens.

Some makers mount the glass disc together with a circular cover-glass in such a way that when placed in position in any Huyghenian eyepiece of their own manufacture, the scale is exactly in focus for normal[Pg 64] vision. Special eyepieces are also obtainable having a sledging adjustment to the eye lens for focussing the micrometer.

The value of one division of the micrometer scale must first be ascertained for each optical combination by the aid of the stage micrometer, thus:

1. Insert the eyepiece micrometer inside the ocular and adjust the stage micrometer on the stage of the microscope.

2. Focus the scale of the stage micrometer accurately; the lines will appear to be immediately below those of the eyepiece micrometer. Make the lines on the two micrometers parallel by rotating the ocular.

3. Make two of the lines on the ocular micrometer coincide with those bounding one division of the stage micrometer; this is effected by increasing or diminishing the tube length; and note the number of included divisions.

4. Calculate the value of each division of the eyepiece micrometer in terms of µ, by means of the following formula:

5. Note the optical combination employed in this experiment and record it with the calculated micrometer value.

Repeat this process for each of the other combinations. Carefully record the results.

To measure an object by this method read off the number of divisions of the eyepiece micrometer it occupies and express the result in micra by a reference to the standard value for the particular optical combination employed.[Pg 65]

Zeiss prepares a compensating eyepiece micrometer for use with his apochromatic objectives, the divisions of which are so computed that (with a tube length of 160 mm.) the value of each is equivalent to as many micra as there are millimetres in the focal length of the objective employed.

Wright's Eikonometer is really a modification of the eyepiece micrometer for rapidly measuring microscopical objects by direct inspection, having previously determined the magnifying power of the particular optical combination employed. It is a small piece of apparatus resembling an eyepiece, with a sliding eye lens, which can be accurately focussed on a micrometer scale fixed within the instrument. When placed over the microscope ocular the divisions of this scale measure the actual size of the virtual image in millimetres.

In order to use this instrument for direct measurement, it is first necessary to determine the magnifying power of each combination of ocular, tube length and objective.

Place a stage micrometer divided into hundredths of a millimetre on the microscope stage and focus accurately.

Rest the eikonometer on the eyepiece. Observation through the eikonometer shows its micrometer scale superposed on the image of the stage micrometer.

Rotate the eikonometer until the lines on the two scales are parallel, and make the various adjustments to ensure that two lines on the eikonometer scale coincide with two lines on the stage micrometer.

For the sake of illustration it may be assumed that five of the divisions on the stage micrometer accurately fill one of the divisions of the eikonometer scale; this indicates a magnifying power of 500 as the constant for that particular optical combination, and a record should be made of the fact.

The magnification constants of the various other[Pg 66] optical combinations should be similarly made and recorded.

To measure any object subsequently it should be first focussed carefully in the ordinary way.

The eikonometer should then be applied to the eyepiece and the size of the object read off on the eikonometer scale as millimetres, and the actual size calculated by dividing the observed size by the magnification constant for the particular optical combination employed in the observation.

(c) By means of the filar micrometer.

Fig. 58.—Ramsden's Filar micrometer.

Fig. 59.—Ramsden's micrometer field, a, fixed wire; b, reference wire (fixed); c, travelling wire.

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.[Pg 67]

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[Pg 68] 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.

Fig. 60.—Electric microscope lamp.

[Pg 69]

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.)

Fig. 61.—Disinfectant Jar.

3. Glass jar filled with 2 per cent. lysol solution for the reception of infected cover-glasses and infected pipettes, etc.[Pg 70]

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).

Fig. 62.—Hanging-drop slides: a, Double cell seen from above; b, single cell seen from the side.

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).

[Pg 71]

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.

Fig. 63.—Ends of platinum rods. a, loop; b, spatula; c, needle.

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[Pg 72] 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.

Fig. 64.—Platinum rod in aluminium handle—method of mounting. The platinum wire may be fused into the end of a piece of glass rod, but such a handle is vastly inferior to aluminium and is not to be recommended.

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

Fig. 65.—Staining rack, rubber change mat and lysol pot.

Fig. 66.—Drop bottle.

Fig. 67.—Canada balsam pot.

And two pots with air-tight glass caps (Fig. 67), each provided with a piece of glass rod and filled respectively[Pg 74] with Canada balsam dissolved in xylol, and sterile vaseline.

METHODS OF EXAMINATION.

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.

Fig. 68.—Holding tubes for removing bacterial growth, as seen from the front.

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[Pg 77] 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

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:

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.[Pg 78]

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.

Fig. 69.—Spreading negative film.

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.[Pg 79]

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[Pg 80] 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).[Pg 81]

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:[Pg 82]

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:

6. Gulland's solution, for five minutes. This solution is prepared by mixing:

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.)[Pg 83]

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.[Pg 84]

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.,

[Pg 85]

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.

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[Pg 86] 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[Pg 87] 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[Pg 88] 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.

Fig. 70.—Immersion oil bottle.

Microscopical Examination of the Stained Specimen.—(The body tube of the microscope may be vertical or inclined to an angle.)

1. Secure the slide on the stage of the microscope by means of the spring clips.

2. Place a drop of cedarwood oil on the centre of the cover-slip.

3. Use the 1/12-inch oil immersion lens of the microscope. Rack down the body tube till the front lens[Pg 89] 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.

[Pg 90]

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

Place in a stoppered bottle having a capacity of from 150 to 200 c.c. and add

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

[Pg 91]

Place in a stoppered bottle of 150 c.c. capacity and add

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

and dissolve in

and add

Filter.

4. Alkaline Methylene-blue (Loeffler).

Measure out and mix

Filter.

Gentian Violet.—

5. Saturated Aqueous Solution.

Weigh out

and proceed as in preparing the corresponding solution of methylene-blue.

6. Saturated Alcoholic Solution.

Weigh out

and proceed as in preparing the corresponding solution of methylene-blue.[Pg 92]

7. Carbolic Gentian Violet (Nicollé).

Measure out and mix

Filter.

8. Anilin Water Solution (Koch-Ehrlich).

Measure out

Add anilin oil drop by drop (shaking well after the addition of each drop) until the solution is opaque.

Filter until clear.

and add

Filter.

Thionine Blue (or Lauth's Violet).—

9. Carbolic Thionine Blue (Nicollé).

Weigh out

and dissolve in

Filter.

Before use dilute with equal quantity of distilled water and again filter.

Fuchsin (Basic).—

10. Saturated Aqueous Solution.

Weigh out

and proceed as in preparing the corresponding solution of methylene-blue (q. v.).[Pg 93]

11. Saturated Alcoholic Solution.

Weigh out

and proceed as in preparing the corresponding solution of methylene-blue.

12. Carbolic Fuchsin (Ziehl).

Weigh out

dissolve in

and add

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:

13. Eosin Aqueous Solution (Yellowish or Bluish Shade), 1 per cent.

Weigh out

dissolve in

and add

Filter.[Pg 94]

14. Eosin Alcoholic Solution, 0.5 per cent.

Weigh out

and dissolve in

Filter.

Safranine.—

15. Aqueous Solution.

Weigh out.

and dissolve in

Filter.

Neutral Red.—

16. Aqueous Solution.

Weigh out

and dissolve in

Filter.

Vesuvin (or Bismarck Brown).—

17. Saturated Aqueous Solution.

Weigh out

and dissolve in

Filter.[Pg 95]

TISSUE STAINS.

Aniline Gentian Violet (For Weigert's Fibrin Stain).—

Weigh out

and dissolve in

then add

Shake well and filter before use.

Hæmatoxylin (Ehrlich).—

1. Weigh out

and dissolve in

2. Weigh out

and dissolve in

3. Mix 1 and 2, allow the mixture to stand forty-eight hours, then filter.

4. Add

5. Allow the stain to stand for one month exposed to light; then filter again ready for use.

Hæmatin (Mayer's).—

A. Weigh out

and dissolve in

[Pg 96]

B. Weigh out

and dissolve in

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.

Alum Carmine (Mayer).—

Weigh out

and place in a glass beaker.

Measure out in a measuring cylinder,

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

and dissolve in

Filter.

Picrocarmine.—

Weigh out

[Pg 97]

and dissolve in

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:

Dissolve in

This will make a thick solution.

B. Weigh out:

Dissolve in

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° C. When a paste[Pg 98] 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:

Dissolve in

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:

Dissolve in

Keep at 65° C. for 12 hours in either a hot incubator or a water-bath; then stand in dark place at room temperature (20°C.) for ten days.

B. Weigh out:

Dissolve in

1. Mix the two solutions A and B in equal volumes,[Pg 99] 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

[Pg 100]

rub up in a mortar with

Add

and make up to 200 c.c. by the addition of

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:

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.[Pg 101]

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:

Warm to 36° 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.—

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.[Pg 102]

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.—

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°C, twenty-four to thirty hours if incubated at 22°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.[Pg 103]

9. Wash well in water.

10. Dry and mount.

[Pg 104]

with 8 c.c. distilled water, filtering and replacing the loss from evaporation.

Alternative Mordant (Bunge's Mordant).—

(b) Stain.—

Weigh out

and dissolve in

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.—

[Pg 105]

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

and dissolve in

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.[Pg 106]

3. Wash thoroughly in water.

4. Stain in hæmatoxylin 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[Pg 107] 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.

[Pg 108]

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.

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

[Pg 109]

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

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.

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.[Pg 110]

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.)[Pg 111]

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.

4. Wash in water.

5. Dry and mount.

Neisser's Method—Modified.—(To demonstrate diphtheroid bacilli.)

Stain I.—

Measure out and mix

Filter.

Stain II.—

Weigh out

[Pg 112]

and dissolve in

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.

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.

[Pg 113]

[Pg 114]

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.[Pg 115]

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.

Fig. 71.—Washing tissues.

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[Pg 116] 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.—

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.—

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.

Fig. 72.—L-shaped brass moulds.

Fig. 73.—Paraffin kettle.

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[Pg 118] 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[Pg 119] 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.

Fig. 74.—Paraffin mould.

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.[Pg 120]

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.

Fig. 75.—Section rack.

Mayer's Albumin.—

Weigh out

and dissolve in

Add

[Pg 121]

Schallibaum's solution:

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

[Pg 122]

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.

Fig. 76.—Staining pot.

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.[Pg 123]

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

and dissolve in

Measure out

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.—

[Pg 124]

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.[Pg 125]

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:

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.

[Pg 126]

VIII. CLASSIFICATION OF FUNGI.

For practical purposes Fungi may be divided into:

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.[Pg 127]

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).

Fig. 77.—Mucor mucedo.

Fig. 78.—Aspergillus

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).

Fig. 79.—Penicillium.

Note the asexual reproduction.

1. A filament (b) grows upward, its termination becomes clubbed; on the clubbed extremity flask-shaped cells appear—"sterigmata" (c).[Pg 128]

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.

Fig. 80.—Oïdium.

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).[Pg 129]

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.

Fig. 81.—Saccharomyces with ascospores.

Fig. 82.—Torula.

2. Spore formation—"ascospores" (e). These are formed at definite temperatures and within well-defined periods; e. g., Saccharomyces cerevisiæ, thirty hours at 25° to 37°C., or ten days at 12°C.[Pg 130]

Torulæ (Fig. 82).—Torulæ, 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.

[Pg 131]

IX. SCHIZOMYCETES.

Classification and Morphology.—Bacteria are often classified, in general terms, according to their life functions, into—

or according to their food requirements into—

or according to their metabolic products into—

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[Pg 132]—

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.

Fig. 83.—Types of bacteria—cocci: 1, Diagram of sphere indicating planes of fission; 2, diplococci; 3, streptococci; 4, tetrads; 5, sarcinæ; 6, staphylococci.

Sarcinæ, where division takes place in three planes successively, and the individuals remain attached in cubical packets of eight and its multiples.

Fig. 84.—Types of bacteria—bacilli, etc.: 1, Bacilli; 2, diplobacilli; 3 streptobacilli; 4, spirilla; 5, vibrios; 6, spirochætæ.

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.

2. Bacilli (Fig. 84, 1 to 3).—Rod-shaped cells. A bacillus, however short, can usually be distinguished[Pg 133] from a coccus in that two sides are parallel. Some bacilli after fission retain a characteristic arrangement and may be spoken of as Diplobacilli or Streptobacilli.

Leptothrix is a term that in the past has been loosely used to signify a long thread, but is now restricted to such forms as belong to the leptothriciæ (vide infra).

3. Spirilla (Fig. 84, 4 to 6).—Curved and twisted filaments. Classified, according to shape, into—

Many Spirochætes appear to belong to the animal kingdom and are grouped under protozoa; other organisms to which this name has been given are undoubtedly bacteria.

Higher forms of bacteria are also met with, which possess the following characteristics: They are attached, unbranched, filamentous forms, showing—

(a) Differentiation between base and apex;

(b) Growth apparently apical;

(c) Exaggerated pleomorphism;

(d) "Pseudo-branching" from apposition of cells; and are classified into—

The morphology of the same bacterium may vary greatly under different conditions.

For example, under one set of conditions the examination of a pure cultivation of a bacillus may show a short oval rod as the predominant form, whilst another[Pg 134] culture of the same bacillus, but grown under different conditions, may consist almost entirely of long filaments or threads. This variation in morphology is known as "pleomorphism."

Some of the factors influencing pleomorphism are:

1. The composition, reaction, etc., of the nutrient medium in which the organism is growing.

2. The atmosphere in which it is cultivated.

3. The temperature at which it is incubated.

4. Exposure to or protection from light.

The various points in the anatomy morphology and physiology of bacteria upon which stress is laid in the following pages should be studied as closely as is possible in preparations of the micro-organisms named in connection with each.

ANATOMY.

1. Capsule (Fig. 85, b).—A gelatinous envelope (probably akin to mucin in composition) surrounding each individual organism, and preventing absolute contact between any two. In some species the capsule (e. g., B. pneumoniæ) is well marked, but it cannot be demonstrated in all. In very well marked cases of gelatinisation of the cell wall, the individual cells are cemented together in a coherent mass, to which the term "zooglœa" is applied (e. g., Streptococcus mesenteroides). In some species colouring matter or ferric oxide is stored in the capsule.

2. Cell Wall (Fig. 85, c).—A protective differentiation of the outer layer of the cell protoplasm; difficult to demonstrate, but treatment with iodine or salt solution sometimes causes shrinkage of the cell contents—"plasmolysis"—and so renders the cell wall apparent (e. g., B. megatherium) in the manner shown in figure 85. Stained bacilli, when examined with the polarising microscope, often show a doubly[Pg 135] refractile cell wall (e. g., B. tuberculosis and B. anthracis).

In some of the higher bacteria the cell wall exhibits this differentiation to a marked degree and forms a hard sheath within which the cell protoplasm is freely movable; and during the process of reproduction the cell protoplasm may be extruded, leaving the empty tube unaltered in shape.

Fig. 85.—Dragrammatic sketch of composite bacterium to illustrate details of anatomical structure.

Fig. 86.—Plasmolysis.

3. Cell Contents.—Protoplasm (mycoprotein) contains a high percentage of nitrogen, but is said to differ from proteid in that it is not precipitated by C₂H₆O. It is usually homogeneous in appearance—sometimes granular—and may contain oil globules or sap vacuoles (Fig. 85, d), chromatin granules, and even sulphur granules. Sap vacuoles must be distinguished from spores, on the one hand, and the vacuolated appearance due to plasmolysis, on the other.

The cell contents may sometimes be differentiated into a parietal layer, and a central body (e. g., beggiotoa) when stained by hæmatoxylin.

4. Nucleus.—This structure has not been conclusively[Pg 136] proved to exist, but in some bacteria chromatin particles have been observed near the centre of the bacterial cell and denser masses of protoplasm situated at the poles which exhibit a more marked affinity than the rest of the cell protoplasm for aniline dyes. These latter are termed polar granules or Polkoerner (Fig. 85, e). Occasionally these aggregations of protoplasm alter the colour of the dye they take up. They are then known as metachromatic bodies or Ernstschen Koerner (e. g., B. diphtheriæ).

5. Flagella (Organs of Locomotion, Fig. 85, a).—These are gelatinous elongations of the cell protoplasm (or more probably of the capsule), occurring either at one pole, at both poles, or scattered around the entire periphery. Flagella are not pseudopodia. The possession of flagella was at one time suggested as a basis for a system of classification, when the following types of ciliation were differentiated (Fig. 87):

Fig. 87.—Types of ciliation.

1. Polar: (a) Monotrichous (a single flagellum situated at one pole; e. g., B. pyocyaneus).

(b) Amphitrichous (a single flagellum at each pole; e. g., Spirillum volutans).

(c) Lophotrichous (a tuft or bunch of flagella situated at each pole; e. g., B. cyanogenus).

2. Diffuse: Peritrichous (flagella scattered around the entire periphery e. g., B. typhosus).

PHYSIOLOGY.

Reproduction.—Active Stage.—Vegetative, i. e., by the division of cells, or "fission."

1. The cell becomes elongated and the protoplasm aggregated at opposite poles.

2. A circular constriction of the organism takes[Pg 137] place midway between these aggregations, and a septum is formed in the interior of the cell at right angles to its length.

3. The division deepens, the septum divides into two lamellæ, and finally two cells are formed.

Fig. 88.—Fission o£ cocci.

Fig. 89.—Fission of bacteria.

4. The daughter cells may remain united by the gelatinous envelope for a variable time. Eventually they separate and themselves subdivide.

Cultures on artificial media, after growing in the same medium for some time—i. e., when the pabulum is exhausted—show "involution forms" (Fig. 90), well exemplified in cultures of B. pestis on agar two days old, B. diphtheriæ on potato four to six days old.

Fig. 90.—Involution forms.

They are of two classes, viz.:

(a) Involution forms characterised by alterations of shape (Fig. 90). (Not necessarily dead.)

(b) Involution forms characterised by loss of staining power. (Always dead.)

Resting Stage.—Spore Formation.—Conditions influencing spore formation: In an old culture nothing may be left but spores. It used to be supposed that spores were always formed, so that the species might not become extinct, when

(a) The supply of nutrient was exhausted.[Pg 138]

(b) The medium became toxic from the accumulation of metabolic products.

(c) The environment became unfavourable; e. g., change of temperature.

This is not altogether correct; e. g., the temperature at which spores are best formed is constant for each bacterium, but varies with different species; again, aerobes require oxygen for sporulation, but anaerobes will not spore in its presence.

(A) Arthrogenous: Noted only in the micrococci. One complete element resulting from ordinary fission becomes differentiated for the purpose, enlarges, and develops a dense cell wall. One or more of the cells in a series may undergo this alteration.

This process is probably not real spore formation, but merely relative increase of resistance. These so-called arthrospores have never been observed to "germinate," nor is their resistance very marked, as they fail to initiate new cultures, after having been exposed to a temperature of 80° C. for ten minutes.

(B) Endogenous: The cell protoplasm becomes differentiated and condensed into a spherical or oval mass (very rarely cylindrical). After further contraction the outer layers of the mass become still more highly differentiated and form a distinct spore membrane, and the spore itself is now highly refractile. It has been suggested, and apparently on good grounds, that the spore membrane consists of two layers, the exosporium and the endosporium. Each cell forms one spore only, usually in the middle, occasionally at one end (some exceptions, however, are recorded; e. g., B. inflatus). The shape of the parent cell may be unaltered, as in the anthrax bacillus, or altered, as in the tetanus bacillus, and these points serve as the basis for a classification of spore-bearing bacilli, as follows:

(A) Cell body of the parent bacillus unaltered in shape (Fig. 91, a).[Pg 139]

(B) Cell of the parent bacillus altered in shape.

1. Clostridium (Fig. 91, b): Rod swollen at the centre and attenuated at the poles; spindle shape; e. g., B. butyricus.

2. Cuneate (Fig. 91, c): Rods swollen slightly at one pole and more or less pointed at the other; wedge-shaped.

Fig. 91—Types of spore-bearing bacilli.

3. Clavate (Fig. 91, d): Rods swollen at one pole and cylindrical (unaltered) at the other; keyhole-shaped; e. g., B. chauvei.

4. Capitate (Fig. 91, e): Rods with a spherical enlargement at one pole; drumstick-shaped; e. g., B. tetani.

The endo-spores remain within the parent cell for a variable time (in one case it is stated that germination of the spore occurs within the interior of the parent cell—"endo-germination"), but are eventually set free, as a result of the swelling up and solution of the cell membrane of the parent bacillus in the surrounding liquid, or of the rupture of that membrane. They then present the following characteristics:

1. Well-formed, dense cell membranes, which renders them extremely difficult to stain, but when once stained equally difficult to decolourise.

2. High refractility, which distinguished them from vacuoles.

3. Higher resistance than the parent organism to such lethal agents as heat, desiccation, starvation, time, etc., this resistance being due to

(a) Low water contents of plasma of the spore.

This resistance varies somewhat with the particular species—e. g., some spores may resist boiling for a few[Pg 140] minutes—but practically all are killed if the boiling is continued for ten minutes.

Germination.—When transplanted to suitable media and placed under favourable conditions, the spores germinate, usually within twenty-four to thirty-six hours, and successively undergo the following changes which may be followed in hanging-drop cultures on a warm stage:

1. Swell up slowly and enlarge, through the absorption of water.

2. Lose their refrangibility.

3. At this stage one of three processes (but the particular process is always constant for the same species) may be observed:

(a) The spore grows out into the new bacillus without discarding the spore membrane (which in this case now becomes the cell membrane); e. g., B. leptosporus.

(b) It loses its spore membrane by solution; e. g., B. anthracis.

(c) It loses its spore membrane by rupture.

In this process the rupture may be either polar (at one pole only e. g., B. butyricus), or bipolar (e. g., B. sessile), or equatorial; (e. g., B. subtilis).

In those cases where the spore membrane is discarded the cell membrane of the new bacillus may either be formed from—

(a) The inner layer of the spore membrane, which has undergone a preliminary splitting into parietal and visceral layers; e. g., B. butyricus.

(b) The outer layers of the cell protoplasm, which become differentiated for that purpose; e. g., B. megatherium.

The new bacillus now increases in size, elongates, and takes on a vegetative growth—i. e., undergoes fission—the bacilli resulting from which may in their turn give rise to spores.[Pg 141]

Fig. 92. Simple.

Fig. 93. Solution.

Fig. 94. Polar.

Fig. 95. Bipolar.

Fig. 96. Equatorial.

[Pg 142]

Food Stuffs.—1. Organic Foods.—

(a) The pure parasites (e. g., B. lepræ) will not live outside the living body.

(b) Both saprophytic and facultative parasitic bacteria agree in requiring non-concentrated food.

(c) The facultative parasites need highly organised foods; e. g., proteids or other sources of nitrogen and carbon, and salts.

(d) The saprophytic bacteria are more easily cultivated; e. g.,

1. Some bacteria will grow in almost pure distilled water.

2. Some bacteria will grow in pure solutions of the carbohydrates.

3. Water is absolutely essential to the growth of bacteria.

Food of a definite reaction is needed for the growth of bacteria. As a general rule growth is most active in media which react slightly acid to phenolphthalein—that is, neutral or faintly alkaline to litmus. Mould growth, on the other hand, is most vigourous in media that are strongly acid to phenolphthalein.

Environment.—The influence of physical agents upon bacterial life and growth is strongly marked.

1. Atmosphere.—The presence of oxygen is necessary for the growth of some bacteria, and death follows when the supply is cut off. Such organisms are termed obligate aerobes.

Some bacteria appear to thrive equally well whether supplied with or deprived of oxygen. These are termed facultative anaerobes.

A third class will only live and multiply when the access of free oxygen is completely excluded. These are termed obligate anaerobes.

2. Temperature.—Practically no bacterial growth occurs below 5°C, and very little above 40° C. 30°C.[Pg 143] to 37° C is the most favorable for the large majority of micro-organisms.

The maximum and minimum temperatures at which growth takes place, as well as the optimum, are fairly constant for each bacterium.

Bacteria have been classified, according to their optimum temperature, into—

The thermal death-point of an organism is another biological constant; and is that temperature which causes the death of the vegetative forms when the exposure is continued for a period of ten minutes (see pages 298-301).

3. Light.—Many organisms are indifferent to the presence of light. On the other hand, light frequently impedes growth, and alters to a greater or lesser extent the biochemical characters of the organisms—e. g., chromogenicity or power of liquefaction. Pathogenic bacteria undergo a progressive loss of virulence when cultivated in the presence of light.

4. Movements.—Movements, if slight and simply of a flowing character, do not appear to injuriously affect the growth of bacteria; but violent agitation, such as shaking, absolutely kills them.

A condition of perfect rest would seem to be that most conducive to bacterial growth.

The Metabolic Products of Bacteria.—Pigment Production.—Many micro-organisms produce one or more vivid pigments—yellow, orange, red, violet, fluorescent, etc.—during the course of their life and growth. The colouring matter usually exists as an intercellular excrementitious substance. Occasionally, however, it[Pg 144] appears to be stored actually within the bodies of the bacteria. The chromogenic bacteria are therefore classified, in accordance with the final destination of the colouring matter they elaborate, into—

Chromoparous Bacteria: in which the pigment is diffused out upon and into the surrounding medium.

Chromophorous Bacteria: in which the pigment is stored in the cell protoplasm of the organism.

Parachromophorous Bacteria: in which the pigment is stored in the cell wall of the organism.

Different species of chromogenic bacteria differ in their requirements as to environment, for the production of their characteristic pigments; e. g., some need oxygen, light, or high temperature; others again favor the converse of these conditions.

Light Production.—Some bacteria, and usually those originally derived from water, whether fresh or salt, exhibit marked phosphorescence when cultivated under suitable conditions. These are classed as "photogenic."

Enzyme Production.—Many bacteria produce soluble ferments or enzymes during the course of their growth, as evidenced by the liquefaction of gelatine, the clotting of milk, etc. These ferments may belong to either of the following well-recognised classes: proteolytic, diastatic, invertin, rennet.

Toxin Production.—A large number, especially of the pathogenic bacteria, elaborate or secrete poisonous substances concerning which but little exact knowledge is available, although many would appear to be enzymic in their action.

These toxins are usually differentiated into—

Extracellular (or Soluble) Toxins: those which are diffused into, and held in solution by, the surrounding medium.

Intracellular (or Inseparate) Toxins: those which are so closely bound up with the cell protoplasm of the bacteria elaborating them that up to the present time[Pg 145] no means has been devised for their separation or extraction.

End-products of Metabolism.—Under this heading are included—

Organic Acids (e. g., lactic, butyric, etc.).

Alkalies (e. g., ammonia).

Aromatic Compounds (e. g., indol, phenol).

Reducing Substances (e. g., those reducing nitrates to nitrites).

Gases (e. g., sulphuretted hydrogen, carbon dioxide, etc.).

And while the discussion of their formation, etc., is beyond the scope of a laboratory handbook, the methods in use for their detection and separation come into the ordinary routine work and will therefore be described (vide page 276 et seq.).

[Pg 146]

X. NUTRIENT MEDIA.

In order that the life and growth of bacteria may be accurately observed in the laboratory, it is necessary—

1. To isolate individual members of the different varieties of micro-organisms.

2. To cultivate organisms, thus isolated, apart from other associated or contaminating bacteria—i. e., in pure culture.

For the successful achievement of these objects it is necessary to provide nutriment in a form suited to the needs of the particular bacterium or bacteria under observation, and in a general way it may be said that the nutrient materials should approximate as closely as possible, in composition and character, to the natural pabulum of the organism.

The general requirements of bacteria as to their food-supply have already been indicated (page 142) and many combinations of proteid and of carbohydrate have been devised, from time to time, on those lines. These, together with various vegetable tissues, physiological or pathological fluid secretions, etc., are collectively spoken of as nutrient media or culture media.

The greater number of these media are primarily fluid, but, on account of the rapidity with which bacterial growth diffuses itself through a liquid, it is impossible to study therein the characteristics of individual organisms. Many such media are, therefore, subsequently rendered solid by the addition of substances like gelatine or agar, in varying proportions, the proportions of such added material being generally mentioned when referring to the media; e. g., 10 per cent. gelatine, 2 per cent. agar. Gelatine is employed[Pg 147] for the solidification of those media it is intended to use in the cultivation of bacteria at the room temperature or in the "cold" incubator. In the percentages usually employed, gelatine media become fluid at 25°C.; higher percentages remain solid at somewhat higher temperatures, but the difficulty of filtering strong solutions of gelatine militates against their general use.

Media, on the other hand which have been solidified by the addition of agar, only become liquid when exposed to 90° C. for about ten minutes, and again solidify when the temperature falls to 40°C.

When it becomes necessary to render these media fluid, heat is applied, upon the withdrawal of which they again assume their solid condition. Such media should be referred to as liquefiable media; in point of fact, however, they are usually grouped together with the solid media.

Other media, again—e. g., potato, coagulated blood-serum, etc.—cannot be again liquefied by physical means, and these are spoken of as solid media.

The following pages detail the method of preparing the various nutrient media, in ordinary use (see also Chapter XI), those which are only occasionally required for more highly specialised work are grouped together in Chapter XII. It must be premised that scrupulous cleanliness is to be observed with regard to all apparatus, vessels, funnels, etc., employed in the preparation of media; although in the preliminary stages of[Pg 148] the preparation of most media absolute sterility of the apparatus used is not essential.

MEAT EXTRACT.

A watery solution of the extractives, etc., of lean meat (usually beef) forms the basis of several nutrient media. This solution is termed "meat extract" and it has been determined empirically that its preparation shall be carried out by extracting half a kilo of moist meat with one litre of water. For many purposes, however, it is more convenient to have a more concentrated extract; one kilo of meat should therefore be extracted with one litre of water, to form "Double Strength" meat extract.

It was customary at one time, and is even now in some laboratories to use either "shin of beef" or "beef-steak"—both contain muscle sugar which often needs to be removed before the nutrient medium can be completed. Heart muscle (bullock's heart or sheep's heart) is much to be preferred and from the point of economy, ease and cleanliness of manipulation, and extractive value, the imported frozen bullock's hearts provide the best extract.

Meat extract (Fleischwasser) is prepared as follows:

1. Measure 1000 c.c. of distilled water into a large flask (or glass beaker, or enamelled iron pot) and add 1000 grammes (roughly, 2-1/2 pounds) of fresh lean meat—e. g., bullock's heart—finely minced in a mincing machine.

2. Heat the mixture gently in a water-bath, taking care that the temperature of the contents of the flask does not exceed 40° C. for the first twenty minutes. (This dissolves out the soluble proteids, extractives, salts, etc.)

3. Now raise the temperature of the mixture to the boiling-point, and maintain at this temperature for[Pg 149] ten minutes. (This precipitates some of the albumins, the hæmoglobin, etc., from the solution.)

4. Strain the mixture through sterile butter muslin or a perforated porcelain funnel, then filter the liquid through Swedish filter paper into a sterile "normal" litre flask, and when cold make up to 1000 c.c. by the addition of distilled water—to replace the loss from evaporation.

5. If not needed at once, sterilise the meat extract in bulk in the steam steriliser for twenty minutes on each of three consecutive days.

Calf, sheep, or chicken flesh is occasionally substituted for the beef; or the meat extract may be prepared from animal viscera, such as brain, spleen, liver, or kidneys.

Reaction of Meat Extract.—Meat extract thus prepared is acid in its reaction, owing to the presence of acid phosphates of potassium and sodium, weak acids of the glycolic series, and organic compounds in which the acid character predominates. Owing to the nature of the substances from which it derives its reaction, the total acidity of meat extract can only be estimated accurately when the solution is at the boiling-point.

Moreover, it has been observed that prolonged boiling (such as is involved in the preparation of nutrient media) causes it to undergo hydrolytic changes which increase its acidity, and the meat extract only becomes stable in this respect after it has been maintained at the boiling-point for forty-five minutes.[Pg 150]

Although meat extract always reacts acid to phenolphthalein, it occasionally reacts neutral or even alkaline to litmus; and again, meat extract that has been rendered exactly neutral to litmus still reacts acid to phenolphthalein. This peculiar behaviour depends upon two factors:

1. Litmus is insensitive to many weak organic acids the presence of which is readily indicated by phenolphthalein.

2. Dibasic sodium phosphate which is formed during the process of neutralisation is a salt which reacts alkaline to litmus, but neutral to phenolphthalein. In order, therefore, to obtain an accurate estimation of the reaction of any given sample of meat extract, it is essential that—

1. The meat extract be previously exposed to a temperature of 100° C. for forty-five minutes.

2. The estimation be performed at the boiling-point.

3. Phenolphthalein be used as the indicator.

The estimation is carried out by means of titration experiments against standard solutions of caustic soda, in the following manner:

Method of Estimating the Reaction.—

Method.—Arrange the apparatus as indicated in figure 97.

(A) 1. Fill the burette with n/10 NaOH.

2. Fill the pipette with n/1 NaOH.

Fig. 97.—Arrangement of apparatus for titrating media.

3. Measure 25 c.c. of the meat extract (previously heated in the steamer at 100° C. for forty-five minutes) into one of the beakers by means of the measure; rinse out the measure with a very small quantity of boiling distilled water from the wash-bottle, and then add this rinse water to the meat extract already in the beaker.

4. Run in about 0.5 c.c. of the phenolphthalein solution and immerse the beaker in the water-bath, and raise to the boil.

5. To the medium in the beaker run in n/10 NaOH cautiously from the burette until the end-point is reached, as indicated by the development of a pinkish[Pg 152] tinge, shown in figure 98 (b). Note the amount of decinormal soda solution used in the process.

Fig. 98.—a, Sample of filtered meat extract or nutrient gelatine to which phenolphthalein has been added. The medium is acid, as evidenced by the unaltered colour of the sample. b, The same neutralised by the addition of n/10 NaOH. The production of this faint rose-pink colour indicates that the "end-point," or neutral point to phenolphthalein, has been reached. If such a sample is cooled down to say 30° or 20° C., the colour will be found to become more distinct and decidedly deeper and brighter, resembling that shown in c. c, Also if, after the end-point is reached, a further 0.5 c.c. or 1.0 c.c. n/10 NaOH be added to the sample, the marked alkalinity is evidenced by the deep colour here shown.

(B) Perform a "control" titration (occasionally two controls may be necessary), as follows:

1. Measure 25 c.c. of the meat extract into one of the beakers, wash out the measure with boiling water, and add the phenolphthalein as in the first estimation.

2. Run in n/1 NaOH from the pipette, just short of the equivalent of the amount of deci-normal soda solution required to neutralise the 25 c.c. of medium. (For example, if in the first estimation 5 c.c. of n/10 NaOH were required to render 25 c.c. of medium neutral to phenolphthalein, only add 0.48 c.c. of n/1 NaOH.) Immerse the beaker in the water-bath.

3. Complete the titration by the aid of the n/10 NaOH.[Pg 153]

4. Note the amount of n/10 NaOH solution required to complete the titration, and add it to the equivalent of the n/1 NaOH solution previously run in. Take the total as the correct estimation.

Method of Expressing the Reaction.—

The reaction or titre of meat extract, medium, or any solution estimated in the foregoing manner, is most conveniently expressed by indicating the number of cubic centimetres of normal alkali (or normal acid) that would be required to render one litre of the solution exactly neutral to phenolphthalein.

Fig. 99.—Stock bottle for dekanormal soda solution.

The sign + (plus) is prefixed to this number if the original solution reacts acid, and the sign - (minus) if it reacts alkaline.[Pg 154]

For example, "meat extract + 10," indicates a sample of meat extract which reacts acid to phenolphthalein, and would require the addition of 10 c.c. of normal NaOH per litre, to neutralise it.

Conversely, if as the result of our titration experiments we find that 25 c.c. of meat extract require the addition of 5 c.c. n/10 NaOH to neutralise, then 1000 c.c. of meat extract will require the addition of 200 c.c. n/10 NaOH = 20 c.c. n/1 NaOH.

And this last figure, 20, preceded by the sign + (i. e., +20), to signify that it is acid, indicates the reaction of the meat extract.

STANDARDISATION OF MEDIA.

Differences in the reaction of the medium in which it is grown will provoke not only differences in the rate of growth of any given bacterium, but also well-marked differences in its cultural and morphological characters; and nearly every organism will be found to affect a definite "optimum reaction"—a point to be carefully determined for each. For most bacteria, however, the "optimum" usually approximates fairly closely to +10; and as experiment has shown that this reaction is the most generally useful for routine laboratory work, it is the one which may be adopted as the standard for all nutrient media derived from meat extract.[Pg 155]

Briefly, the method of standardising a litre of media to +10 consists in subtracting 10 from the initial titre of the medium mass; the remainder indicates the number of cubic centimetres of normal soda solution that must be added to the medium, per litre, to render the reaction +10.

Standardising Nutrient Bouillon.—For example, 1000 c.c. bouillon are prepared; at the first titration it is found

1. 25 c.c. require the addition of 5.50 c.c. n/10 NaOH to neutralise.

Two controls give the following results:

2. 25 c.c. require the addition of 5.70 c.c. n/10 NaOH to neutralise.

3. 25 c.c. require the addition of 5.60 c.c. n/10 NaOH to neutralise.

Averaging these two controls, 25 c.c. require the addition of 5.65 c.c. n/10 NaOH to neutralise, and therefore 1000 c.c. require the addition of 226 c.c. n/10 NaOH, or 22.60 c.c. n/1 NaOH, or 2.26 c.c. n/10 NaOH.

Initial titre of the bouillon = +22.6, and as such requires the addition of (22.6 c.c. - 10 c.c.) = 12.6 c.c. of n/1 NaOH per litre to leave its finished reaction +10.

But the three titrations, each on 25 c.c. of medium, have reduced the original bulk of bouillon to (1000 - 75 c.c.) = 925 c.c. The amount of n/1 NaOH required to render the reaction of this quantity of medium +10 may be deduced thus:

Then x = 11.65 c.c. n/1 NaOH.

Whenever possible, however, the required reaction is produced by the addition of dekanormal soda solution, on account of the minute increase it causes in the bulk, and the consequent insignificant disturbance of the percentage composition of the medium. By means of a pipette graduated to 0.01 c.c. it is possible to deliver[Pg 156] very small quantities; but if the calculated amount runs into thousandth parts of a cubic centimetre, these are replaced by corresponding quantities of normal or even decinormal soda.

In the above example it is necessary to add 11.65 c.c. normal NaOH or its equivalent, 1.165 c.c. dekanormal NaOH. The first being too bulky a quantity, and the second inconveniently small for exact measurement, the total weight of soda is obtained by substituting 1.16 c.c. dekanormal soda solution, and either 0.05 c.c. of normal soda solution or 0.5 c.c. of decinormal soda solution.

Standardising Nutrient Agar and Gelatine.—The method of standardising agar and gelatine is precisely similar to that described under bouillon.

THE FILTRATION OF MEDIA.

Fluid media are usually filtered through stout Swedish filter paper (occasionally through a porcelain filter candle), and in order to accelerate the rate of filtration the filter paper should be folded in that form which is known as the "physiological filter," not in the ordinary "quadrant" shape, as by this means a large surface is available for filtration and a smaller area in contact with the glass funnel supporting it.

To fold the filter proceed thus:

1. Take a circular piece of filter paper and fold it exactly through its centre to form a semicircle (Fig. 100, a).

2. Fold the semicircle exactly in half to form a quadrant; make the crease 2, distinct by running the thumbnail along it, then open the filter out to a semicircle again.

3. Fold each end of the semicircle in to the centre and so form another quadrant; smooth down the two new creases 3 and 3a, thus formed and again open out to a semicircle.[Pg 157]

4. The semicircle now appears as in figure 100, a, the dark lines indicating the creases already formed.

5. Fold the point 1 over to the point 3, and 1a to 3a, to form the creases 4 and 4a, indicated in the diagram by the light lines. Fold point 1 over to 3a, and 1a to 3, to form the creases 5 and 5a.

Fig. 100.—Filter folding: a, Filter folded in half, showing creases; b, appearance of filter on completion of folding; c, filter opened out ready for use.

6. Thus far the creases have all been made on the same side of the paper. Now subdivide each of the eight sectors by a crease through its centre on the opposite side of the paper, indicated by the faint broken lines in the diagram. Fold up the filter gradually as each crease is made, and when finished the filter has assumed the shape of a wedge, as in figure 100, b.[Pg 158]

When opened out the filter assumes the shape represented in figure 100, c.

The folded filter is next placed inside a glass funnel supported on a retort stand, and moistened with hot distilled water before the filtration of the medium is commenced.

Liquefiable solid media are filtered through a specially made filter paper—"papier Chardin"—which is sold in boxes of twenty-five ready-folded filters.

Fig. 101.—Hot-water filter funnel and ring burner.

Gelatine, when properly made, filters through this paper as quickly as bouillon does through the Swedish filter paper, and does not require the use of the hot-water funnel.

Agar, likewise, if properly made, filters readily, although not at so rapid a rate as gelatine. If badly "egged," and also during the winter months, it is necessary to surround the glass funnel, in which the filtration of the agar is carried on, by a hot-water jacket. This is done by placing the glass funnel inside a double-walled copper funnel—the space between the[Pg 159] walls being filled with water at about 90° C.—and supporting the latter on a ring gas burner fixed to a retort stand (Fig. 101). The gas is lighted and the water jacket maintained at a high temperature until filtration is completed. If the steam steriliser of the laboratory is sufficiently large, it is sometimes more convenient to place the flask and filtering funnel bodily inside, close the steriliser and allow filtration to proceed in an atmosphere of live steam, than to use the gas ring and hot-water funnel.

STORING MEDIA IN BULK.

After filtration fill the medium into sterile litre flasks with cotton-wool plugs and sterilise in the steamer for twenty minutes on each of three consecutive days. After the third sterilisation, and when the flasks and contents are cool, cut off the top of the cotton-wool plug square with the mouth of the flask; push the plug a short distance down into the neck of the flask and fill in with melted paraffin wax to the level of the mouth. When the wax has set the flasks are stored in a cool dark cupboard for future use.

Fig. 102.—Rubber cap closing store bottle. a, before, and b, after sterilizing.

This plan is not absolutely satisfactory, although very generally employed on occasion, and it is preferable to fill the medium into long-necked flint glass bottles (the quart size, holding nearly 1000 c.c., such as those in which Pasteurised milk is retailed) and to close the neck of the bottle by a special rubber cap.[3] This cap is made of soft rubber, the lower part, dome-shaped with thin walls, being slipped over the neck of the bottle (Fig. 102, a). The upper part is solid,[Pg 160] but with a sharp clean-cut (made with a cataract or tenotomy knife) running completely through its axis from the centre of the disc to the top of the dome. During sterilisation the air in the neck of the bottle, expanded by the heat, is driven out through the valvular aperture in the solid portion of the stopper. On removing the bottle from the steam chamber, the liquid contracts as it cools, and the pressure of the external air drives the solid piece of rubber down into the neck of the bottle, and forces together the lips of the slit (Fig. 102, b). Thus sealed, the bottle will preserve its contents sterile for an indefinite period without loss from evaporation.

TUBING NUTRIENT MEDIA.

After the final filtration, the nutrient medium is usually "tubed"—i. e., filled into sterile tubes in definite measured quantities, usually 10 c.c. This process is sometimes carried out by means of a large separator funnel fitted with a "three-way" tap which communicates with a small graduated tube (capacity 20 c.c. and graduated in cubic centimetres) attached to the side. The shape of this piece of apparatus, known as Treskow's funnel, renders it particularly liable to damage. It is better, therefore, to arrange a less expensive piece of apparatus which will serve the purpose equally well (Fig. 103).

A Geissler's three-way stop-cock has the tube on one side of the tap ground obliquely at its extremity, and the tube on the opposite side cut off within 3 cm. of the tap. The short tube is connected by means of a perforated rubber cork with a 10 cm. length of stout glass tubing (1.5 cm. bore). The third channel of the three-way tap is connected, by means of rubber tubing, with the nozzle of an ordinary separator funnel. Finally, the receiving cylinder above the three-way tap is graduated[Pg 161] in cubic centimetres up to 20, by pouring into it measured quantities of water and marking the various levels on the outside with a writing diamond.

Fluid media containing carbohydrates are filled into fermentation tubes (vide Fig. 21); or into ordinary media tubes which already have smaller tubes, inverted, inside them (Fig. 104), to collect the products of growth of gas-forming bacteria. When first filled, the small tubes float on the surface of the medium after the first sterilisation nearly all the air is replaced by the medium, and after the final sterilisation the gas tubes will be submerged and completely filled with the medium.

Fig. 103.—Separatory funnel and three-way tap arranged for tubing media.

Fig. 104.—Gas tube (Durham).

Storing "Tubed" Media.—Media after being tubed are best stored by packing, in the vertical position, in oblong boxes having an internal measurement of 37 cm. long by 12 cm. wide by 10 cm. deep. Each box (Fig. 105) has a movable partition formed by the[Pg 162] vertical face of a weighted triangular block of wood, sliding free on the bottom (Fig. 105, A); or by a flat piece of wood sliding in a metal groove in the bottom of the box, which can be fixed at any spot by tightening the thumbscrew of a brass guide rod which transfixes the partition (Fig. 105, B). The front of the box is provided with a handle and a celluloid label for the name of the contained medium. These boxes are arranged upon shelves in a dark cupboard—or preferably an iron safe—which should be rendered as nearly air-tight as possible, and should have the words "media stores" painted on its doors.

Fig. 105.—Medium box, showing alternative partitions A and B.

[Pg 163]

XI. CULTURE MEDIA.

ORDINARY OR STOCK MEDIA.

Nutrient Bouillon.—

1. Measure out double strength meat extract, 500 c.c., into a litre flask and add 300 c.c. distilled water.

2. Weigh out Witté's peptone, 10 grammes (= 1 per cent.), salt, 5 grammes (= 0.5 per cent.), and mix into a smooth paste with 200 c.c. of distilled water previously heated to 60° C. (Be careful to leave no unbroken globular masses of peptone.)

3. Add the peptone emulsion to the meat extract in the flask and heat in the steamer for forty-five minutes (to completely dissolve the peptone, and to render the acidity of the meat extract stable).

4. Estimate the reaction of the medium; control the result; render the reaction of the finished medium +10 (vide page 155).

5. Heat for half an hour in the steamer at 100°C. (to complete the precipitation of the phosphates, etc.).

6. Filter through Swedish filter paper into a sterile flask.

7. Fill into sterile tubes (10 c.c. in each tube).

8. Sterilise in the steamer for twenty minutes on each of three consecutive days—i. e., by the discontinuous method (vide page 35).

Nutrient Gelatine.—

1. Weigh a 2-litre flask on a trip balance (Fig. 106) and note the weight, or counterpoise carefully.

Fig. 106.—Trip balance.

An extremely useful counterpoise is a small sheet-brass cylinder about 38 mm. high and 38 mm. in diameter, with a funnel-shaped top and provided with a side tube by which its contents, fine "dust" shot, may be emptied out (Fig. 107).

Fig. 107.—Counterpoise; weight when empty, 35 grammes; when full of dust shot, 200 grammes.

[Pg 165]

2. Measure out double strength meat extract, 500 c.c., into the "tared" flask.

3. Weigh out and mix 10 grammes of peptone, 5 grammes of salt, and make into a thick paste with 150 c.c. distilled water; then add the emulsion to the meat extract in the flask; also add 100 grammes sheet gelatine cut into small pieces; place the flask in the water-bath and raise to the boil.

Fig. 108.—Arrangement of steam can and water-bath for the preparation of media.

4. Arrange a 5-litre tin can (with copper bottom, such as is used in the preparation of distilled water) by the side of the water bath, fill the can with boiling water and place a lighted Bunsen burner under it. Fit a long safety tube to the neck of the can and also a delivery tube, bent twice at right angles; adjust the tube to reach to the bottom of the interior of the flask containing the gelatine, etc. (Fig. 108).[Pg 166]

5. Keep the water in the steam can vigourously boiling, and so steam at 100°C, bubbling through the medium mass, for ten minutes, by which time complete solution of the gelatine is effected. A certain amount of steam will condense as water in the medium flask during this process—hence the necessity for the use of double strength meat extract—but if the water bath is kept boiling this condensation will not exceed 100 c.c.

6. Weigh the flask and its contents; then (1115[4] grammes + weight of the flask) minus (weight of the flask and its contents) equals the weight of water required to make up the bulk to 1 litre. The addition of the requisite quantity of water is carried out as follows:

In one pan of the trip balance place the counterpoise of the tared flask (or its equivalent in weights) together with the weights making up the calculated medium weight. In the opposite pan place the flask containing the medium mass. Now add boiling distilled water from a wash bottle until the two pans are exactly balanced.

7. Titrate and estimate the reaction of the medium mass; control the result. Calculate the amount of soda solution required to make the reaction of the medium mass +10 (i. e., calculate for 1000 c.c., less the quantity used for the titrations).

8. Add the necessary amount of soda solution and heat in the steamer at 100° C. for twenty minutes, to precipitate the phosphates, etc.

9. Allow the medium mass to cool to 60° C. Well whip the whites of two eggs, add to the contents of the flask and replace in the steamer at 100° C. for about half an hour (until the egg-albumen has coagulated[Pg 167] and formed large, firm masses floating on and in clear gelatine).

10. Filter through papier Chardin into a sterile flask.

11. Tube in quantities of 10 c.c.

12. Sterilise in the steamer at 100° C. for twenty minutes on each of three consecutive days—i. e., by the discontinuous method.

Nutrient Agar-agar.—

1. Weigh a 2-litre flask and note the weight—or counterpoise exactly.

2. Measure out double strength meat extract, 500 c.c., into the "tared" flask.

3. Weigh out and mix 10 grammes of peptone, 5 grammes of salt, and 20 grammes of powdered agar, and make into a thick paste with 150 c.c. distilled water, and add to the meat extract in the flask; place the flask in a water-bath.

4. Arrange the steam can and water-bath as already directed (for the preparation of gelatine) and figured.

5. Bubble live steam (at 100° C.) through the medium mass, for twenty-five minutes, by which time complete solution of the agar is effected.

6. Now weigh the flask and its contents; then (1035[5] grammes + weight of flask) minus (weight of flask and its contents) equals the weight of water required to make up the bulk of the medium to 1 litre. Add the requisite amount (see preparation of gelatine, page 166, step 6).

7. Titrate, and estimate the reaction of the medium mass; control the result. Calculate the amount of[Pg 168] soda solution required to make the reaction of the medium mass + 10 (i. e., calculated for 1000 c.c., less the quantity used for the titrations).

8. Add the necessary amount of soda solution and replace in the steamer for twenty minutes (to complete the precipitation of the phosphates, etc.).

9. Allow the medium mass to cool to 60° C. Well whip the whites of two eggs, add to the contents of the flask, and replace in the steamer at 100° C. for about one hour (until the egg-albumen has coagulated and formed large, firm masses floating on and in clear agar.)

10. Filter through papier Chardin, by the aid of a hot-water funnel, if necessary (Fig. 101), into a sterile flask.

11. Tube in quantities of 10 c.c. or 15 c.c.

12. Sterilise in the steamer at 100° C. for thirty minutes on each of three consecutive days—i. e., by the discontinuous method.

Blood-serum (Inspissated).—

1. Sterilise cylindrical glass jar (Fig. 109) and its cover by dry heat, or by washing first with ether and then with alcohol and drying.

2. Collect blood at the slaughter house from ox or sheep in the sterile cylinder.

3. Allow the vessel to stand for fifteen minutes for the blood to coagulate. (This must be done before leaving the slaughterhouse, otherwise the serum will be stained with hæmoglobin.)

4. Separate the clot from the sides of the vessel by means of a sterile glass rod (the yield of serum is much smaller when this is not done), and place the cylinder in the ice-chest for twenty-four hours.

5. Remove the serum with sterile pipettes, or syphon it off, and fill into sterile tubes (5 c.c. in each) or flasks.[Pg 169]

6. Heat tubes containing serum to 56° C. in a water-bath for half an hour on each of two successive days.

7. On the third day, heat the tubes, in a sloping position, in a serum inspissator to about 72° C. (A coagulum is formed at this temperature which is fairly transparent; above 72° C., a thick turbid coagulum is formed.)

Fig. 109.—Blood-serum jar with wicker basket for transport.

The serum inspissator (Fig. 110) in its simplest form is a double-walled rectangular copper box, closed in by a loose glass lid, and cased in felt or asbestos—the space between the walls is filled with water. The inspissator is supported on adjustable legs so that the serum may be solidified at any desired "slant," and is heated from below by a Bunsen burner controlled by a thermo-regulator. The more elaborate forms resemble the hot-air oven (Fig. 26) in shape and are provided with adjustable shelves so that any desired obliquity of the serum slope can be obtained.

8. Place the tubes in the incubator at 37° C. for[Pg 170] forty-eight hours in order to eliminate those that have been contaminated. Store the remainder in a cool place for future use.

Alternative Method.

Steps 1-5 as above.

6. Sterilise the serum by the fractional method—that is, by exposure in a water-bath to a temperature of 56° C. for half an hour on each of six consecutive days; store in the fluid condition.

7. Coagulate in the inspissator when needed.

Fig. 110.—Serum inspissator.

[Pg 171]

Citrated Blood Agar. Guy's.—

1. Kill a small rabbit with chloroform vapour, and nail it out on a board (as for a necropsy); moisten the hair thoroughly with 2 per cent. solution of lysol.

2. Sterilise several pairs of forceps, scissors, etc. by boiling.

3. Reflect the skin over the thorax with sterile instruments.

4. Open the thoracic cavity by the aid of a fresh set of sterile instruments.

5. Open the pericardium with another set of sterile instruments.

6. Sear the surface of the left ventricle with a red-hot iron.

7. Take a sterile capillary pipette (Fig. 13, c); break off the sealed extremity with a pair of sterile forceps.

8. Steady the heart in a pair of forceps and thrust the point of the pipette through the wall of the ventricle and through the seared area, apply suction to the plugged end of the pipette and fill it with blood.

9. Transfer the entire quantity of blood collected from the rabbit's heart to a small Erlenmeyer flask containing a number of sterile glass beads and 5 c.c. concentrated sod. citrate solution. (See page 378.)

10. Agitate thoroughly and set aside for a couple of hours.

11. Melt up several tubes of nutrient agar (see page 167) and cool to 42° C.

12. With a sterile 10 c.c. graduated pipette transfer 1 c.c. citrated blood from the Erlenmeyer flask to each tube of liquefied agar. Rotate the tube between the hands in order to diffuse the citrated blood evenly throughout the agar.

13. Place the tubes in a sloping position and allow the medium to set.

14. Place tubes of blood agar for forty-eight hours in[Pg 172] the incubator at 37° C. and at the end of that time eliminate any contaminated tubes.

15. Store such tubes as remain sterile for future use.

Milk.—

1. Pour 1 litre of fresh cow's or goat's milk into a large separating funnel, and heat in the steamer at 100° C. for one hour.

2. Remove from the steamer and estimate the reaction of the milk (normal cows' milk averages +17). If of higher acidity than +20, or lower than +10, reject this sample of milk and proceed with another supply of milk from a different source.

Reject milk to which antiseptics have been added as preservatives.

3. Allow the milk to cool, when the fat or cream will rise to the surface and form a thick layer.

4. Draw off the subnatant fat-free milk into sterile tubes (10 c.c. in each).

5. Sterilise in the steamer at 100° C. for twenty minutes on each of five successive days.

6. Incubate at 37° C. for forty-eight hours and eliminate any contaminated tubes. Store the remainder for future use.

Litmus Milk.—

1. Prepare milk as described above, sections 1 to 3.

2. Draw off the subnatant fat-free milk into a flask.

3. Add sterile litmus solution, sufficient to colour the milk a deep lavender.

4. Tube, sterilise, etc., as for milk.

Nutrose Agar (Eyre).—

(This is a modification of the well known Drigalski-Conradi medium originally introduced for the isolation of B. typhosus).

1. Collect 250 c.c. perfectly fresh ox serum (vide[Pg 173] Blood Serum, page 168, steps 1 to 5) and add to it 450 c.c. sterile distilled water.

2. Weigh out agar powder, 20 grammes, and emulsify it with 250 c.c. of the cold serum water.

3. Weigh out

and dissolve in 200 c.c. of serum water heated to 80° C.

4. Mix the agar emulsion and the peptone-nutrose solution in a "tared" flask of 2-litre capacity and add a further 100 c.c. serum water.

5. Complete the solution of the various ingredients by bubbling live steam through the flask as in making nutrient agar.

6. Add further 250 c.c. serum water.

7. Weigh the flask and its contents: then (1045 grammes + weight of flask) minus (weight of flask and its present contents) = weight of fluid required to make up the bulk of the medium to 1 litre. Add the requisite amount of sterile distilled water.

8. Titrate and estimate the reaction of the medium mass. Then standardise to reaction of +2.5.

9. Clarify with egg, and filter as for nutrient agar. (In clarifying, after the addition of the egg white the mixture should be in the steamer for full two hours.)

10. After filtration is complete measure the filtrate, and to every 150 c.c. of the medium add:

11. Tube in quantities of 15 c.c.

12. Sterilise in the steamer at 100° C. for thirty minutes on each of three successive days—i. e., by the discontinuous method for three days.[Pg 174]

Egg Medium (Dorset).—

1. Prepare 1000 c.c. of a 0.85 per cent. solution of sodium chloride in a stout 2-litre flask.

2. Sterilise in the autoclave at 120° C. for twenty minutes. Cool to 20° C.

3. Take 12 fresh eggs; wash the shells first with water then with undiluted formalin: allow the shells to dry.

4. Break the eggs into a sterile graduated cylinder and measure the total volume of the mixed whites and yolks. Add one part sterile saline solution to three parts mixed eggs.

5. Transfer this mixture to a large wide-mouthed stoppered bottle previously sterilised. Add sterile glass beads and shake thoroughly in a mechanical shaker for about thirty minutes, or whip with an egg-whisk.

6. Filter through coarse butter muslin into a sterile flask.

7. Tube in quantities of 10 c.c.

8. Solidify in the sloping position in the inspissator at 75° C. for one hour.

9. Place the tubes for forty-eight hours in the incubator at 37° C., and eliminate any contaminated tubes.

To prevent drying, 0.5 c.c. glycerine bouillon (see page 209) may be added to each tube between steps 8 and 9.

10. Cap those tubes of media which remain sterile with india-rubber caps and store for future use.

Potato.—

1. Choose fairly large potatoes, wash them well, and scrub the peel with a stiff nail-brush.[Pg 175]

2. Peel and take out the eyes.

3. Remove cylinders from the longest diameter of each potato by means of an apple-corer or a large cork-borer (i. e., one of about 1.4 cm. diameter).

The reaction of the fresh potato is strongly acid to phenolphthalein. If, therefore, the potatoes are required to approximate +10, as for the cultivation of some of the vibrios, the cylinders should be soaked in a 1 per cent. solution of sodium carbonate for thirty minutes.

4. Cut each cylinder obliquely from end to end, forming two wedge-shaped portions.

5. Place a small piece of sterilised cotton-wool, moistened with sterile water, at the bottom of a sterile test-tube; insert the potato wedge into the tube so that its base rests upon the cotton-wool. Now plug the tube with cotton-wool (Fig. 111).

6. Sterilise in the steamer at 100° C. for twenty minutes on each of five consecutive days.

Fig. 111.—Potato tube.

Beer Wort.—Wort is chiefly used as a medium for the cultivation of yeasts, moulds, etc., both in its fluid form and also when made solid by the addition of gelatine or agar. The wort is prepared as follows:

1. Weigh out 250 grammes crushed malt and place in a 2-litre flask.

2. Add 1000 c.c. distilled water, heated to 70° C., and close the flask with a rubber stopper.[Pg 176]

3. Place the flask in a water-bath regulated to 60°C. and allow the maceration to continue for one hour.

4. Strain through butter muslin into a clean flask and heat in the steamer for thirty minutes.

5. Filter through Swedish filter paper.

6. Tube in quantities of 10 c.c. or store in flasks.

7. Sterilise in the steamer at 100° C. for twenty minutes on each of three consecutive days.

The natural reaction of the wort should not be interfered with.

Wort Gelatine.—

1. Measure out wort (prepared as above), 900 c.c., into a sterile flask.

2. Weigh out gelatine, 100 grammes (= 10 per cent.), and add it to the wort in the flask.

3. Bubble live steam through the mixture for ten minutes, to dissolve the gelatine.

4. Cool to 60°C.; clarify with egg as for nutrient gelatine (vide page 164).

5. Filter through papier Chardin.

6. Tube, and sterilise as for nutrient gelatine.

Wort Agar.—

1. Measure out wort (as above), 700 c.c., into a sterile flask.

2. Weigh out powdered agar, 20 grammes; mix into a smooth paste with 200 c.c. of cold wort and add to the wort in the flask.

3. Bubble live steam through the mixture for twenty minutes, to dissolve the agar.[Pg 177]

4. Cool to 60° C.; clarify with egg as for nutrient agar (vide page 167).

5. Filter through papier Chardin, using the hot-water funnel.

6. Tube, and sterilise as for nutrient agar.

Peptone Water (Dunham).—

1. Weigh out Witté's peptone, 10 grammes, and salt, 5 grammes, and emulsify with about 250 c.c. of distilled water previously heated to 60° C.

2. Pour the emulsion into a litre flask and make up to 1000 c.c. by the addition of distilled water.

3. Heat in the steamer at 100° C. for thirty minutes.

4. Filter through Swedish filter paper.

5. Tube in quantities of 10 c.c. each.

6. Sterilise in the steamer at 100° C. for twenty minutes on each of three consecutive days.

"Sugar" or "Carbohydrate" Media.—

Formerly the ability of bacteria to induce hydrolytic changes in carbohydrate substances was observed only in connection with a few well-defined sugars, but of recent years it has been shown that when using litmus as an indicator these so-called "fermentation reactions" facilitate the differentiation of closely allied species, and the list of substances employed in this connection has been considerably extended. The media prepared with them are now no longer regarded as special, but are comprised in the "stock media" of the laboratory. The chief of these substances are the following, arranged in accordance with their chemical constitution:

These substances should be obtained from Kahlbaum (of Berlin); in the pure form, and when possible as large crystals, and the method of preparing a medium containing either of them may be exemplified by describing Dextrose Solution.

Dextrose Solution.—

1. Weigh out

and grind together in a mortar; then emulsify in 100 c.c. of distilled water heated to 60° C.

2. Place in a flask and add

3. Steam in the steamer at 100° C. for twenty minutes to dissolve the peptone and glucose.

4. Add

(The substances enumerated above react acid to phenolphthalein, but variously toward the neutral litmus solution. To such as react acid, add very cautiously n/1 sodium hydrate solution to the medium in bulk until the neutral tint has returned).

5. Fill into tubes in which have previously been placed the inverted Durham's gas tubes.

6. Sterilise in the steamer at 100° C. for twenty minutes on each of three successive days.

Neutral Litmus Solution.

The most satisfactory is the Kubel-Tiemann, prepared by Kahlbaum. It can however be made in the laboratory as follows:

1. Weigh out

and place in a well stoppered 500 c.c. bottle; measure out and add 300 c.c. alcohol 95 per cent.

2. Shake well at least once a day for seven days—the alcohol acquires a green colour.

3. Decant off the green alcohol and fill a further 300 c.c. 95 per cent. alcohol into the bottle and repeat the shaking.

4. Repeat this process until on adding fresh alcohol the fluid only becomes tinged with violet.

5. Pour off the alcohol, leaving the litmus as dry as possible. Connect up the bottle to an air pump and evaporate off the last traces of alcohol.

6. Transfer the dry litmus to a litre flask, measure in 600 c.c. distilled water and allow to remain in contact 24 hours with frequent shakings.

7. Filter the solution into a clean flask and add one or two drops of pure concentrated sulphuric acid until the litmus solution is distinctly wine-red in colour.

8. Add excess of pure solid baryta and allow to stand until the reaction is again alkaline.

9. Filter.

10. Bubble CO₂ through the solution until reaction is definitely acid.

11. Sterilise in the steamer at 100° C. for thirty minutes on each of three consecutive days. This sterilises the solution and also drives off the carbon dioxide, leaving the solution neutral.[Pg 180]

Media for anaerobic cultures. In addition to the foregoing media, all of which can be, and are employed in the cultivation of anaerobic bacteria, certain special media containing readily oxidised substances are commonly used for this purpose. The principal of these are as follows:

[Pg 182]

XII. SPECIAL MEDIA.

In this chapter are collected a number of media which have been elaborated by various workers for special purposes, grouped together under headings which indicate their chief utility. In many instances the name of the originator of the medium is given, but without reference to his original instructions, since these are in many cases inadequate to the requirements of the isolated worker, who would probably fail to reproduce the medium in a form giving the results attributed to it by its author. Such modifications have therefore been introduced as make for uniformity between the different batches of media.

A considerable number of coloured media, chiefly intended for work with intestinal bacteria, have been included; but beyond the fact that the author's modification of the Drigalski-Conradi medium has been included amongst the routine media of the laboratory, no comment has been made upon their relative values, since only by observation and practice can the skill necessary to utilise their full value be acquired.

The instructions as to sterilisation are rarely given in full; the routine method of exposure in the steam steriliser at 100° C. (without pressure) for twenty minutes on each of three successive days for all fluid media, and thirty minutes on each of three successive days for all liquefiable or solid media must be carried out; and only when these general rules are to be departed from are further details given.[Pg 183]

Media for the Study of the Chemical Composition of Bacteria.

Asparagin Medium (Uschinsky).—

1. Weigh out and mix

2. Dissolve the mixture in distilled water 1000 c.c.

3. Add glycerine, 40 c.c.

4. Tube, and sterilise as for nutrient bouillon.

Asparagin Medium (Frankel and Voges).—

1. Weigh out and mix

and dissolve in

2. Tube, and sterilise as for nutrient bouillon.

Proteid Free Broth (Uschinsky).—

1. Weigh out and mix

2. Dissolve the mixture in distilled water 1000 c.c.

3. Add glycerine 30 c.c.

4. Tube and sterilise as for nutrient broth.

Media for the Study of Biochemical Reaction.

Inosite-free Media—Bouillon (Durham).—

1. Prepare meat extract, 1000 c.c. (vide page 148), from bullock's heart which has been "hung" for a couple of days.

2. Prepare nutrient bouillon (+10), 1000 c.c. (vide, page 161), from the meat extract, and store in 1-litre flask.[Pg 184]

3. Inoculate the bouillon from a pure cultivation of the B. lactis aerogenes, and incubate at 37° C. for forty-eight hours.

4. Heat in the steamer at 100° C. for twenty minutes to destroy the bacilli and some of their products.

5. Estimate the reaction of the medium and if necessary restore to +10.

6. Inoculate the bouillon from a pure cultivation of the B. coli communis and incubate at 37° C. for forty-eight hours.

7. Heat in the steamer at 100° C. for twenty minutes.

Now fill two fermentation tubes with the bouillon, tint with litmus solution, and sterilise; inoculate with B. lactis aerogenes. If no acid or gas is formed, the bouillon is in a sugar-free condition; but if acid or gas is present, again make the bouillon in the flask +10, reinoculate with one or other of the above-mentioned bacteria, and incubate; then test again. Repeat this till neither acid nor gas appears in the medium when used for the cultivation of either of the bacilli referred to above.

8. After the final heating, stand the flask in a cool place and allow the growth to sediment. Filter the supernatant broth through Swedish filter paper. If the filtrate is cloudy, filter through a porcelain filter candle.

9. Tube, and sterilise as for bouillon.

Bouillon prepared in the above-described manner will prove to be absolutely sugar-free; and from it may be prepared nutrient sugar-free gelatine or agar, by dissolving in it the required percentage of gelatine or agar respectively and completing the medium according to directions given on pages 166 and 167. The most important application of inosite-free bouillon is its use in the preparation of sugar bouillons, whether glucose, maltose, lactose, or saccharose, of exact percentage composition.

Sugar (Dextrose) Bouillon.—

1. Measure out nutrient bouillon, 1000 c.c. (vide page 163, sections 1 to 6) or sugar-free bouillon (vide supra).

2. Weigh out glucose (anhydrous), 20 grammes (= 2 per cent.), and dissolve in the fluid.

3. Tube, and sterilise as for bouillon.

Ordinary commercial glucose serves the purpose equally well, but is not recommended, as during the process of sterilisation it causes the medium to gradually deepen in colour.

Sugar Gelatine.—

1. Prepare nutrient gelatine (vide page 164, sections 1 to 7). Measure out 1000 c.c.

2. Weigh out glucose, 20 grammes (= 2 per cent.), and dissolve in the hot gelatine.[Pg 185]

3. Filter through papier Chardin.

4. Tube, and sterilise as for nutrient gelatine.

Sugar Agar.—

1. Prepare nutrient agar (vide page 167, sections 1 to 8). Measure out 1000 c.c.

2. Weigh out glucose, 20 grammes (= 2 per cent.), and dissolve in the clear agar.

3. Tube, and sterilise as for nutrient agar.

Iron Bouillon.—

1. Measure out nutrient bouillon, 1000 c.c. (vide page 141, sections 1 to 6).

2. Weigh out ferric tartrate, 1 gramme (= 0.1 per cent.), and dissolve it in the bouillon.

3. Tube, and sterilise as for bouillon.

Lead Bouillon.—

1. Measure out nutrient bouillon, 1000 c.c. (vide page 163, sections 1 to 6).

2. Weigh out lead acetate, 1 gramme (= 0.1 per cent.), and dissolve it in the bouillon.

3. Tube, and sterilise as for bouillon.

Nitrate Bouillon.—

1. Measure out nutrient bouillon, 1000 c.c. (vide page 163, sections 1 to 6).

2. Weigh out potassium nitrate, 5 grammes (= 0.5 per cent.), and dissolve it in the bouillon.

3. Tube, and sterilise as for bouillon.

Iron Peptone Solution (Pakes).—

1. Weigh out peptone, 30 grammes, and emulsify it with 200 c.c. tap water, previously heated to about 60°C.

2. Wash the emulsion into a litre flask with 800 c.c. tap water.

3. Weigh out salt, 5 grammes, and sodium phosphate, 3 grammes, and dissolve in the mixture in the flask.

4. Heat the mixture in the steamer at 100° C. for thirty minutes,[Pg 186] to complete the solution of the peptone, and filter into a clean flask.

5. Fill into tubes in quantities of 10 c.c. each.

6. Add to each tube 0.1 c.c. of a 2 per cent. neutral solution of ferric tartrate. (A yellowish-white precipitate forms.)

7. Sterilise as for nutrient bouillon.

Lead Peptone Solution.—

Prepare as for iron peptone solution but in step 6 substitute 0.1 c.c. of a 1 per cent. neutral aqueous solution of lead acetate.

Nitrate Peptone Solution (Pakes).—

1. Weigh out Witté's peptone, 10 grammes, and emulsify it with 200 c.c. ammonia-free distilled water previously heated to 60°C.

2. Wash the emulsion into a flask and make up to 1000 c.c., with ammonia-free distilled water.

3. Heat in the steamer at 100° C. for twenty minutes.

4. Weigh out sodium nitrate, 1 gramme, and dissolve in the contents of the flask.

5. Filter through Swedish filter paper.

6. Tube, and sterilise as for nutrient bouillon.

Litmus Bouillon.—

1. Measure out nutrient bouillon, 1000 c.c. (vide page 163, sections 1 to 6).

2. Add sufficient sterile litmus solution to tint the medium a dark lavender colour. (Media rendered +10 will usually react very faintly alkaline or occasionally neutral to litmus.)

3. Tube, and sterilise as for bouillon.

Rosolic Acid Peptone Solution.—

1. Weigh out rosolic acid (corallin), 0.5 gramme, and dissolve it in 80 per cent. alcohol, 100 c.c. Keep this as a stock solution.

2. Measure out peptone water (Dunham), 100 c.c., and rosolic acid solution, 2 c.c., and mix.

3. Heat in the steamer at 100° C. for thirty minutes.

4. Filter through Swedish filter paper.

5. Tube, and sterilise as for nutrient bouillon.

Capaldi-Proskauer Medium, No. I.—

1. Weigh out and mix

2. Dissolve in water 1000 c.c. in a 2-litre flask[Pg 187]

3. Weigh out and mix

and add to contents of flask.

4. Measure out 25 c.c. of the solution and titrate it against decinormal sodic hydrate, using litmus as the indicator. Control the result and estimate the amount of sodic hydrate necessary to be added to render the remainder of the solution neutral to litmus. Add this quantity of sodic hydrate.

5. Filter.

6. Add litmus solution 47.5 c.c. (= 5 per cent.).

7. Tube, and sterilise as for nutrient bouillon.

Capaldi-Proskauer Medium No. II.—

1. Weigh out and mix

2. Dissolve in water 1000 c.c. in a 2-litre flask.

3. Neutralise to litmus as in No. I (vide supra, Step 4).

4. Filter.

5. Add litmus solution 47.5 c.c. (= 5 per cent.).

6. Tube, and sterilise as for nutrient bouillon.

Urine Media. Bouillon.—

1. Collect freshly passed urine in sterile flask.

2. Place the flask in the steamer at 100° C. for thirty minutes.

3. Filter through two thicknesses of Swedish filter paper.

4. Tube, and sterilise as for nutrient bouillon. (Leave the reaction unaltered.)

Urine Gelatine.—

1. Collect freshly passed urine in sterile flask.

2. Take the specific gravity, and, if above 1010, dilute with sterile water until that gravity is reached.

3. Estimate (with control) at the boiling-point, and note the reaction of the urine.

4. Weigh out gelatine, 10 per cent., and add to the urine in the flask.

5. Heat in the steamer at 100° C. for one hour to dissolve the gelatine.

6. Estimate the reaction and add sufficient caustic soda solution to restore the reaction of the medium mass to the equivalent of the original urine.

7. Cool to 60° C. and clarify with egg as for nutrient gelatine (vide page 166).

8. Filter through papier Chardin.

9. Tube, and sterilise as for nutrient gelatine.[Pg 188]

Urine Gelatine (Heller).—

1. Collect freshly passed urine in sterile flask.

2. Filter through animal charcoal to remove part of the colouring matter.

3. Take the specific gravity, and if above 1010, dilute with sterile water till this gravity is reached.

4. Add Witté's peptone, 1 per cent.; salt, 0.5 per cent.; gelatine, 10 per cent.

5. Heat in the steamer at 100° C. for one hour, to dissolve the gelatine, etc.

6. Add normal caustic soda solution in successive small quantities, and test the reaction from time to time with litmus paper, until the fluid reacts faintly alkaline.

7. Cool to 60° C. and clarify with egg as for nutrient gelatine (vide page 166).

8. Filter through papier Chardin.

9. Tube, and sterilise as for nutrient gelatine.

Urine Agar.—

1. Collect freshly passed urine in sterile flask.

2. Take the specific gravity and if above 1010, dilute with sterile water till this gravity is reached.

3. Weigh out 1.5 per cent. or 2 per cent. powdered agar, and add it to the urine.

4. Heat in the steamer at 100° C. for ninety minutes to dissolve the agar.

5. Cool to 60° C. and clarify with egg as for nutrient agar (vide page 168).

6. Filter through papier Chardin, using the hot-water funnel.

7. Tube, and sterilise as for nutrient agar.

(Leave the reaction unaltered.)

Serum Sugar Media (Hiss).—

In these media the fermentation of carbohydrate substance by bacterial action is indicated by the coagulation of the serum proteids in addition to the production of an acid reaction.

Serum Dextrose Water (Hiss).—

1. Measure out into a litre flask

2. Weigh out

and dissolve in the serum water.

3. Filter through Swedish filter paper.

4. Measure out and add to the medium

[Pg 189]

5. Tube in quantities of 10 c.c. and sterilise in the steamer at 100° C. for twenty minutes on each of three successive days.

Lævulose, galactose, maltose, lactose, etc., can be substituted in similar amounts for dextrose and the medium completed as above.

Omeliansky's Nutrient Fluid (For Cellulose Fermenters).—

1. Weigh out and mix

2. Dissolve in distilled water 4000 c.c.

3. Flask in quantities of 250 c.c.

4. Weigh out and add 5 grammes precipitated chalk to each flask.

5. Sterilise in the steamer at 100° C. for twenty minutes on each of three successive days.

Media for the Study of Chromogenic Bacteria.

Milk Rice (Eisenberg).—

1. Measure out nutrient bouillon, 70 c.c., and milk, 210 c.c., and mix thoroughly.

2. Weigh out rice powder, 100 grammes, and rub it up in a mortar with the milk and broth mixture.

3. Fill the paste into sterile capsules, spreading it out so as to form a layer about 0.5 cm. thick, over the bottom of each.

4. Heat over a water-bath at 100° C. until the mixture solidifies.

5. Replace the lids of the capsules. Sterilise in the steamer at 100° C. for thirty minutes on each of three consecutive days.

(A solid medium of the colour of café au lait is thus produced.)

Milk Rice (Soyka).—

1. Measure out nutrient bouillon, 50 c.c., and milk, 150 c.c., and mix thoroughly.

2. Weigh out rice powder, 100 grammes, and rub it up in a mortar with the milk and broth mixture.

3. Fill the paste into sterile capsules, to form a layer over the bottom of each.

4. Replace the lids of the capsules.

5. Sterilise in the steamer at 100° C. for thirty minutes on each of three consecutive days.

(A pure white, opaque medium is thus formed.)[Pg 190]

Media for the Study of Phosphorescent and Photogenic Bacteria.

Fish Bouillon.—

1. Weigh out herring, mackerel, or cod, 500 grammes, and place in a large porcelain beaker (or enamelled iron pot).

2. Weigh out sodium chloride, 26.5 grammes; potassium chloride, 0.75 gramme; magnesium chloride, 3.25 grammes; and dissolve in 500 c.c. distilled water. Add the solution to the fish in the beaker.

3. Place the beaker in a water-bath and proceed as in preparing meat extract—i. e., heat gently at 40° C. for twenty minutes, then rapidly raise the temperature to, and maintain at, the boiling-point for ten minutes.

4. Strain the mixture through butter muslin into a clean flask.

5. Weigh out peptone, 5 grammes, and emulsify with about 200 c.c. of the hot fish water; incorporate thoroughly with the remainder of the fish water in the flask.

6. Heat in the steamer at 100° C. for twenty minutes to complete the solution of the peptone.

7. Filter through Swedish filter paper.

8. When the fish bouillon is cold, if it is to be used as fluid medium, make up to 1000 c.c. by the addition of distilled water. If, however, it is to be used as the basis for agar or gelatine media store it in the "Double Strength" condition.

9. Tube and sterilise as for nutrient bouillon.

As an alternative method "Marvis" fish food (16 grammes) may be substituted for the 500 grammes of fresh fish.

Fish Gelatine.—

1. Measure out double strength fish bouillon, 500 c.c., into a "tared" 2-litre flask.

2. Add sheet gelatine, 100 grammes, cut into small pieces.

3. Bubble live steam through the mixture for fifteen minutes to dissolve the gelatine.

4. Weigh the flask and its contents; adjust the weight to the calculated figure for one litre of medium (1135.5 grammes) by the addition of distilled water at 100° C. (vide page 166).

5. Cool to below 60°C., and clarify with egg.

6. Filter through papier Chardin.

7. Tube, and sterilise as for nutrient gelatine.

Shake well after the final sterilisation, to aerate the medium.

Fish Gelatine-Agar.—

1. Weigh out powdered agar, 5 grammes, and emulsify it with 200 c.c. double strength fish bouillon.

2. Wash the emulsion into a "tared" 2-litre flask with 300 c.c. fish bouillon.[Pg 191]

3. Weigh out sheet gelatine, 70 grammes, cut it into small pieces and add it to the contents of the flask.

4. Bubble live steam through the mixture to dissolve the gelatine and agar.

5. Weigh the flask and contents. Adjust the weight to the calculated figure for one litre of medium (1110.5 grammes) by the addition of distilled water at 100° C. (vide page 166).

6. Cool to below 60° C. and clarify with egg.

7. Filter through papier Chardin.

8. Tube, and sterilise as for nutrient gelatine.

Shake well after the final sterilisation, to aerate the medium.

Media for the Study of Yeasts and Moulds.

Pasteur's Solution.—

(Reaction alkaline).

1. Weigh out and mix the ash from 10 grammes of yeast; ammonium tartrate, 10 grammes; cane sugar, 100 grammes.

2. Dissolve the mixture in distilled water, 1000 c.c.

3. Tube or flask, and sterilise as for nutrient bouillon.

Yeast Water (Pasteur).—

1. Weigh out pressed yeast, 75 grammes; place in a 2-litre flask and add 1000 c.c. distilled water.

2. Heat in the steamer at 100° C. for thirty minutes.

3. Filter through papier Chardin.

4. Tube or flask, and sterilise as for nutrient bouillon.

Cohn's Solution.—

1. Weigh out and mix

and dissolve in

2. Tube, or flask and sterilise as for nutrient bouillon.

Naegeli's Solution.—

1. Weigh out and mix

and dissolve in

2. Tube or flask; sterilise as for nutrient bouillon.[Pg 192]

Plaster-of-Paris Discs.—

1. Take large corks, 2.5 cm. diameter, and roll a piece of stiff note-paper round each, so that about a centimetre projects as a ridge above the upper surface of the cork, and secure in position with a pin (Fig. 112).

2. Mix plaster-of-Paris into a stiff paste with distilled water, and fill each of the cork moulds with the paste.

3. When the plaster has set, remove the paper from the corks, and raise the plaster discs.

4. Place the plaster discs on a piece of asbestos board and sterilise by exposing in the hot-air oven to 150° C. for half an hour.

Fig. 112.—Cork and paper mould for plaster-of-Paris disc.

5. Remove the sterile discs from the oven by means of sterile forceps, place each inside a sterile capsule, and moisten with a little sterile water.

6. Sterilise in the steamer at 100° C. for thirty minutes on each of three consecutive days.

Gypsum Blocks (Engel and Hansen).—

These are in the form of truncated cones and for their preparation small tin moulds are required, each having a diameter of 5.5 cm. at the base and 4 cm. at the truncated apex. The height (or depth) of a mould is 4.5 to 5 cm.

1. Mix powdered calcined gypsum into a stiff paste with distilled water.

2. Fill the paste into the moulds and allow it to set and dry by exposure to air.

3. Remove the block from the mould and transfer it to a double glass dish of adequate size (7 cm. diameter × 7 cm. high).

4. Sterilise block in its dish for one hour in the hot-air oven at 115°C.

5. Carefully open the dish and add sterile distilled water to moisten the block and form a layer in the bottom of the dish 1 cm. deep.

Wine Must.—(Wine must is obtained from Sicily, in hermetically sealed tins, in a highly concentrated form—as a thick syrup—but not sterilised.)

1. Weigh out "wine must," 200 grammes, place in a 2-litre flask and add distilled water, 800 c.c.

2. Weigh out ammonium tartrate, 5 grammes, and add to the dilute must.

3. Place the flask in a water-bath regulated to 60° C. for one hour and incorporate the mixture thoroughly by frequent shaking.

4. Filter through papier Chardin.

5. Tube, and sterilise as for nutrient bouillon.[Pg 193]

Wheat Bouillon (Gasperini).—

1. Weigh out and mix wheat flour, 150 grammes; magnesium sulphate, 0.5 gramme; potassium nitrate, 1 gramme; glucose, 15 grammes.

2. Dissolve the mixture in 1000 c.c. of water heated to 100°C.

3. Filter through papier Chardin.

4. Tube, and sterilise as for nutrient bouillon.

Bread Paste.—

1. Grate stale bread finely on a bread-grater.

2. Distribute the crumbs in sterile Erlenmeyer flasks, sufficient to form a layer about one centimetre thick over the bottom of each.

3. Add as much distilled water as the crumbs will soak up, but not enough to cover the bread.

4. Plug the flasks and sterilise in the steamer at 100° C. for thirty minutes on each of four consecutive days.

Media for the Study of Parasitic Moulds.

French Proof Agar (Sabouraud).—

1. Weigh out Chassaing's peptone, 10 grammes, and emulsify it with 200 c.c. distilled water previously heated to 60°C.

2. Weigh out powdered agar, 13 grammes, and emulsify with 200 c.c. cold distilled water.

3. Mix the two emulsions and wash into a tared 2-litre flask with 600 c.c. distilled water.

4. Bubble live steam through the mixture for twenty minutes, to dissolve the agar.

5. Cool to 60° C. and clarify with egg as for nutrient agar (vide page 168).

6. Filter through Papier Chardin, using the hot-water funnel.

7. Weigh out French maltose, 40 grammes, and dissolve in the agar.

8. Tube, and sterilise as for nutrient agar.

English Proof Agar (Blaxall).—Substitute Witté's peptone for that of Chassaing, and proceed as for French proof agar.

French Mannite Agar, Sabouraud.—(For cultivation of Favus.)

Proceed exactly as in preparing French Proof agar vide supra substituting Mannite (38 grammes) for maltose.

Media for the Study of Milk Bacteria.

Gelatine Agar.—This medium is prepared by adding to nutrient gelatine sufficient agar to ensure the solidity of the medium when incubated at temperatures above 22° C. If it is intended[Pg 194] to employ an incubating temperature of 30°C., 10 per cent. gelatine and 0.5 per cent. agar must be dissolved in the meat extract before the addition of the peptone and salt; while for incubating at 37°C., 12 per cent. gelatine and 0.75 per cent. agar must be used. Avoid the addition of more agar than is absolutely necessary, otherwise the action upon the medium of such organisms as elaborate a liquefying ferment may be retarded or completely absent.

1. Measure out 400 c.c. double strength meat extract into a "tared" 2-litre flask, and add to it gelatine, 100 grammes.

2. Weigh out powdered agar, 5 grammes, emulsify with 100 c.c., cold distilled water and add to the contents of the flask.

3. Dissolve the agar and gelatine by bubbling live steam through the flask for twenty minutes.

4. Weigh out peptone, 10 grammes; salt, 5 grammes; emulsify with 100 c.c. double strength meat extract previously heated to 60°C., and add to the contents of the flask.

5. Replace in the steamer for fifteen minutes. Then adjust the weight to the calculated figure for one litre (in this instance 1120 grammes) by the addition of distilled water at 100°C.

6. Estimate the reaction; control the result. Then add sufficient caustic soda solution to render the reaction +10.

7. Replace in the steamer at 100° C. for twenty minutes.

8. Cool to 60° C. Clarify with egg as for nutrient agar.

9. Filter through papier Chardin, using the hot-water funnel.

10. Tube, and sterilise as for nutrient agar.

Agar Gelatine (Guarniari).—

1. Measure out double strength meat extract, 400 c.c., into a "tared" 2-litre flask, and add to it gelatine, 50 grammes.

2. Weigh out powdered agar, 3 grammes; emulsify with cold distilled water, 50 c.c., and add to the contents of the flask.

3. Dissolve the agar and gelatine by bubbling live steam through the flask for twenty minutes.

4. Weigh out Witté's peptone, 25 grammes; salt, 5 grammes, and emulsify with 100 c.c. double strength meat extract previously heated to 60°C., and add to the contents of the flask.

5. Replace in the steamer for fifteen minutes.

6. Weigh the flask and make up the medium mass to the calculated figure for one litre (1083 grammes) by the addition of distilled water at 100°C.

7. Neutralise carefully to litmus paper by the successive additions of small quantities of normal soda solution.

8. Replace in the steamer at 100° C. for twenty minutes.

9. Cool to 60° C. Clarify with egg as for nutrient agar.

10. Filter through papier Chardin, using the hot-water funnel.

11. Tube, and sterilise as for nutrient agar.[Pg 195]

Whey Gelatine.—

1. Curdle fresh milk by warming to 60°C., and adding rennet; filter off the whey into a sterile "tared" flask.

2. Estimate and note the reaction of the whey.

3. Weigh out gelatine, 10 per cent., and add it to the whey in the flask.

4. Bubble live steam through the mixture fifteen minutes to dissolve the gelatine; and weigh.

5. Estimate the reaction of the medium mass; then add sufficient caustic soda solution to restore the reaction of the medium mass (i. e., total weight minus weight of flask) to the equivalent of the original whey.

6. Cool to 60° C. and clarify with egg as for nutrient gelatine (vide page 166).

7. Filter through papier Chardin.

8. Tube, and sterilise as for nutrient gelatine.

Whey Agar.—

1. Curdle fresh milk by warming to 60°C., and adding rennet; filter off the whey into a sterile flask.

2. Weigh out agar, 1.5 or 2 per cent., and add it to the whey in the flask.

3. Bubble live steam through the mixture for twenty minutes, to dissolve the agar.

4. Cool to 60°C.; clarify with egg as for nutrient agar (vide page 168).

5. Filter through papier Chardin, using the hot-water funnel.

6. Tube, and sterilise as for nutrient agar.

Litmus Whey.—

1. Curdle fresh milk by warming to 60° C. and adding rennet.

2. Filter off the whey through butter muslin into a sterile flask.

3. Neutralise to litmus by the cautious addition of citric acid solution 4 per cent. (Do not neutralise with mineral acid.)

4. Heat in the steamer at 100° C. for one hour to coagulate all the proteid.

(If the whey is cloudy when removed from the steamer allow it to stand for forty-eight hours in the ice chest and then decant off the clear fluid—or filter through a Berkefeld filter candle.)

5. Filter into a sterile flask.

6. Tint the whey with litmus solution to a deep purple red.

7. Tube, and sterilise as for milk.

Litmus Whey (Petruschky).—

1. Measure out into a flask

[Pg 196]

2. Add

and boil.

3. Filter off coagulated casein.

4. Neutralise to litmus by the addition of n/1 caustic soda solution and boil. Whey now cloudy and acid again.

5. Again neutralise to litmus by addition of n/10 caustic soda solution.

6. Filter.

7. Tint the whey with neutral litmus solution to a deep purple colour.

8. Tube and sterilise as for milk.

Litmus Whey Gelatine.—

1. Measure out milk 1000 c.c. into a tared 2-litre flask.

2. Add hydrochloric acid (or glacial acetic acid) 1.5 c.c. and boil for five minutes.

3. Filter off the casein, and make the whey faintly alkaline to litmus.

4. Weigh out

and emulsify in a few cubic centimeters of the whey and return to the flask.

5. Weigh out

add it to the whey in the flask and incorporate the mixture by bubbling through live steam.

6. Clear with egg and filter.

7. Make the weight of the medium mass to the calculated figure for one litre (1060 grammes) by the addition of distilled water.

8. Weigh out

and dissolve in the fluid whey gelatine.

9. Add sterile litmus solution to the required tint.

10. Tube and sterilise for twenty minutes in steamer at 100°C. on each of five successive days.

This medium will remain semi-fluid at the room temperature, and may be used for cultures in the cool or hot incubator.

Litmus Whey Agar is prepared in a similar manner to Whey Gelatine, with the substitution of 15 grammes of agar for the gelatine.

Malt Extract Solution (Herschell).—

1. Measure into a flask distilled water 1000 c.c.

2. Weigh out

and add to distilled water in flask.[Pg 197]

3. Boil for five minutes, allow to stand, and decant off clear fluid from sediment.

4. Tube and sterilise as for nutrient bouillon.

Media for the Study of Earth Bacteria, Nitrogen Fixers.

Earthy Salts Agar (Lipman and Brown).—(For the enumeration of soil organisms.)

1. Measure out

Emulsify in 200 c.c. distilled water.

2. Wash the agar emulsion into a tared 2-litre flask with 400 c.c. distilled water.

3. Weigh out

Emulsify in 50 c.c. distilled water and add to the contents of the flask.

4. Bubble live steam through the mixture for twenty minutes to dissolve the agar.

5. Weigh out and mix

and add to the contents of the flask.

6. Adjust the weight of the medium mass to the calculated figure for one litre (1025 grammes) by the addition of distilled water at 100°C.

7. Titrate the medium mass and adjust the reaction to +5.

8. Cool to 60° C. Clarify with egg and filter.

9. Tube in quantities of 10 c.c. and sterilise as for nutrient agar.

Beyrinck's Solution. I.—(For the cultivation of nitrogen fixing organisms.)

1. Weigh out and mix 1 gramme potassium hydrogen phosphate, 0.2 gramme magnesium sulphate, and 0.02 gramme sodium chloride.

2. Dissolve in water 1000 c.c., in a 2-litre flask.

3. Add 1 c.c. of a one per thousand aqueous solution of ferrous sulphate.

4. Add 1 c.c. of a one per thousand solution manganese sulphate.

5. Weigh out 20 grammes dextrose and add to the contents of the flask (dextrose up to 40 grammes may be used for the different organisms).

6. Steam for twenty minutes, filter.

7. Tube, and sterilise as for nutrient bouillon.[Pg 198]

Beyrinck's Solution. II.—(For growth of Azobacter.)

Proceed as in preparing solution No. I, substituting mannite for dextrose in step 5.

Winogradsky's Solution (for Nitric Organisms).—

1. Weigh out and mix.

and dissolve in

2. Fill into flasks, in quantities of 20 c.c. and add to each a small quantity of freshly washed magnesium carbonate.

3. Sterilise in the steamer at 100° C. for twenty minutes on each of three consecutive days.

4. Add to each flask containing 20 c.c. solution, 2 c.c. of a sterile 2 per cent. solution of ammonium sulphate.

5. Incubate at 37° C. for forty-eight hours and eliminate any contaminated culture flasks. Store the remainder for future use.

Winogradsky's Solution (for Nitrous Organisms).—

1. Weigh out and mix

and dissolve in

2. Add 5 to 10 grammes basic magnesium carbonate, previously sterilised by boiling.

3. Fill into flasks and sterilise, etc., as for previous solution.

Silicate Jelly (Winogradsky).—

1. Weigh out and mix

and dissolve in

Label—Solution A.

2. Weigh out and mix

and dissolve in

Label—Solution B.[Pg 199]

3. Weigh out

and dissolve in

4. Pour the silicic acid solution into a large porcelain basin.

5. Mix equal quantities of the solutions A and B; then add successive small quantities of the mixed salts to the silicic acid solution, stirring continuously with a glass rod, until a jelly of sufficiently firm consistence has been formed.

6. Spread a layer of this jelly over the bottom of each of several large capsules or "plates."

7. Sterilise in the steamer at 100° C. for thirty minutes on each of three consecutive days.

Media for the Study of Water Bacteria.

Naehrstoff Agar (Hesse and Niedner).—(For enumeration of water organisms.)

1. Weigh out: agar, 12.5 grammes and emulsify in 250 c.c. distilled water.

2. Wash the agar emulsion into a tared 2-litre flask with a further 250 c.c. distilled water.

3. Dissolve by bubbling live steam through the mixture.

4. Emulsify Naehrstoff-Heyden (albumose) 7.5 grammes in 200 c.c. cold distilled water and add to melted agar.

5. Adjust weight of medium mass to the calculated figure for one litre (1020 grammes) by addition of distilled water at 100° C.

6. Clarify with white of egg and filter.

7. Tube in quantities of 10 c.c. and sterilise in the steamer at 100° C. for twenty minutes on each of three successive days.

Bile Salt Broth—Double Strength.—

1. Weigh out Witté's peptone, 40 grammes, and emulsify with 300 c.c. distilled water previously warmed to 60° C.

2. Wash the peptone emulsion into a litre flask with 600 c.c. distilled water.

3. Weigh out sodium taurocholate, 10 grammes, and glucose, 10 grammes; dissolve in 100 c.c. distilled water and add to the peptone emulsion in the flask.

4. Heat in the steamer at 100° C. for twenty minutes.

5. Filter through Swedish filter paper into a sterile flask.

6. Add sterile neutral litmus solution sufficient to colour the medium to a deep purple.

7. Fill into small Erlenmeyer flasks in quantities of 25 c.c.

8. Sterilise as for nutrient bouillon.[Pg 200]

Media for the Study of Plant Bacteria.

Hay Infusion.—

1. Weigh out dried hay, 10 grammes, chop it up into fine particles and place in a flask.

2. Add 1000 c.c. distilled water, heated to 70° C.; close the flask with a solid rubber stopper.

3. Macerate in a water-bath at 60° C. for three hours.

4. Replace the stopper by a cotton-wool plug, and heat in the steamer at 100° C. for one hour.

5. Filter through Swedish filter paper.

6. Tube, and sterilise as for nutrient bouillon.

Haricot Bouillon.—(For cultivation of bacteria from tubercles of Legumes.)

1. Measure out 1000 c.c. distilled water into a 2-litre flask.

2. Weigh out 250 grammes haricot beans and add to the water in the flask.

3. Weigh out 10 grammes sodium chloride and add to the contents of the flask.

4. Add 1 c.c. of a 1 per cent. solution of sodium bicarbonate.

5. Place in the steamer at 100° C. for thirty minutes.

6. Filter.

7. Weigh out 20 grammes saccharose and add to the filtrate.

8. Tube, and sterilise as for nutrient bouillon.

Haricot Agar.—

1. Measure out 400 c.c. distilled water into a "tared" 2-litre flask.

2. Weigh out 15 grammes agar and mix into a thick paste with 100 c.c. cold distilled water, and add to the flask.

3. Dissolve the agar by bubbling live steam through the mixture as in making nutrient agar.

4. Weigh out 250 grammes haricot beans, place in the flask with the agar mixture.

5. Add 1 c.c. of 1 per cent. aqueous solution sodium bicarbonate.

6. Weigh out 10 grammes sodium chloride and add to the contents of the flask.

7. Place in the steamer at 100° C. for thirty minutes.

8. Adjust the weight of the medium mass to 1030 grammes (the figure per litre obtained experimentally) by the addition of distilled water at 100° C.[Pg 201]

9. Cool to 60°C., clarify with egg and filter.

10. Weigh out 20 grammes saccharose and add to the contents of the flask.

11. Tube, and sterilise as for nutrient agar.

Wood Ash Agar.—

1. Measure 400 c.c. distilled water into a tared 2-litre flask.

2. Weigh out 10 grammes agar and make into a thick paste with 100 c.c. cold distilled water.

3. Add this agar paste to the distilled water in the flask.

4. Dissolve the agar by passing live steam through it, as in preparing nutrient agar.

5. Weigh out 5 grammes clean wood ash and place in a second flask containing 200 c.c. distilled water with some sterile glass beads: shake thoroughly in a mechanical shaker for ten minutes.

6. Heat in steamer at 100°C., for thirty minutes.

7. After removal from the steamer dry the outside of the flask thoroughly, place it over a Bunsen flame and boil for one minute.

8. Filter directly into the flask containing the melted agar mixture.

9. Weigh out 4 grammes maltose. Add to the contents of the flask.

10. Adjust the weight of the medium mass to the calculated figure for one litre (1019 grammes) by the addition of distilled water at 100°C.

11. Replace the flask in the steamer for twenty minutes, cool to 60°C., and clarify with egg and filter.

12. Tube, and sterilise as for nutrient agar.

Media for the Study of Special Bacilli.

B. Acnes.

Oleic Acid Agar (Fleming).—

1. Measure out into a sterile stout glass bottle which already contains about 10 sterile glass beads

2. Weigh out

and add it to the ascitic fluid in the bottle.

3. Emulsify evenly by shaking (either by hand or in a shaking machine) for ten minutes.

4. Liquefy and measure out into a flask

then cool to 55°C.

5. Mix the oleic acid emulsion with the agar.[Pg 202]

6. Add 10 c.c. sterile neutral red, 1 per cent. aqueous solution.

7. Tube in quantities of 10 c.c., slant, and allow to set.

8. Incubate for forty-eight hours at 37° C. and reject any contaminated tubes. Store the sterile tubes for future use.

Coli-typhoid Group.

Parietti's Bouillon.—

1. Measure out pure hydrochloric acid, 4 c.c., and add to it carbolic acid solution (5 per cent.), 100 c.c. Allow the solution to stand at least a few days before use.

2. This solution is added in quantities of 0.1, 0.2. and 0.3 c.c. (delivered by means of a sterile graduated pipette) to tubes each containing 10 c.c. of previously sterilised nutrient bouillon (vide page 163).

3. Incubate at 37° C. for forty-eight hours to eliminate contaminated tubes. Store the remainder for future use.

Carbolised Bouillon.—

1. Prepare nutrient bouillon (vide page 163, sections 1 to 6). Measure out 1000 c.c.

2. Weigh out carbolic acid, 1 gramme (2.5 or 5 grammes may be needed for special purposes), and dissolve it in the medium.

3. Tube, and sterilise as for bouillon.

Carbolised Gelatine.—

1. Prepare nutrient gelatine (vide page 164, sections 1 to 7). Measure out 1000 c.c.

2. Weigh out carbolic acid, 5 grammes (= 0.5 per cent.), and dissolve it in the gelatine.

3. Filter if necessary through papier Chardin.

4. Tube, and sterilise as for nutrient gelatine.

One or 2.5 grammes of carbolic acid (= 0.1 per cent. or 0.25 per cent.) are occasionally used in place of the 5 grammes to meet special requirements.

Carbolised Agar.—

1. Prepare nutrient agar (vide page 167, sections 1 to 8). Measure out 1000 c.c.

2. Weigh out 1 gramme pure phenol and dissolve in the medium.

3. Filter if necessary through papier Chardin.

4. Tube, and sterilise as for nutrient agar.

Litmus Gelatine.—

1. Prepare nutrient gelatine (vide page 164, sections 1 to 8).

2. Add sterile litmus solution, sufficient to tint the medium a deep lavender colour.

3. Tube, and sterilise as for nutrient gelatine.[Pg 203]

Lactose Litmus Bouillon (Lakmus Molke).—

1. Weigh out peptone, 4 grammes, and emulsify it with 200 c.c. meat extract (vide page 148), previously heated to 60°C.

2. Weigh out salt, 2 grammes, and lactose, 20 grammes, and mix with the emulsion.

3. Wash the mixture into a sterile litre flask with 200 c.c. meat extract and add 600 c.c. distilled water.

4. Heat in the steamer at 100° C. for thirty minutes, to completely dissolve the peptone, etc.

5. Neutralise carefully to litmus paper by the successive additions of small quantities of decinormal soda solution.

6. Replace in the steamer for twenty minutes to precipitate phosphates, etc.

7. Filter through two thicknesses of Swedish filter paper.

8. Add sterile litmus solution, sufficient to colour the medium a deep purple.

9. Tube, and sterilise as for bouillon.

Lactose Litmus Gelatine (Wurtz).—

1. Prepare nutrient gelatine (vide page 164, sections 1 to 4).

2. Render the reaction of the medium mass -5.

3. Replace in the steamer at 100° C. for twenty minutes.

4. Clarify with egg as for gelatine.

5. Weigh out lactose, 20 grammes (= 2 per cent.), and dissolve it in the medium.

6. Filter through papier Chardin.

7. Add sufficient sterile litmus solution to colour the medium pale lavender.

8. Tube, and sterilise as for nutrient gelatine.

Lactose Litmus Agar (Wurtz).—

1. Prepare nutrient agar (vide page 167, sections 1 to 4).

2. Render the reaction of the medium mass -5.

3. Replace in the steamer at 100° C. for twenty minutes.

4. Cool to 60° C. and clarify with egg as for nutrient agar.

5. Weigh out lactose, 20 grammes (= 2 per cent.), and dissolve it in the medium.

6. Filter through papier Chardin, using the hot-water funnel.

7. Add sterile litmus solution, sufficient to colour the medium a pale lavender.

8. Tube, and sterilise as for nutrient agar.

Glycerine Potato Bouillon.—

1. Take 1 kilo of potatoes, wash thoroughly in water, peel, and grate finely on a bread-grater.

2. Weigh the potato gratings, place them in a 2-litre flask,[Pg 204] and add distilled water in the proportion of 1 c.c. for every gramme weight of potato. Allow the flask to stand in the ice-chest for twelve hours.

3. Strain the mixture through butter muslin and filter through Swedish filter paper into a graduated cylinder. Note the amount of the filtrate.

4. Place the filtrate in a flask, add an equal quantity of distilled water, and heat in the steam steriliser for sixty minutes.

5. Add glycerine, 4 per cent., mix thoroughly, and again filter.

6. Tube and sterilise as for nutrient bouillon.

Potato Gelatine (Elsner).—

1. Take 1 kilo of potatoes, wash thoroughly in water, peel, and finally grate finely on a bread-grater.

2. Weigh the potato gratings, place them in a 2-litre flask, and add distilled water in the proportion of 1 c.c. for every gramme weight of potato. Allow the flask to stand in the ice-chest for twelve hours.

3. Strain the mixture through butter muslin, and filter through Swedish filter paper into a graduated cylinder.

4. Add 15 per cent. gelatine to the potato decoction and bubble live steam through the mixture for ten minutes.

5. Estimate the reaction; adjust the reaction of the medium mass to +25.

6. Cool the medium to below 60°C.; clarify with egg as for nutrient gelatine (vide page 166).

7. Add 1 per cent. potassium iodide (powdered) to the medium.

8. Filter through papier Chardin.

9. Tube and sterilise as for nutrient gelatine.

Aesculin Agar.—(B. coli and allied organisms give black colonies surrounded by black halo.)

1. Measure out 400 c.c. distilled water into a tared 2-litre flask.

2. Weigh out

and make into a thick paste with 150 c.c. distilled water.

3. Add this paste to the distilled water in the flask.

4. Dissolve the ingredients by bubbling live steam through the mixture.

5. Weigh out

and dissolve in a second flask containing 100 c.c. distilled water.

6. Mix the contents of the two flasks—adjust the weight to[Pg 205] the calculated medium figure (in this case 1031.5 grammes) by the addition of distilled water at 100°C.

7. Clarify with egg and filter.

8. Tube and sterilise as for nutrient agar.

Bile Salt Agar (MacConkey).—

1. Weigh out powdered agar, 15 grammes (= 1.5. per cent.), and emulsify with 200 c.c. cold tap water.

2. Weigh out peptone, 20 grammes (= 2 per cent.), and emulsify with 200 c.c. tap water previously warmed to 60°C.

3. Mix the peptone and agar emulsions thoroughly.

4. Weigh out sodium taurocholate, 5 grammes (= 0.5 per cent.), dissolve it in 300 c.c. tap water, and use the solution to wash the agar-peptone emulsion into a tared 2-litre flask.

5. Bubble live steam through the mixture for twenty minutes.

6. Adjust the weight of the medium mass to the calculated figure for one litre (1040 grammes).

7. Cool to 60° C. and clarify with egg as for nutrient agar (vide page 168).

8. Filter through papier Chardin, using the hot-water funnel.

9. Weigh out lactose, 10 grammes (= 1 per cent.), and dissolve it in the agar.

If desired, add 5 c.c. of a 1 per cent. (= 0.5 per cent.) aqueous solution of neutral red.

10. Tube, and sterilise as for nutrient agar.

Litmus Nutrose Agar (Drigalski-Conradi).—

This medium should be prepared in precisely the same manner as the Nutrose agar described on page 172 substituting meat extract for serum water, and increasing the percentage of agar added per litre to 3 per cent.

Fuchsin Agar (Braun).—

1. Liquefy and measure out into a sterile flask:

2. Weigh out: lactose 10 grammes and dissolve in the fluid agar.

3. Adjust the reaction to -5 and filter.

4. Measure out and mix thoroughly with agar:

The fuchsin solution is prepared by mixing:

Allow to stand twenty-four hours, then centrifugalise thoroughly and decant the supernatant fluid into a well-stoppered bottle.[Pg 206]

5. Measure out and add to the nutrient agar, sodium sulphite, 10 per cent. aqueous solution, freshly prepared 25 c.c.

6. Tube and sterilise as for nutrient agar.

7. Store in a dark cupboard.

Fuchsin Sulphite Agar (Endo).—

1. Liquefy and measure out into a sterile flask:

2. Weigh out

and dissolve in the fluid agar.

3. Adjust the reaction to +3 and filter.

4. Measure out and mix thoroughly with the fluid agar.

5. Measure out and add to the medium

6. Tube and sterilise as for nutrient agar.

Brilliant Green Agar (Conradi).—

1. Liquefy and measure out into a sterile flask

2. Adjust reaction to +30 by the addition of normal phosphoric acid; and filter.

3. Measure out and mix thoroughly with the fluid medium

4. Measure out and add to the medium

5. Tube and sterilise as for nutrient agar.

Brilliant Green Bile Salt Agar (Fawcus).—

1. Weigh out agar 20 grammes and emulsify in 100 c.c. cold distilled water.

2. Wash the emulsion into a "tared" 2-litre flask with 500 c.c. distilled water.

3. Dissolve the agar by bubbling live steam through the flask.

4. Cool, clarify with egg and filter.

5. Weigh out

and add to the medium in the flask.[Pg 207]

6. Weigh out

and add to the medium in the flask.

7. Adjust reaction to +15 and filter if necessary.

8. Measure out

and mix thoroughly with the fluid agar.

9. Measure out and add to the medium

10. Tube and sterilise as for nutrient agar.

China Green Agar (Werbitski).—

1. Liquefy and measure out into a sterile flask

2. Adjust the reaction accurately to +13 and filter.

3. Measure out and mix thoroughly with the fluid agar

4. Tube and sterilise as for nutrient agar.

Malachite Green Agar (Loeffler).—

1. Liquefy and measure out into a sterile flask

2. Weigh out

and dissolve in nutrient agar.

3. Adjust the reaction to +3, and filter.

4. Measure out and mix thoroughly in the fluid agar

4a. To the filtered agar add

5. Tube and sterilise as for nutrient agar.

Double Sugar Agar (Russell).—

1. Liquefy and measure out into a sterile flask

2. Add 100 c.c. litmus solution to the fluid agar.

3. Weigh out and dissolve in the fluid agar.

[Pg 208]

4. Render the reaction of the medium neutral to litmus paper by the cautious addition of normal caustic soda.

5. Tube in quantities of 10 c.c. and sterilise in the steamer at 100° C. for twenty minutes on each of three successive days.

6. Store for use in a cool dark place.

B. Diphtheriæ.

Glycerine Blood-serum.—

1. Prepare blood-serum as described, page 168, sections 1 to 4.

2. Add 5 per cent. pure glycerine.

3. Complete as described above for ordinary blood-serum, sections 5 to 7.

Blood-serum (Loeffler).—

1. Prepare nutrient bouillon (vide page 163), using meat extract made from veal instead of beef.

2. Add 1 per cent. glucose to the bouillon, and allow it to dissolve completely.

3. Now add 300 c.c. clear blood-serum (vide page 168, sections 1 to 4) to every 100 c.c. of this bouillon.

4. Fill into sterile tubes and complete as for ordinary blood-serum.

Blood-serum (Lorrain Smith).—

1. Collect blood-serum (vide page 168, sections 1 to 4), as free from hæmoglobin as possible.

2. Weigh out 0.15 per cent. sodium hydrate and dissolve it in the fluid (or add 0.375 c.c. of dekanormal soda solution for every 100 c.c. of serum).

3. Tube, and stiffen at 100° C. in the serum inspissator.

4. Incubate at 37° C. for forty-eight hours to eliminate any contaminated tubes. Store the remainder for future use.

Blood Serum (Councilman and Mallory).—

1. Collect blood serum in slaughterhouse, coagulate, remove serum and tube (vide page 168).

Great care must be taken to avoid the inclusion of air bubbles—indeed if only a few tubes are filled at one time, it is a good plan to stand them upright in the receiver of an air pump and to exhaust as completely as possible before transferring to the serum inspissator.

2. Heat the tubes in a slanting position in hot-air steriliser at 90° C. till firmly coagulated, say half an hour.[Pg 209]

3. Sterilise in steam steriliser at 100° C. for 20 minutes on each of three successive days.

Resulting medium not translucent, but opaque and firm.

B. Tuberculosis.

Egg Medium (Lubenau).—

This modification of Dorset's egg medium (quod vide page 174) is preferred by some for the growth of the tubercle bacillus of the human type. It consists in the addition of one part of 6 per cent. glycerine in normal saline solution, to the egg mixture between steps 4 and 5.

Glycerine Bouillon.—

1. Measure out nutrient bouillon, 1000 c.c. (vide page 163, sections 1 to 6).

2. Measure out glycerine, 60 c.c. (= 6 per cent.), and add to the bouillon.

3. Tube, and sterilise as for bouillon.

Glycerine Agar.—

1. Prepare nutrient agar (vide page 167, sections 1 to 8). Measure out 1000 c.c.

2. Measure out pure glycerine, 60 c.c. (= 6 per cent.), and add to the agar.

3. Tube, and sterilise as for nutrient agar.

Glycerine Blood-serum.—

1. Prepare blood-serum as described, page 168, sections 1 to 4.

2. Add 5 per cent. pure glycerine.

3. Complete as described above for ordinary blood-serum, sections 5 to 7.

Glycerinated Potato.—

1. Prepare ordinary potato wedges (vide page 174, sections 1 to 4).

2. Soak the wedges in 25 per cent. solution of glycerine for fifteen minutes.

3. Moisten the cotton-wool pads at the bottom of the potato tubes with a 25 per cent. solution of glycerine.

4. Insert a wedge of potato in each tube and replug the tubes.

5. Sterilise in the steamer at 100° C. for twenty minutes on each of five consecutive days.[Pg 210]

Animal Tissue Media (Frugoni).—

1. Take a number of sterile test-tubes 16 × 3 or 4 cm., plugged with cotton wool, and into each insert a 2 cm. length of stout glass tubing (about 1 cm. diameter); fill in glycerine (6 per cent.) bouillon to the upper level of the piece of glass tubing. Sterilise in the steamer at 100° C. for twenty minutes on each of three successive days.

2. Kill a small rabbit by means of chloroform vapour.

3. Under strictly aseptic precautions remove the lungs, liver and other solid organs and transfer them to a sterile double glass dish.

4. With the help of sterile scissors and forceps divide the organs into roughly rectangular blocks 3 × 1.5 × 1 cm.

5. Pour into the dish a sufficient quantity of sterile glycerine solution (6 per cent. in normal saline), cover, and allow to stand for one hour.

6. Introduce a block of tissue into each tube so that it rests upon the upper end of the piece of glass tubing. (The surface of the tissue will now be kept moist by capillary attraction and condensation).

7. Sterilise in the autoclave at 120° C. for thirty minutes.

8. Cap the tubes and store them in the ice chest for future use.

Tissues obtained at postmortems can also be used after preliminary sterilisation by boiling or autoclaving.

Media for the Study of Special Cocci.

Diplococcus GonorrhϾ.

Ascitic Bouillon (Serum Bouillon).—

1. Collect ascitic fluid (pleuritic fluid, hydrocele fluid, etc.), by aspiration directly into sterile flasks, under strictly aseptic precautions.

2. Mix the serum with twice its bulk of sterile nutrient bouillon (vide page 163).

3. If considered necessary (on account of the presence of blood, crystals, etc.), filter the serum bouillon through porcelain filter candle.

4. Tube, and sterilise in the water bath at 56° C. for half an hour on each of five consecutive days.

5. Incubate at 37° C. for forty-eight hours and eliminate contaminated tubes. Store the remainder for future use.

Serum Agar (Heiman).—

1. Prepare nutrient agar (vide page 167), to following formula:

[Pg 211]

2. Make reaction of medium + 10.

3. Filter; tube in quantities of 6 c.c.

4. Sterilise as for nutrient agar.

5. After the third sterilisation cool the tubes to 42°C., and add to each 3 c.c. of sterile hydrocele fluid, ascitic fluid, or pleuritic effusion (previously sterilised, if necessary, by the fractional method); allow the tubes to solidify in a sloping position.

6. When solid, incubate at 37° C. for forty-eight hours, and eliminate any contaminated tubes. Store the remainder for future use.

Serum Agar (Wertheimer).—

1. Prepare nutrient agar (vide page 167), to the following formula:

2. Make reaction of medium +10.

3. Filter; tube in quantities of 5 c.c.

4. Sterilise as for nutrient agar.

5. After the last sterilisation cool to 42°C., then add 5 c.c. sterile blood-serum from human placenta (sterilised, if necessary, by the fractional method) to each tube; slope the tubes.

6. When solid, incubate at 37° C. for forty-eight hours, and eliminate any contaminated tubes. Store the remainder for future use.

Serum Agar (Kanthack and Stevens).—

1. Collect ascitic, pleuritic, or hydrocele fluid in sterile flasks and allow to stand in the ice-chest for twelve hours to sediment.

2. Decant 1000 c.c. of the clear fluid into a measuring cylinder and transfer to sterile litre flask.

3. Add 0.5 c.c. dekanormal NaOH solution for every 100 c.c. serum (i. e., 5.0 c.c.), and mix thoroughly.

4. Heat in the steamer for twenty minutes.

5. Weigh out 15 grammes agar, emulsify in a separate vessel with 200 c.c. of the alkaline fluid previously cooled to about 20°C., and then add to the remainder of the fluid in the flask.

6. Bubble live steam through the mixture for twenty minutes to dissolve the agar.

7. Filter through papier Chardin, using a hot-water funnel.

8. Weigh out glucose 10 grammes (= 1 per cent.), and dissolve it in the clear agar.

8a. If desired, add glycerine, 5 per cent., to the clear agar.

9. Tube, and sterilise as for nutrient agar.[Pg 212]

Serum Agar (Libman).—

1. Prepare nutrient agar (vide, page 167) using, however, 1.5 per cent. peptone (that is 15 grammes per litre instead of 10 grammes).

2. Adjust the reaction to 0 (i. e., neutral to phenolphthalein).

3. Filter and transfer 1000 c.c. liquefied medium to a sterile flask.

4. Weigh out dextrose 20 grammes and dissolve in the fluid agar.

5. Tube in quantities of 6 c.c.; and sterilise in the steamer at 100° C. for thirty minutes on each of three consecutive days.

6. After the third sterilisation cool to 42° C. and add to each tube 3 c.c. of sterile hydrocele fluid, ascitic fluid or pleuritic effusion (previously sterilised, if necessary, by the fractional method); allow the tubes to solidify in a sloping position.

7. When solid, incubate at 37° C. for forty-eight hours, and eliminate any contaminated tubes. Store the remainder for future use.

Egg-albumen, Inspissated.—

1. Break several fresh eggs (hens', ducks', or turkeys' eggs), and collect the "whites" in a graduated cylinder, taking care to avoid admixture with the yolks.

2. Add 40 per cent. distilled water, and incorporate the mixture thoroughly by the aid of an egg-whisk.

3. Weigh out 0.15 per cent. sodium hydrate and dissolve it in the fluid (or add the amount of dekanormal caustic soda solution calculated to yield the required percentage of soda in the total bulk of the fluid—i. e., 0.375 c.c. of dekanormal NaOH solution per 100 c.c. of the mixture).

3a. Glucose to the extent of 1 to 2 per cent. may now be added, if desired.

4. Strain the mixture through butter muslin and filter through a porcelain filter candle into a sterile filter flask.

5. Tube, and stiffen at 100° C. in the serum inspissator.

6. Incubate at 37° C. for forty-eight hours and eliminate any contaminated tubes; store the remainder for future use.

Egg-albumen (Tarchanoff and Kolesnikoff).—

1. Place unbroken hens' eggs in dekanormal caustic soda solution for ten days. (After this time the white becomes firm like gelatine.)

2. Carefully remove the shell and cut the egg into fine slices.

3. Wash for two hours in running water.

4. Place the egg slices in a large beaker and sterilise in the steamer at 100° C. for one hour.

5. Transfer each slice of egg by means of a pair of sterilised forceps to a Petri dish or large capsule.[Pg 213]

6. Sterilise in the steamer at 100° C. for twenty minutes on each of three consecutive days.

Egg Albumin Broth (Lipschuetz).—

1. Weigh out

and place in a 2-litre flask with a number of sterile glass beads.

2. Measure out distilled water 200 c.c. into a half-litre flask and warm to 37° C. in the incubator.

3. Add the water to the flask containing the albumin and beads and dissolve by shaking.

4. Add n/10-NaOH, 40 c.c. Allow the mixture to stand for thirty minutes with frequent shaking.

5. Filter through Swedish filter paper.

6. Sterilise by boiling two or three times at intervals of two hours.

7. Add ordinary nutrient bouillon 600 c.c.

8. Fill into small Erlenmeyer flasks in quantities of 50 c.c.

9. Incubate for forty-eight hours at 37°C.—discard any contaminated flasks and store the remainder for future use.

Egg Albumin Agar.—

1. Prepare egg albumin solution as above 1-6.

2. Liquefy and measure out ordinary nutrient agar 600 c.c. and add to the egg albumin solution (in place of the nutrient broth).

3. Complete as above 8-9.

Diplococcus Meningitidis Intracellularis.

Ascitic Fluid Agar (Wassermann) Synonym N-as-gar (Mervyn Gordon).

1. Liquefy and measure out into a sterile flask:

2. Measure out into a half litre flask

and add to it

3. Heat over a bunsen flame, shaking constantly until the fluid boils, and the nutrose is dissolved.

4. Add the nutrose ascitic solution to the fluid agar.

5. Heat in the steamer for thirty minutes, then filter.

6. Tube and sterilise as for nutrient agar.

[Pg 214]

Diplococcus Pneumoniæ.

Blood Agar (Washbourn).—

1. Melt up several tubes of nutrient agar (vide page 167) and allow them to solidify in the oblique position.

2. Place the tubes, in the horizontal position, in the "hot" incubator for forty-eight hours, to evaporate off some of the condensation water.

3. Kill a small rabbit with chloroform and nail it out on a board (as for a necropsy). Moisten the hair thoroughly with 2 per cent. solution of lysol.

4. Sterilise several pairs of forceps, scissors, etc., by boiling.

5. Reflect the skin over the thorax with sterile instruments.

6. Open the thoracic cavity by the aid of a fresh set of sterile instruments.

7. Open the pericardium with another set of sterile instruments.

8. Sear the surface of the left ventricle with a red-hot iron and remove fluid blood from the heart by means of sterile pipettes (e. g., those shown in Fig. 13, c).

9. Deliver a small quantity of the blood on the slanted surface of the agar in each of the tubes, and allow it to run over the entire surface of the medium.

10. Place the tubes in the slanting position and allow the blood to coagulate.

11. Return the "blood agar" to the hot incubator for forty-eight hours and eliminate any contaminated tubes. Store the remainder for future use.

Media for the Study of Mouth Bacteria Generally.

Potato Gelatine (Goadby).—

1. Prepare glycerine potato broth (see page 203, sections 1 to 5).

2. Add 10 per cent. gelatine to the potato decoction and bubble live steam through the mixture for ten minutes.

3. Estimate the reaction; adjust the reaction of the medium to +5.

4. Cool the medium to below 60°C., clarify with egg as for nutrient gelatine.

5. Filter through papier Chardin.

6. Tube, and sterilise as for nutrient gelatine.

Media for the Study of Protozoa.

Tissue Medium (Noguchi).—For spirochætes (cultivations must be grown anaerobically).

1. Plug and sterilise test-tubes 20 × 2 cm.

2. Kill a small rabbit with chloroform vapour. Open the abdomen[Pg 215] with all aseptic precautions, remove kidneys and testicles and transfer to a sterile glass dish. Cut up the organs with sterile scissors into small pieces—say 4 millimetre cubes. The four organs should yield from 25 to 30 pieces of tissue.

3. Drop a small piece of sterile tissue into the bottom of each sterilised tube.

4. Take a flask containing about 400 c.c. nutrient agar (+10 reaction), liquefy the medium by heat and cool in a water bath to 50°C.

5. Add 200 c.c. ascitic or hydrocele fluid (horse or sheep serum may be employed, but is not so good) to the liquid agar and mix carefully to avoid formation of air bubbles.

6. Fill about 20 c.c. of the ascitic agar into each of the sterilised tubes which already contains a piece of sterile rabbit's tissue, stand all the tubes upright in racks or a jar, and allow agar to set.

7. After solidification pour sterile paraffin oil on the surface of the medium in each tube to the depth of 3 centimetres.

8. Incubate tubes at 37° C. for several days and discard any which prove to be contaminated.

9. Store such tubes as are sterile for future use.

[Pg 216]

XIII. INCUBATORS.

Fig. 113.—Incubator.

An incubator (Fig. 113) consists essentially of a chamber for the reception of cultivations, etc., surrounded by a water jacket, the walls of which are of metal, usually copper, and outside all an asbestos or felt jacket, or wooden casing. The water in the jacket is heated by gas or electricity and maintained at some constant temperature by a thermo-regulator. The cellular incubator (Fig. 114) which was made for me[7] some years ago is of the greatest practical utility. Here the[Pg 217] central cavity is subdivided by five double-walled partitions (in which water circulates in connection with the water tanks at the top and base of the incubator) and again by iron shelves to form twenty-four pigeon holes. Into each of these slides an iron drawer 35 cm. long × 12 cm. wide × 22 cm. high forming a self-contained incubator. The drawer is fitted with a wooden form to which is fixed a handle and a numbered label. The thermo-regulating apparatus is the well-known Hearson capsule.

Fig. 114.—Cellular incubator.

Two incubators at least are required in the laboratory, for the cultivation of bacteria the one regulated to maintain a temperature of 37°C., and known as the "hot" incubator; the other, 20° C. to 22°C., and known as the "cool" or "cold" incubator.

Two other incubators, regulated to 42° C. and 60°C. respectively, whilst not absolutely, necessary very soon justify their purchase.

Thermo-regulators.—The thermo-regulator is the[Pg 218] most essential portion of the incubator, as upon its efficient working depends the maintenance of a constant temperature in the cultivation chamber. It is also used in the fitting up of water and paraffin baths, and for many other purposes.

Fig. 115.—Reichert's thermo-regulator.

Of the many forms and varieties of thermo-regulator (other than electrical), two only are of sufficiently general use to need mention. In one of these the flow of gas to the gas-jet is controlled by the expansion or contraction of mercury within a glass bulb; in the other, by alterations in the position of the walls of a metallic capsule containing a fluid, the boiling-point of which corresponds to the temperature at which the incubator is intended to act. They are:

(a) Reichert's (Fig. 115), consists of a bulb containing mercury which is to be suspended in the medium, whether air or water, the temperature of which it is desired to regulate. Gas enters at A, and passes out to the jet by B. As the temperature rises the mercury expands and cuts off the main gas supply. As the temperature falls the mercury contracts and reopens the narrow tube C. By means of a thumbscrew D (which mechanically raises or lowers the column of mercury irrespective of the temperature) and the aid of a thermometer the apparatus can be set to keep the incubator at any desired temperature. With this form a special gas burner is required, with separate supply of gas to a pilot jet at the side.

(b) Hearson's capsule regulator consists of a metal capsule hermetically sealed and filled with a liquid which boils at the required temperature, this is adjusted in the interior of the incubator. Soldered to the upper side of the capsule is a thick piece of metal having a central[Pg 219] cup to receive the lower end of a rigid rod, through which the movements of the walls of the capsule are transmitted to the gas valve fixed outside the incubator.

The gas valve or governor is shown in figure 116. A is the inlet for gas, C the outlet to burner heating the water jacket, B D a lever pivoted to standards at G, and acted upon by the capsule, through the rigid rod which enters the socket below the screw P.

Fig. 116.—Capsule thermo-regulator.

The construction of the valve is such that, whenever the short arm of the lever B D presses on the disc below the end B, the main supply of gas is entirely cut off. At such times, however, a very small portion of gas passes from A to C, through an aperture inside the valve, the size of which aperture can be adjusted by the screw needle S, hence the gas flame below the incubator is never extinguished.

The expansion of the metal walls of the capsule, which takes place upon the boiling of its contents, provides the motive force, transmitted through the rigid rod to raise the long arm of the lever B D, and as this expansion only takes place at a predetermined temperature, the lever will only be acted upon when the critical temperature is reached, no sensible effect being produced at even 1° C. below that at which the capsule is destined to act.

W is a weight sliding on the lever rod D; by increasing the distance between the weight and the fulcrum[Pg 220] of the lower increased pressure is brought to bear upon the walls of the capsule with the result that the boiling-point of the liquid in the capsule is slightly raised, and a range of about two degrees can thus be obtained with any particular capsule.

[Pg 221]

XIV. METHODS OF CULTIVATION.

Cultivations of micro-organisms are usually prepared in the laboratory in one of three ways:

These may be incubated either aerobically (i. e., in the presence of oxygen) or anaerobically (i. e., in the absence of oxygen, or in the presence of an indifferent gas, such as hydrogen, nitrogen, or carbon dioxide).

With regard to the temperature at which the cultivations are grown, it may be stated as a general rule that all media rendered solid by the addition of gelatine are incubated at 20°C., or at any rate at a temperature not exceeding 22° C. (that is, in the "cold" incubator); whilst fluid media and all other solid media are incubated at 37° C. (that is, in the "hot" incubator). Exceptions to this rule are numerous. For instance, in studying the growth of the psychrophylic bacteria, the yeasts and the moulds, the cold incubator is employed for all media.

Tube cultivations are usually packed in the incubator in small tin cylinders, such as those in which American cigarettes are sold, or in square tin boxes. Beakers or tumblers may be used for the same purpose, but being fragile are not so convenient. Metal test-tube racks, long enough to just fit into the interior of the incubator and each accommodating two rows of tubes, are also exceedingly useful.[Pg 222]

AEROBIC.

The Preparation of Tube Cultivations.

The preparation of a tube cultivation consists in:

(a) Inoculating a tube of sterile nutrient medium with a portion of the material to be examined.

(b) Incubating the inoculated tube at a suitable temperature.

The details of the first of these processes must be varied somewhat according to whether the tubes of nutrient media are inoculated or "planted" from—

1. Pre-existing cultivations.

2. Morbid material previously collected (vide page 373).

3. Fluids, tissues, etc., or from the animal body direct.

The method of preparing tube cultivations from pre-existing cultivations is as follows:

Fig. 117.—Inoculating tubes, seen from the front.

1. Fluid Media (e. g., Nutrient Bouillon).—

1. Flame the cotton-wool plug of the tube containing the cultivation and also that of the tube of sterile bouillon.

2. Hold the two tubes, side by side, between the left thumb and the first and third fingers, allowing the sealed ends to rest on the dorsum of the hand, and separating the mouths of the tubes (which are pointed to the right) by the tip of the second finger. Keep[Pg 223] the tubes as nearly horizontal as is possible without allowing the fluid in the bouillon tube to reach the cotton-wool plug (Fig. 117).

3. Sterilise the platinum loop and allow it to cool.[8]

4. Grasp the plug of the tube containing the cultivation between the little finger and palm of the hand and remove it from the tube.

5. Grasp the plug of the bouillon tube between the fourth finger and the ball of the thumb and remove it from the tube.

6. Pass the platinum loop into the tube containing the culture—do not allow the loop to touch the sides of the tube, or the handle to touch the medium—and remove a small portion of the growth; withdraw the loop from the tube, keeping the infected side of the loop downward.

7. Pass the loop into the bouillon tube almost down to the level of the fluid, reverse the loop so that the infected side faces upward, emulsify the portion of the growth in the moisture adhering to the side of the tube which is uppermost. Withdraw the loop.

8. Replug both tubes.

9. Sterilise the platinum loop.

10. Label the bouillon tube with (a) the name of the organism and (b) the date of inoculation.

11. Incubate.

2. Solid Media.—Solid media are stored in tubes in one of two ways:

1. Oblique tube or slanted tube (Fig. 118), in which the medium has been allowed to solidify whilst the tube was retained in an inclined position, so forming an extensive surface of medium extending from the bottom of the tube almost to its mouth.

This is employed for "streak" or "smear" cultivations (Strichcultur).

2. Straight tube (Fig. 119), in which the medium[Pg 224] forms a cylindrical mass in the lower portion of the tube and presents an upper surface which is at right angles to the long axis of the tube.

This is employed for "stab" or "stick" cultivations (Stichcultur), or by inoculating the medium whilst fluid, and allowing to solidify in this position, for "shake" cultivations.

Streak Culture.—

1. Flame the plugs, sterilise the platinum loop (or spatula). Open the tubes and charge the loop as in previous inoculation.

2. Pass the infected loop to the bottom of the tube to be inoculated and draw it, as lightly as possible, along the centre of the surface of the medium, terminating the "streak" over the thin layer of medium near the mouth of the tube.

3. Replug the tubes, sterilise the platinum loop.

4. Label the newly inoculated tube and incubate.

Smear Culture.—Proceed generally as in streak culture, but rub the infected loop all over the surface of the medium, instead of restricting the inoculation to a narrow line.

Stab Culture.—

1. Flame the plugs, open the tubes, sterilise the platinum needle and charge it with the inoculum as in the previous cultivations.

2. Pass the platinum needle into the tube to be inoculated until it touches the centre of the surface of the medium. Now thrust it deeply into the substance of the medium, keeping the needle as nearly as possible in the axis of the cylinder of medium. Then withdraw the needle.

3. Replug the tubes. Sterilise the platinum needle.[Pg 225]

4. Label the newly planted tube and incubate.

Fig. 118.—Sloped or slanted medium for streak or smear culture.

Fig. 119.—Straight tube.

Shake Culture.—

1. Liquefy a tube of nutrient gelatine (or agar, or other similar medium), by heating in a water-bath (Fig. 121).

2. Inoculate the liquefied medium and label it, etc., precisely as if dealing with a tube of bouillon.[Pg 226]

3. Place the newly planted tube in the upright position (e. g., in a test-tube rack) and allow it to solidify.

4. Label the tube; when solid, incubate.

Esmarch's Roll Cultivation.—

1. Liquefy three tubes of gelatine by heat.

2. Prepare three dilutions of the inoculum (as described for plate cultivations, page 228, steps 4 to 7).

3. Roll the tubes, held almost horizontally, in a groove made in a block of ice, until the gelatine has set in a thin film on the inner surface of tube (Fig. 120); or under the cold-water tap.

Fig. 120. Esmarch's roll culture on block of ice.

In order that the medium may adhere firmly to the glass, the agar used for roll cultivation should have 1 per cent. gelatine or 1 per cent. gum arabic added to it before sterilisation.

Roll cultivations, which served a most important purpose in the days before the introduction of Petri dishes for plate cultivations, are now obsolete in modern laboratories and are merely mentioned for the benefit of students, since examiners who are interested in the academic and historical aspects of bacteriology sometimes expect candidates to be acquainted with the method of preparing them.

The Preparation of Plate Cultures.

If a small number of bacteria are suspended in liquefied gelatine, agar, or other similar medium, and the infected medium spread out in an even layer over a flat surface and allowed to solidify, each individual micro-organism becomes fixed to a certain spot and its further development is restricted to the vicinity of this spot. After a variable interval the growth of this[Pg 227] organism becomes visible to the naked eye as a "colony." This is the principle upon which the method of plate cultivation is based and its practice enables the bacteriologist to study the particular manner of development affected by each species of microbe when growing (a) unrestricted upon the surface of the medium, (b) in the depths of the medium. The method itself is as follows:

Fig. 121.—Handy form of water-bath for melting tubes of agar and gelatine previous to slanting them; or to making shake cultures or pouring plates.

Method of "Pouring" Plates.—

1. Place the glass dish on the levelling tripod (Figs. 122, 123); if gelatine plates are to be poured fill the dish with ice water—gelatine solidifies so slowly that it is necessary to hasten the process; if agar is to be used fill with water at 50°C.—agar sets almost immediately at the room temperature and by slightly retarding the process lumpiness is avoided; cover the dish with the square sheet of glass.

2. Place the spirit level on the sheet of glass and by means of the levelling screws adjust the surface of the glass to the horizontal.[Pg 228]

This leveling is an important matter since the development of a colony is to some extent proportionate to the supply of medium available for its nutrition. Thus in a "smear" on sloped tube culture, the colonies at the upper part of the medium are stunted and small but increase in size and luxuriance of growth the nearer they approach to the bottom of the tube, where there is the greatest depth of medium.

Fig. 122.—Plate-levelling stand.

3. Place three sterile Petri dishes in a row on the surface of the glass plate and number them 1, 2, and 3, from left to right.

Fig. 123.—Plate-levelling stand, side view.

4. Number the previously liquefied tubes of nutrient media 1, 2, and 3. Flame the plugs and see that each plug can be readily removed from the mouth of its tube.

5. Add one loopful of the inoculum to tube No. 1,[Pg 229] treating the liquefied medium as bouillon. After replugging, grasp the tube near its mouth by the thumb and first finger of the right hand, and with an even circular movement of the whole arm, diffuse the inoculum throughout the medium; avoid jerky movements, as these cause bubbles of air to form in the medium.

Fig. 124.—Mixing emulsion for plates.

The knack of mixing evenly without producing air bubbles, is not always easily acquired, by this method. An alternative plan is to hold the inoculated tube vertically upright between the opposed palms and to rotate it between them by rapid backward and forward movements of the two hands (Fig. 124).

Fig. 125.—Pouring plates.

6. Sterilise the platinum loop, and add two loopfuls of diluted inoculum to tube No. 2, and mix as before.

7. In a similar manner transfer three loopfuls of liquefied medium from tube No. 2 to tube No. 3, and mix thoroughly.

8. Flame the plug of tube No. 1, remove it, then flame the lips of the tube; slightly raise the cover of Petri dish No. 1, introduce the mouth of the tube; then,[Pg 230] elevating the bottom of the tube, pour the liquefied medium into the Petri dish, to form a thin layer. Remove the mouth of the tube and close the "plate." If the medium has failed to flow evenly over the bottom of the plate, raise the plate from the levelling platform and by tilting in different directions rectify the fault.

9. Pour plates No. 2 and No. 3, in a similar manner, from tubes Nos. 2 and 3.

10. Label the plates with the distinctive name or number of the inoculum, also the date; the number of the dilution having been previously indicated (step 3).

11. Place in the cool incubator for three or more days, as may be necessary.

In this way colonies may be obtained quite pure and separate from each other.

In plate No. 1, probably, the colonies will be so numerous and crowded, and therefore so small, as to render it useless. In plate No. 2 they will be more widely separated, but usually No. 3 is the plate reserved for careful examination, as in this the colonies are usually widely separated, few in number, and large in size.

Agar plates are poured in a similar manner, but the agar must be melted in boiling water and then allowed to cool to 45° C. or 42° C. in a carefully regulated water-bath before being inoculated, and the entire process must be carried out very rapidly, otherwise the agar will have solidified before the operation is completed.

Surface Plates.—

This method of pouring what may be termed "whole" plates (since colonies may appear both on the surface and in the depths of the medium) is essential to the accurate study of the formation of colonies under[Pg 231] various conditions, but when the main object of the separation of the bacteria is to obtain subcultivations from a number of individual bacteria, "surface" plates must be prepared, since here colony formation is restricted to the surface of the medium. The method adopted varies slightly according to whether the medium employed is gelatine or agar, or one of the derivatives or variants of the latter.

(a) Gelatine Surface Plates.—

1. Liquefy three tubes of nutrient gelatine.

2. Pour each tube into a separate Petri dish and allow it to solidify. Then turn each plate and its cover upside down.

Fig. 126.—Surface plate spreader.

3. When quite cold raise the bottom of plate 1, revert it and deposit a drop of the inoculum (whether a fluid culture or an emulsion from solid culture) upon the surface of the gelatine with a platinum loop—close to one side of the plate; replace the bottom half of the Petri dish in its cover.

4. Take a piece of thin glass rod, stout platinum wire or best of all a piece of aluminium wire (say 2 mm. diameter) about 28 cm. long. Bend the terminal 4 cm. at right angles to the remainder, making an L-shaped rod (Fig. 126). Sterilise the short arm and adjacent portion of the long arm, in the Bunsen flame, and allow it to cool.

5. Now raise the bottom of the Petri dish in the left hand, leaving the cover on the laboratory bench, and holding it vertically, smear the drop of inoculum all over the surface of the gelatine with the short arm of the spreader by a rotatory motion, (Fig. 127). Replace the dish in its cover.

6. Raise the bottom of plate 2 and rub the infected[Pg 232] spreader all over the surface of the gelatine—then go on in like manner to the third plate in the series.

7. Sterilise the spreader.

8. Label and incubate the plates.

Fig. 127.—Spreading surface plate.

After incubation, plate No. 1 will probably yield an enormous number of colonies; plate 2 will show fewer colonies, since only those bacteria adhering to the rod after rubbing over plate 1 would be deposited on its surface, and by the time the rod reached plate 3 but very few organisms should remain upon it. So that the third plate as a rule will only show a very few scattered colonies, eminently suitable for detailed study.

(b) Agar Surface Plates.—

1. Liquefy three tubes of nutrient agar—nutrose agar or the like.

2. Pour each tube into a separate Petri dish and allow it to solidify.

3. When quite solid invert each dish, raise the bottom half and rest it obliquely on its inverted cover (Fig. 128) and place it in this position in an incubator at 60° C. for forty-five minutes (or in an incubator at 42° C. for[Pg 233] two hours). This evaporates the water of condensation and gives the medium a firm, dry surface.

4. On removing the plates from the incubator close each dish and place it—still upside down—on the laboratory bench.

Fig. 128.—Drying surface plate of agar.

5. Inoculate the plates in series of three, as described for gelatine surface plates 3-8.

Hanging-drop Cultivation.

(a) Fluid Media.—

1. Prepare first and second dilutions of the inoculum as directed for plate cultivations (vide pages 228-229, sections 4 to 6), substituting tubes of nutrient broth for the liquefied gelatine.

2. Sterilise a hanging-drop slide by washing thoroughly in water and drying, then plunging it into a beaker of absolute alcohol, draining off the greater part of the spirit, grasping the slide in a pair of forceps, and burning off the remainder of the alcohol in the flame.

3. Place the hanging-drop slide on a piece of blotting paper moistened with 2 per cent. lysol solution and[Pg 234] cover it with a small bell glass that has been rinsed out with the same solution and not dried.

4. Raise the bell glass slightly and smear sterile vaseline around the rim of the metal cell by means of a sterile spatula of stout platinum wire.

5. Remove a clean cover-slip from the alcohol pot with sterile forceps and burn off the alcohol; again raise the bell glass and place the sterile cover-slip on the blotting paper by the side of the hanging-drop slide.

6. Remove a drop of the broth from the second dilution tube with a large platinum loop; raise the bell glass and deposit the drop on the centre of the cover-slip. Sterilise the loop.

7. Raise the bell glass sufficiently to allow of the cover-slip being grasped with forceps, inverted, and adjusted over the cell of the hanging-drop slide. Remove the bell glass altogether and press the cover-slip firmly on to the cell.

8. Either incubate and examine at definite intervals, or observe continuously with the microscope, using a warm stage if necessary (Fig. 53).

(b) Solid Media.—Observing precisely similar technique, a few drops of liquefied gelatine or agar from the second dilution tube may be run over the surface of the sterile cover-slip and a hanging-drop plate cultivation thereby prepared.

This method is extremely useful in connection with the study of yeasts, when the circular cell on the hanging-drop slide should be replaced by a rectangular cell some 38 by 19 mm., and the gelatine spread over a cover-slip of similar size. After sealing down the preparation, the upper surface of the cover-slip may be ruled into squares by the aid of the grease pencil or a writing diamond and numbered to facilitate the subsequent identification of the colonies which are observed to develop from solitary germs.[Pg 235]

Hanging-block Culture (Hill).—

Apparatus required: As for hanging-drop cultivation with the addition of a scalpel.

Carry out the method as far as possible under cover of a bell glass, to avoid aerial contamination.

1. Liquefy a tube of nutrient agar (or gelatine) and pour into a Petri dish to the depth of about 4 mm. and allow to set.

2. With a sharp scalpel cut out a block some 8 mm. square, from the entire thickness of the agar layer.

3. Raise the agar block on the blade of the scalpel and transfer it, under side down, to the centre of a sterile slide.

4. Spread a drop of fluid cultivation (or an emulsion of growth from a solid medium) over the upper surface of the agar block as if making a cover-slip film.

5. Place the slide and block covered by the bell glass in the incubator at 37° C. for ten minutes to dry slightly.

6. Take a clean dry sterile cover-slip in a pair of forceps, and with the help of a second pair of forceps lower it carefully on the inoculated surface of the agar (avoiding air bubbles), so as to leave a clear margin of cover-slip overlapping the agar block.

7. Invert the preparation and with the blade of the scalpel remove the slide from the agar block.

8. With a platinum loop run a drop or two of melted agar around the edges of the block. This solidifies at once and seals the block to the cover-slip.

9. Prepare a sterile hanging-drop slide, and smear hard vaseline or melted white wax on the rim of the metal cell.

10. Invert the cover-slip with the block attached on to the hanging-drop slide, and seal the cover-slip firmly in place.

11. Observe as for hanging-drop cultivations.[Pg 236]

ANAEROBIC CULTIVATIONS.

Numerous methods have been devised for the cultivation of anaerobic bacteria, the majority requiring the employment of special apparatus. The principle upon which any method is based and upon which it depends for its success falls under one or another of the following headings:

(a) Exclusion of air from the cultivation.

(b) Exhaustion of air from the vessel containing the cultivation by means of an air pump—i. e., cultivation in vacuo.

(c) Absorption of oxygen from the air in contact with the cultivation by means of pyrogallic acid rendered alkaline with caustic soda—i. e., cultivation in an atmosphere of nitrogen.

(d) Displacement of air by an indifferent gas, such as hydrogen or coal gas—i. e., cultivation in an atmosphere of hydrogen.

(e) A combination of two or more of the above methods.

A selection of the simplest and most generally useful methods is given here.

Whenever possible, the nutrient media that are employed in any of the processes should contain some easily oxidisable substance, such as sodium formate (0.4 per cent.) or sodium sulphindigotate (0.1 per cent.), which will absorb all the available oxygen held in solution by the medium. The further addition of glucose, 2 per cent., favors the growth of anaerobic bacteria (vide, pages 189-190).

Further, it is advisable to seal all joints between india-rubber stoppers and tubulures or the mouths of the tubes with melted paraffin; glass stoppers and taps should be lubricated with resin ointment or a mixture of beeswax 1 part, olive oil 4 parts.[Pg 237]

(A) Method I (Hesse's Method).—

1. Make a stab culture in gelatine or agar, choosing for the purpose a straight tube containing a deep column of medium, and thrusting the inoculating needle to the bottom of the tube.

2. Pour a layer of sterilised oil (olive oil, vaseline, or petroleum), 1 or 2 cm. deep, upon the surface of the medium.

3. Incubate.

Method II.—This method is only available when dealing with pure cultivations.

1. Liquefy a tube of gelatine (or agar) by heat, pour it into a Petri dish, and allow it to solidify.

2. Inoculate the surface of the medium in one spot only.

3. Remove a cover-slip from the pot of absolute alcohol with sterile forceps; burn off the alcohol in the gas flame.

4. Lower the now sterile cover-slip carefully on to the inoculated surface of the medium, carefully excluding air bubbles, and press it down firmly with the points of the forceps. (A sterile disc of mica may be substituted for the cover-slip.)

5. Incubate.

Method III (Roux's Physical Method).—

1. Prepare tube cultures of fluid media (or solid media rendered fluid by heat) in the usual way.

2. Aspirate some of the inoculated media into capillary pipettes.

3. Seal both ends of each pipette in the blowpipe flame.

4. Incubate.

Method IV (Roux's Biological Method).—

1. Plant a deep stab, as in method I.

2. Pour a layer, 1 or 2 cm. deep, of broth cultivation of a vigourous aerobe—e. g., B. aquatilis sulcatus or B.[Pg 238] prodigiosus—upon the surface of the medium; or an equal depth of liquefied gelatine, which is then inoculated with the aerobic organism.

3. Incubate.

The growth of the aerobe will use up all the oxygen that reaches it and will not allow any to pass through to the medium below, which will consequently remain in an anaerobic condition.

(B) Method V.—

1. Prepare tube or flask cultivations in the usual way.

2. Replace the cotton-wool plug by an india-rubber stopper perforated with one hole and fitted with a length of glass tubing which has a constriction about 3 cm. above the stopper and is then bent at right angles (Fig. 129). The stopper and glass tubing are sterilised by being boiled in a beaker of water for five minutes.

Fig. 129.—Vacuum culture.

3. Connect the tube leading from the culture vessel with a water or air pump, interposing a Wulff's bottle fitted as a wash-bottle and containing sulphuric acid.

4. Exhaust the air from the culture vessel.

5. Before disconnecting the apparatus, seal the glass tube from the culture vessel at the constriction, using the blowpipe flame.

6. Incubate.

(C) Method VI (Buchner's Method).

Apparatus and Solutions Required.—

[Pg 239]

Method.—

1. Prepare the tube cultivation in the usual way.

2. Moisten the india-rubber stopper of the Buchner's tube with water and see that it fits the mouth of the tube accurately.

3. Remove the stopper from the caustic soda bottle.

4. Drop one of the pyrogallic acid tablets[9] into the Buchner's tube (roughly, use 1 gramme pyrogallic acid for every 100 c.c. air capacity of the receiving vessel).

5. Add about 1 c.c. of the soda solution.

6. Place the inoculated tube inside the Buchner's tube. The pyrogallic tablet acts as a buffer and prevents damage to either the inoculated tube or the Buchner's tube even should it be slipped in hurriedly.

7. Fit the india-rubber stopper tightly into the mouth of the Buchner's tube.

Fig. 130.—Buchner's tube.

The pyrogallic acid tablet dissolves slowly in the soda solution and its oxidation proceeds very slowly at first so that ample time is available when this method is adopted.

8. Restopper the caustic soda bottle.

9. Place Buchner's tube in a wire support, and incubate.

Method VII (Wright's Method).—

1. Prepare tube cultivation in the usual way.

2. Cut off that portion of the cotton-wool plug projecting above the mouth of the tube with scissors, then push the plug into the tube for a distance of 2 or 3 cm.[Pg 240]

3. By means of a pipette drop about 1 c.c. of pyrogallic acid 10 per cent. aqueous solution on to the plug. It will immediately be absorbed by the cotton-wool.

4. With another pipette run in an equal quantity of the caustic soda solution.

5. Quickly close the mouth of the tube with a tightly fitting india-rubber stopper.

6. Incubate.

Fig. 131.—McLeod's anaerobic plate base with half petri dish inverted in situ

Method VIII (McLeod's Method).—

Apparatus and Solutions Required.—

Method.—

1. Roll out a long cylinder of plasticine and fit it into the groove on the upper surface of the earthenware base.[Pg 241]

2. Place a tablet of pyrogallic acid in one division of the interior of the plate base, and two tablets of sodic hydroxide in the other.

3. Prepare surface plate culture of the organism to be cultivated.

4. Run a few cubic centimetres of distilled water into that division of the plate base containing the sodic hydroxide.

5. Invert the bottom half of the surface plate over the plate base and press its edges firmly down into the plasticine filling the groove.

6. Label and incubate.

(D) Method IX.—

Apparatus Required.—

Method.—

1. Sterilise a glass vessel, shaped as in a Ruffer's or Woodhead's flask, in the hot-air oven. (The tubulure and the side tubes are plugged with cotton-wool.) After sterilisation, fix a short piece of rubber tubing occluded by a metal clip to each side tube.

2. Inoculate a large quantity (e. g., 200 c.c.) of the medium. Where solid media are employed they must first be liquefied by heat.

3. Remove the cotton-wool plug from the tubulure and pour the inoculated medium into the glass vessel.

4. Close the tubulure by means of an india-rubber stopper previously sterilised by boiling in a beaker of water.[Pg 242]

Fig. 132.—Kipp's hydrogen apparatus, (a) connected up to two washing bottles containing (b) lead acetate 10 per cent. solution, to remove H2S and (c) silver nitrate solution to remove AsH3. A third washing bottle containing pyrogallic acid 10 per cent. solution, rendered alkaline, to remove any trace of oxygen, is sometimes introduced.

Fig. 133.—Improved gas apparatus; the metal is contained in a perforated glass tube which is submerged in acid when the triangular bottle is upright (a), but is above the level of the liquid when the bottle is turned on its side (b).

[Pg 243]

5. Connect up the india-rubber tubing on one of the side tubes with a cylinder of compressed hydrogen (or the delivery tube of a Kipp's Fig. 132 or other hydrogen apparatus, Fig. 133), interposing a short piece of glass tubing; and in like manner connect a long piece of rubber tubing which should be led into a basin of water, to the opposite side tube.

6. Open both metal clips and pass hydrogen through the vessel until the atmospheric air is replaced by hydrogen. This is determined by collecting some of the gas which bubbles through the water in the basin in a test-tube and testing it by means of a lighted taper.

7. Close the metal clip on the tube through which the gas is entering; close the clip on the exit tube.

8. Disconnect the gas apparatus.

9. Incubate.

Method X (Botkin's Method).—

Apparatus Required.—

Method.—

1. Place the leaden cross inside the glass dish, resting on the bottom.

2. Prepare the cultivations in the usual way.

3. Place the tube cultivations in a glass battery jar (or the plate cultivations on a metal frame), resting on the centre of the leaden cross.

4. Cover the cultivations with the bell jar.

5. Adjust the U-shaped pieces of glass tubing in a vertical position on opposite sides of the bell jar, one arm of each inside the jar, the other outside. These tubes are best held in position by embedding the U-shaped[Pg 244] bends in two lumps of plasterine stuck on the bottom of the glass dish. Fix a short length of rubber tubing clamped with a metal clip to each of the outside arms (Fig. 134).

6. Fill the glass dish with glycerine or metallic mercury to a depth of about 5 cm.

Fig. 134.—Botkin's apparatus.

7. Connect up one U-shaped tube with the hydrogen cylinder (or gas apparatus) by means of rubber tubing. Replace the atmospheric air by hydrogen, as in method IX.

8. Clamp the tubes and disconnect the gas apparatus.

9. Incubate.

Method XI (Novy's Method).—

Apparatus Required.—

Method.—

1. Prepare cultivations in the usual way.

2. Place these inside the jar.

3. Lubricate the stopper and insert it in the mouth of the jar, with the handle in a line with the two side tubes.

4. Connect up the delivery tube a with the hydrogen gas supply by means of rubber tubing.

Fig. 135.—Novy's jar for plate cultivations.

Fig. 136.—Novy's jar for tube cultivations.

5. Attach a piece of rubber tubing to the exit tube b and collect samples of the issuing gas (over water) and test from time to time.

6. When the air is completely displaced by hydrogen, turn the handle of the stopper at right angles to the line of entry and exit tubes; this seals the orifice of both tubes.

7. Disconnect the gas apparatus and incubate.

(E) Method XII (Bulloch's Method).—

Apparatus Required.—

Method.—

1. Prepare the cultivations in the usual way.

2. Place the glass dish in the centre of the glass slab, and stand the cultivations inside this.

3. Place a sufficient number of pyrogallic acid tablets at one side of the glass dish (i. e., 1 tablet for each 100 cubic centimeters air capacity of the bell jar). Place a small heap of dry granulated soda (or half a dozen tablets of sodic hydroxide) by the side of the pyro tablets.

4. Smear the flange of the bell jar with resin ointment and apply the jar firmly to the glass slab, covering the cultivations—so arranged that the long tube passes with its lower end into the glass dish at a point directly opposite to the pyrogallic acid tablets. Lubricate the two stop-cocks with resin ointment (Fig. 137).

5. Connect up the short tube a with the gas-supply by means of rubber pressure tubing and open both stop-cocks.

6. Connect a long, straight piece of glass tubing to the long tube b by means of a piece of rubber tubing interposing a screw clamp: and collect samples of the issuing gas from time to time and test.

7. When the air is displaced, shut off the stop-cock of the entry tube, then that of the exit tube b. Screw down the clamp and remove the glass tube from the rubber connection and connect up the short tube a to the air pump by means of pressure tubing.

8. Open the stop-cock of tube a and with two or three[Pg 247] strokes of the air pump, aspirate a small quantity of gas, so creating a slight vacuum. Then shut off the stop-cock and disconnect the air pump.

9. Fill the 10 c.c. bulb pipette with water; insert its point into the rubber tubing on the long tube b as far as the screw clamp. Open the screw clamp and run in water until stopped by the internal pressure. Shut off stop-cock. (The water dissolves the soda and pyrogallic acid converting the latter into alkaline pyro. and so bringing its latent capacity for oxygen into action).

Fig. 137.—Bulloch's jar.

10. Reverse the tubes from the tubulures so that they meet, out of harm's way, over the top of the bell glass; again see that all joints are tight and transfer the apparatus to the incubator.

This last method is the most satisfactory for anaerobic cultivations, as by its means complete anaerobiosis can be obtained with the least expenditure of time and trouble.

[Pg 248]

XV. METHODS OF ISOLATION.

The work in the preceding sections, arranged to demonstrate the chief biological characters of bacteria in general, is intended to be carried out by means of cultivations of various organisms previously isolated and identified and supplied to the student in a state of purity. A cultivation which comprises the progeny of a single cell is termed a "pure culture"; one which contains representatives of two or more species of bacteria is spoken of as an "impure," or "mixed" "cultivation," and it now becomes necessary to indicate the chief methods by which one or more organisms may be isolated in a state of purity from a mixture; whether that mixture exists as an impure laboratory cultivation, or is contained in pus and other morbid exudations, infected tissues, or water or food-stuffs.

Fig. 138.—Hæmatocytometer cell, showing, a, section through the centre of the cell, and b, a magnified image of the cell rulings.

Before the introduction of solid media the only method of obtaining pure cultivations was by "dilution"—by no means a reliable method. "Dilution" consisted in estimating approximately the number of bacteria present in a given volume of fluid (by means of a graduated-celled slide resembling a hæmatocytometer,[Pg 249] Fig. 138), and diluting the fluid by the addition of sterile water or bouillon until a given volume (usually 1 c.c.) of the dilution contained but one organism. By planting this volume of the fluid into several tubes or flasks of nutrient media, it occasionally happened that the resulting growth was the product of one individual microbe. A method so uncertain is now fortunately replaced by many others, more reliable and convenient, and in those methods selected for description here, the segregation and isolation of the required bacteria may be effected—

A. By Mechanical Separation.

1. By surface plate cultivation:

[2. By Esmarch's roll cultivation:

This archaic method (see page 226) is no longer employed for the isolation of bacteria.]

3. By serial cultivation.

B. By Biological Differentiation.

4. By differential media.

5. By differential incubation.

6. By differential sterilisation.

7. By differential atmosphere cultivation.

8. By animal inoculation.

The selection of the method to be employed in any specific instance will depend upon a variety of circumstances, and often a combination of two or more will ensure a quicker and more reliable result than a rigid adherence to any one method. Experience is the only reliable guide, but as a general rule the use of[Pg 250] either the first or the third method will be found most convenient, affording as each of them does an opportunity for the simultaneous isolation of several or all of the varieties of bacteria present in a mixture.

1. Surface Plate Cultivations.—

(a) Gelatine (vide page 164).

(b) Agar (vide page 167).

(c) Alkaline serum agar (vide page 211).

These plates are prepared in a manner precisely similar to that adopted for nutrient gelatine and agar surface plates (vide pages 231-233).

(d) Serum Agar.—

1. Melt three tubes of nutrient agar, label them 1, 2, and 3, and place them, with three tubes of sterile fluid serum, also labelled 1a, 2a, and 3a, in a water-bath regulated at 45° C.; allow sufficient time to elapse for the temperature of the contents of each tube to reach that of the water-bath.

2. Take serum tube No. 1a and agar tube No. 1. Flame the plugs and remove them from the tubes (retaining the plug of the agar tube in the hand); flame the mouths of the tubes, pour the serum into the tube of liquefied agar and replace the plug of the agar tube.

3. Mix thoroughly and pour plate No. 1 secundum artem.

4. Treat the remaining tube of agar and serum in a similar fashion, and pour plates Nos. 2 and 3.

5. Dry the serum agar plates in the incubator running at 60° C. for one hour (see page 232).

6. Inoculate the plates in series as described for gelatine surface plates (page 231).

(e) Blood Agar, Human.—

1. Melt a tube of sterile agar and pour it into a sterile plate; let it set.[Pg 251]

2. Collect a few drops of human blood, under all aseptic conditions, in a sterile capillary teat pipette.

3. Raise the cover of the Petri dish very slightly, insert the extremity of the capillary pipette, and deposit the blood on the centre of the agar surface. Close the dish.

4. Charge a platinum loop with a small quantity of the inoculum. Raise the cover of the plate, introduce the loop, mix its contents with the drop of blood, remove the loop, close the dish and sterilise the loop.

5. Finally smear the mixture over the surface of the agar with a sterilised L-shaped rod.

6. Label and incubate.

(If considered necessary, two, three, or more similar plates may be inoculated in series.)

(f) Blood Agar, Animal.—

When preparing citrated blood agar (page 171) it is always advisable to pour several blood agar tubes into plates, which can be stored in the ice chest ready for use at any moment for surface plate cultures.

(g) Hanging-drop or block culture, (vide page 233).

3. Serial Cultivations.—These are usually made upon agar or blood-serum, although gelatine may also be used.

The method is as follows:

1. Take at least four "slanted" tubes of media and number them consecutively.

2. Flame all the plugs and see that each can be readily removed.

3. Charge the platinum loop with a small quantity of the inoculum, observing the usual routine, and plant tube No. 1, smearing thoroughly all over the surface. If any water of condensation has collected at the bottom of the tube, use this as a diluent before smearing the contents of the loop over the surface of the medium.

4. Without sterilising or recharging the loop, inoculate[Pg 252] tube No. 2, by making three parallel streaks from end to end of the slanted surface.

5. Plant the remainder of the tubes in the series as "smears" like tube No. 1.

6. Label with distinctive name or number, and date; incubate.

The growth that ensues in the first two or three tubes of the series will probably be so crowded as to be useless. Toward the end of the series, however, discrete colonies will be found, each of which can be transferred to a fresh tube of nutrient medium without risk of contamination from the neighbouring colonies.

"Working" up Plates.—

Having succeeded in obtaining a plate (or tube cultivation) in which the colonies are well grown and sufficiently separated from each other, the process of "working up," "pricking out," or "fishing" the colonies in order to obtain subcultures in a state of purity from each of the different bacteria present must now be proceeded with.

Occasionally it happens that this is quite a simple matter. For example, the original mixed cultivation when examined microscopically was found to contain a Gram positive micrococcus, a Gram positive straight bacillus and a Gram negative short bacillus. The third gelatine plate prepared from this mixture, on inspection after four day's incubation, showed twenty-five colonies—seven moist yellow colonies, each sinking into a shallow pit of liquefied gelatine, fourteen flat irridescent filmy colonies, and four raised white slimy colonies. A film preparation (stained Gram) from each variety examined microscopically showed that the yellow liquefying colony was composed of Gram positive micrococci; the flat colony of Gram positive bacilli and the white colony of gram negative bacilli. One of each of these varieties of colonies would be transferred by means of[Pg 253] the sterilised loop to a fresh gelatine culture tube, and after incubation the growth in each subculture would correspond culturally and microscopically with that of the plate colony from which it was derived,—the object aimed at would therefore be achieved.

Usually, however, the colonies cannot be thus readily differentiated, and unless they are "worked up" in an orderly and systematic manner much labour will be vainly expended and valuable time wasted. The following method minimises the difficulties involved.

(A) Inspection.

a. Without opening the plate carefully study the various colonies with the naked eye, with the assistance of a watchmaker's lens or by inverting the plate on the stage of the microscope and viewing with the 1-inch objective through the bottom of the plate and the layer of medium.

b. If gross differences can be detected mark a small circle on the bottom of the plate around the site of each of the selected colonies, with the grease pencil.

c. If no obvious differences can be made out choose nine colonies haphazard and indicate their positions by pencil marks on the bottom of the plate.

(B) Fishing Colonies.—

a. Take a sterile Petri dish and invert it upon the laboratory bench. Rule two parallel lines on the bottom of the dish with a grease pencil, and two more parallel lines at right angles to the first pair—so dividing the area of the dish into nine portions. Number the top right-hand portion 1, and the central bottom portion 8 (Fig. 139). Revert the dish. The numbers 1 and 8 can be readily recognised through the glass and by their positions enable any of the other divisions to be localised by number. This is the stock dish.

b. Slightly raise the cover of the dish, and with a[Pg 254] sterile teat-pipette deposit a small drop of sterile water in the centre of each of the nine divisions.

c. With the sterilised platinum spatula raise one of the marked colonies from the "plate 3" and transfer it to the first division in the ruled plate and emulsify it in the drop of water awaiting it. Repeat this process with the remaining colonies, emulsifying a separate colony in each drop of water.

(C) Preliminary Differentiation of Bacteria.—

a. Prepare a cover-slip film preparation from each drop of emulsion in the "stock dish" and number to correspond to the division from which it was taken. Stain by Gram's method.

b. Examine microscopically, using the oil immersion lens and note the numbers of those cover-slips which morphologically and by Gram results appear to be composed of different species of bacteria.

Fig. 139.—Diagram for stock plate.

(D) Preparing Isolation Subcultures.—

a. Inoculate an agar slope and a broth tube from the emulsion in the stock dish corresponding to each of these specially selected numbers.

b. Ascertain whether the cover-slips from the nine emulsions in the stock dish include all the varieties represented in the cover-slip film preparation made from the original mixture before plating.

c. If some varieties are missing prepare a second stock dish from other colonies on plate 3, and repeat the process until each morphological form or tinctorial variety has been secured in subculture.

d. Place the stock dishes in the ice chest to await the[Pg 255] results of incubation. (If any of the subcultures fail, further material can be obtained from the corresponding emulsion; or if it has dried, by moistening it with a further drop of sterile distilled water.)

e. Incubate all the subcultures and identify the organisms picked out.

4. Differential Media.—

(a) Selective.—Some varieties of media are specially suitable for certain species of bacteria and enable them to overgrow and finally choke out other varieties; e. g., wort is the most suitable medium-base for the growth of torulæ and yeasts and should be employed when pouring plates for the isolation of these organisms. To obtain a pure cultivation of yeast from a mixture containing bacteria as well, it is often sufficient to inoculate wort from the mixture and incubate at 37° C. for twenty-four hours. Plant a fresh tube of wort from the resulting growth and incubate. Repeat the process once more, and from the growth in this third tube plant a streak on wort gelatine, and incubate at 20°C. The resulting growth will almost certainly be a pure culture of the yeast.

(b) Deterrent.—The converse of the above also obtains. Certain media possess the power of inhibiting the growth of a greater or less number of species. For instance, media containing carbolic acid to the amount of 1 per cent. will inhibit the growth of practically everything but the Bacillus coli communis.

5. Differential Incubation.—

In isolating certain bacteria, advantage is taken of the fact that different species vary in their optimum temperature. A mixture containing the Bacillus typhosus and the Bacillus aquatilis sulcatus, for example, may be planted on two slanted agar tubes, the one incubated at 40°C., and the other at 12° C. After twenty-four hours' incubation the first will show a pure cultivation of the[Pg 256] Bacillus typhosus, whilst the second will be an almost pure culture of the Bacillus aquatilis.

6. Differential Sterilisation.—

(a) Non-sporing Bacteria.—Similarly, advantage may be taken of the varying thermal death-points of bacteria. From a mixture of two organisms whose thermal death-points differ by, say, 4°C.—e. g., Bacillus pyocyaneus, thermal death-point 55°C., and Bacillus mesentericus vulgatus, thermal death-point 60°C.—a pure cultivation of the latter may be obtained by heating the mixture in a water-bath to 58° C. and keeping it at that point for ten minutes. The mixture is then planted on to fresh media and incubated, when the resulting growth will be found to consist entirely of the B. mesentericus.

(b) Sporing Bacteria.—This method finds its chief practical application in the differentiation of a spore-bearing organism from one which does not form spores. In this case the mixture is heated in a water-bath at 80° C. for fifteen to twenty minutes. At the end of this time the non-sporing bacteria are dead, and cultivations made from the mixture will yield a growth resulting from the germination of the spores only.

Differential sterilisation at 80° C. is most conveniently carried out in a water-bath of special construction, designed by Balfour Stewart (Fig. 140). It consists of a double-walled copper vessel mounted on legs, and provided with a tubulure communicating with the space between the walls. This space is nearly filled with benzole (boiling-point 80°C.; pure benzole, free from thiophene must be employed for the purpose, otherwise the boiling-point gradually and perceptibly rises in the course of time), and to the tubulure is fitted a long glass tube, some 2 metres long and about 0.75 cm. diameter, serving as a condensing tube (a tube half this length if provided with a condensing bulb at[Pg 257] the centre will be equally efficient). The interior of the vessel is partly filled with water and covered with a lid which is perforated for a thermometer. This latter dips into the water and records its temperature. A very small Bunsen flame under the apparatus suffices to keep the benzole boiling and the water within at a constant temperature of 80° C. The bath is thus always ready for use.

Method.—To use the apparatus.

1. Place some of the mixture itself, if fluid, containing the spores, or an emulsion of the same if derived from solid material, in a test-tube.

2. Immerse the test-tube in the water contained in the benzole bath, taking care that the upper level of the liquid in the tube is at least 2 cm. beneath the surface of the water in the copper vessel.

3. The temperature of the water, of course, falls a few degrees after opening the bath and introducing a tube of colder liquid, but after a few minutes the temperature will have again reached 80°C.

4. When the thermometer again records 80°C., note the time, and fifteen minutes later remove the tube containing the mixture from the bath.

5. Make cultures upon suitable media; incubate.

Fig. 140.—Benzole bath.

7. Differential Atmosphere Cultivation.—

(a) By adapting the atmospheric conditions to the particular organism it is desired to isolate, it is comparatively easy to separate a strict aerobe from a strict anaerobe, and vice versa. In the first case, however, it is important that the cultivations should[Pg 258] be made upon solid media, for if carried out in fluid media the aerobes multiplying in the upper layers of fluid render the depths completely anaerobic, and under these conditions the growth of the anaerobes will continue unchecked.

(b) When it is desired to separate a facultative anaerobe from a strict anaerobe, it is generally sufficient to plant the mixture upon the sloped surface agar, incubate aerobically at 37°C., and examine carefully at frequent intervals. At the first sign of growth, subcultivations must be prepared and treated in a similar manner. As a result of these rapid subcultures, the facultative anaerobe will be secured in pure culture at about the third or fourth generation.

(c) If, on the other hand, the strict anaerobe is the organism required from a mixture of facultative and strict anaerobes, pour plates of glucose formate agar (or gelatine) in the usual manner, place them in a Bulloch's or Novy's jar, and incubate at a suitable temperature. Pick off the colonies of the required organism when the growth appears, and transfer to tubes of the various media.

Incubate under suitable conditions as to temperature and atmosphere.

8. Animal Inoculation.—

Finally, when dealing with pathogenic organisms, it is often advisable to inoculate some of the impure culture (or even some of the original materies morbi) into an animal specially chosen on account of its susceptibility to the particular pathogenic organism it is desired to inoculate. Indeed, with some of the more sensitive and strictly parasitic bacteria this method of animal inoculation is practically the only method that will yield a satisfactory result.

[Pg 259]

XVI. METHODS OF IDENTIFICATION AND STUDY.

In order to identify an organism after isolation, tube, plate, and other cultivations must be prepared, incubated under suitable conditions as to temperature and environment, and examined from time to time (a) macroscopically, (b) by microscopical methods, (c) by chemical methods, (d) by physical methods, (e) by inoculation methods, and the results of these examinations duly recorded.

It must be stated definitely that no micro-organism can be identified by any one character or property, whether microscopical, biological or chemical, but that on the contrary its entire life history must be carefully studied and then its identity established from a consideration of the sum total of these observations.

In order to give to the recorded results their maximum value it is essential that they should be exact and systematic, therefore some such scheme as the following should be adhered to; and especially is this necessary in describing an organism not previously isolated and studied.

SCHEME OF STUDY.

Designation:

Originally isolated by (observer's name) in (date), from (source of organism).

1. Cultural Characters.—(Vide Macroscopical Examination of Cultivation, page 261.)

Special media for the purpose of demonstrating characteristic appearances.

2. Morphology.—(Vide Microscopical Examination of Cultivations, page 272.)

3. Chemical Products of Growth.—(Vide Chemical Examination of Cultivations, page 276.)

4. Biology.—(Vide Physical Examination of Cultures, page 295.)

5. Pathogenicity:

MACROSCOPICAL EXAMINATION OF CULTIVATIONS.

In describing the naked-eye and low-power appearances of the bacterial growth the descriptive terms introduced by Chester (and included in the following scheme) should be employed.

Solid Media.

Plate Cultures.—

Gelatine.—Note the presence or absence of liquefaction of the surrounding medium. If liquefaction is present, note shape and character (vide page 269, "stab" cultures).

Agar.—No liquefaction takes place in this medium. The liquid found on the surface of the agar (or at the bottom of the tube in agar tube cultures) is merely[Pg 262] water which has been expressed during the rapid solidification of the medium and has subsequently condensed.

Gelatine and Agar.—Examine the colonies at intervals of twenty-four hours.

(a) With the naked eye.

(b) With a hand lens or watchmaker's glass.

(c) Under a low power (1 inch) of the microscope, or by means of a small dissecting microscope.

Distinguish superficial from deep colonies and note the characters of the individual colonies.

(A) Size.—The diameter in millimetres, at the various ages.

(B) Shape.—

Punctiform: Dimensions too slight for defining form by naked eye; minute, raised, hemispherical.

Round: Of a more or less circular outline.

Elliptical: Of a more or less oval outline.

Irregular: Outlines not conforming to any recognised shape.

Fusiform: Spindle-shaped, tapering at each end.

Cochleate: Spiral or twisted like a snail shell (Fig. 141, a).

Fig. 141.—Types of colonies: a, Cochleate; b, amœboid; c, mycelioid.

[Pg 263]

Amœboid: Very irregular, streaming (Fig. 141, b).

Mycelioid: A filamentous colony, with the radiate character of a mould (Fig. 141, c).

Filamentous: An irregular mass of loosely woven filaments (Fig. 142, a).

Floccose: Of a dense woolly structure.

Rhizoid: Of an irregular, branched, root-like character (Fig. 142, b).

Conglomerate: An aggregate of colonies of similar size and form (Fig. 142, c).

Toruloid: An aggregate of colonies, like the budding of the yeast plant (Fig. 142, d).

Rosulate: Shaped like a rosette.

Fig. 142.—Types of colonies: a, Filamentous; b, rhizoid; c, conglomerate; d, toruloid.

(C) Surface Elevation.—

1. General Character of Surface as a Whole:

Flat: Thin, leafy, spreading over the surface (Fig. 143, a).

Effused: Spread over the surface as a thin, veily layer, more delicate than the preceding.

Raised: Growth thick, with abrupt terraced edges (Fig. 143, b).

Convex: Surface the segment of a circle, but very flatly convex (Fig. 143, c).[Pg 264]

Pulvinate: Surface the segment of a circle, but decidedly convex (Fig. 143, d).

Capitate: Surface hemispherical (Fig. 143, e).

Umbilicate: Having a central pit or depression (Fig. 143, f).

Conical: Cone with rounded apex (Fig. 143, g).

Umbonate: Having a central convex nipple-like elevation (Fig. 143, h).

2. Detailed Characters of Surface:

Smooth: Surface even, without any of the following distinctive characters.

Alveolate: Marked by depressions separated by thin walls so as to resemble a honeycomb (Fig. 144).

Punctate: Dotted with punctures like pin-pricks.

Bullate: Like a blistered surface, rising in convex prominences, rather coarse.

Vesicular: More or less covered with minute vesicles due to gas formation; more minute than bullate.

Fig. 143.—Surface elevation of colonies: a, Flat; b, raised; c, convex; d, pulvinate; e, capitate; f, umbilicate; g, conical; h, umbonate.

Fig. 144.—Types of colonies—alveolate.

Verrucose: Wart-like, bearing wart-like prominences.

Squamose: Scaly, covered with scales.

Echinate: Beset with pointed prominences.

Papillate: Beset with nipple or mamma-like processes.[Pg 265]

Rugose: Short irregular folds, due to shrinkage of surface growth.

Corrugated: In long folds, due to shrinkage.

Contoured: An irregular but smoothly undulating surface, resembling the surface of a relief map.

Rimose: Abounding in chinks, clefts, or cracks.

(D) Internal Structure of Colony (Microscopical).—

Refraction Weak: Outline and surface of relief not strongly defined.

Refraction Strong: Outline and surface of relief strongly defined; dense, not filamentous colonies.

Fig. 145.—Types of colonies: a, Grumose; b, moruloid; c, clouded.

1. General:

Amorphous: Without any definite structure, such as is specified below.

Hyaline: Clear and colourless.

Homogeneous: Structure uniform throughout all parts of the colony.

Homochromous: Colour uniform throughout.

2. Granulations or Blotchings:

Finely granular.

Coarsely granular.

Grumose: Coarser than the preceding, with a clotted[Pg 266] appearance, and particles in clustered grains (Fig. 145, a).

Moruloid: Having the character of a mulberry, segmented, by which the colony is divided in more or less regular segments (Fig. 145, b).

Clouded: Having a pale ground, with ill-defined patches of a deeper tint (Fig. 145, c).

Fig. 146.—Types of colonies: a, Reticulate; b, gyrose; c, marmorated.

3. Colony Marking or Striping:

Reticulate: In the form of a network, like the veins of a leaf (Fig. 146, a).

Areolate: Divided into rather irregular, or angular, spaces by more or less definite boundaries.

Gyrose: Marked by wavy lines, indefinitely placed (Fig. 146, b).

Marmorated: Showing faint, irregular stripes, or traversed by vein-like markings, as in marble (Fig. 146, c).

Rivulose: Marked by lines like the rivers of a map.

Rimose: Showing chinks, cracks, or clefts.

Fig. 147.—Types of colonies—curled.

4. Filamentous Colonies:

Filamentous: As already defined.

Floccose: Composed of filaments, densely placed.[Pg 267]

Curled: Filaments in parallel strands, like locks or ringlets (Fig. 147).

(E) Edges of Colonies.—

Entire: Without toothing or division (Fig. 148, a).

Undulate: Wavy (Fig. 148, b).

Repand: Like the border of an open umbrella (Fig. 148, c).

Erose: As if gnawed, irregularly toothed (Fig. 148, d).

Fig. 148.—Edges of colonies: a, Entire; b, undulate; c, repand; d, erose.

Lobate.

Lobulate: Minutely lobate (Fig. 149, e).

Auriculate: With ear-like lobes (Fig. 149, f).

Lacerate: Irregularly cleft, as if torn (Fig. 149, g).

Fimbriate: Fringed (Fig. 149, h).

Ciliate: Hair-like extensions, radiately placed (Fig. 149, j).

Tufted.

Filamentous: As already defined.

Curled: As already defined.

Fig. 149.—Edges of colonies: e, Lobar-lobulate; f, auriculate; g, lacerate; h, fimbriate; i, ciliate.

(F) Optical Characters (after Shuttleworth).—

1. General Characters:

Transparent: Transmitting light.[Pg 268]

Vitreous: Transparent and colourless.

Oleaginous: Transparent and yellow; olive to linseed-oil coloured.

Resinous: Transparent and brown, varnish or resin-coloured.

Translucent: Faintly transparent.

Porcelaneous: Translucent and white.

Opalescent: Translucent; greyish-white by reflected light.

Nacreous: Translucent, greyish-white, with pearly lustre.

Sebaceous: Translucent, yellowish or greyish-white.

Butyrous: Translucent and yellow.

Ceraceous: Translucent and wax-coloured.

Opaque.

Cretaceous: Opaque and white, chalky.

Dull: Without lustre.

Glistening: Shining.

Fluorescent.

Iridescent.

2. Chromogenicity:

Colour of pigment.

Pigment restricted to colonies.

Pigment restricted to medium surrounding colonies.

Pigment present in colonies and in medium.

Streak or Smear Cultures.—

Gelatine and Agar.—Note general points as indicated under plate cultivations.

Inspissated Blood-serum.—Note the presence or absence of liquefaction of the medium. (The presence of condensation water at the bottom of the tube must not be confounded with liquefaction of the medium.)

All Oblique Tube Cultures.—

1. Colonies Discrete: Size, shape, etc., as for plate cultivations (vide page 261).[Pg 269]

2. Colonies Confluent: Surface elevation and character of edge, as for plate cultivations (vide page 263).

Chromogenicity: As for plate cultures.

Gelatine Stab Cultures.—

(A) Surface Growth.—As for individual colonies in plate cultures (vide page 261).

Fig. 150.—Stab cultivations—types of growth: a, Filiform; b, beaded; c, echinate; d, villous; e, arborescent.

(B) Line of Puncture.—

Filiform: Uniform growth, without special characters (Fig. 150, a).

Nodose: Consisting of closely aggregated colonies.[Pg 270]

Beaded: Consisting of loosely placed or disjointed colonies (Fig. 150, b).

Papillate: Beset with papillate extensions.

Echinate: Beset with acicular extensions (Fig. 150, c).

Villous: Beset with short, undivided, hair-like extensions (Fig. 150, d).

Plumose: A delicate feathery growth.

Fig. 151.—Stab cultivations—types of growth: f, Crateriform; g, saccate; h, infundibuliform; j, napiform; k, fusiform; l, stratiform.

Arborescent: Branched or tree-like, beset with branched hair-like extensions (Fig. 150, e).

(C) Area of Liquefaction (if present).—

Crateriform: A saucer-shaped liquefaction of the gelatine (Fig. 151, f).[Pg 271]

Saccate: Shape of an elongated sack, tubular cylindrical (Fig. 151, g).

Infundibuliform: Shape of a funnel, conical (Fig. 151, h).

Napiform: Shape of a turnip (Fig. 151, j).

Fusiform: Outline of a parsnip, narrow at either end, broadest below the surface (Fig. 151, k).

Stratiform: Liquefaction extending to the walls of the tube and downward horizontally (Fig. 151, l).

(D) Character of the Liquefied Gelatine.—

1. Pellicle on surface.

2. Uniformly turbid.

3. Granular.

4. Mainly clear, but containing flocculi.

5. Deposit at apex of liquefied portion.

(E) Production of Gas Bubbles.

Shake Cultures.—

1. Presence or absence of liquefaction.

2. Production of gas bubbles.

3. Bulk of growth at the surface—aerobic.

4. Bulk of growth in depths—anaerobic.

Fluid Media.

1. Surface of the Liquid.—

Presence or absence of froth due to gas bubbles.

Presence or absence of pellicle formation.

Character of pellicle.

2. Body of the Liquid.—

Uniformly turbid.

Flocculi in suspension.

Granules in suspension.

Clear, with precipitate at bottom of tube.

Colouration of fluid, presence or absence of.

3. Precipitate.—

Character.[Pg 272]

Amount.

Colour.

Carbohydrate Media.—

Growth.

Reaction.

Gas formation.

Coagulation or not of serum albumen (when serum water media are employed).

Litmus Milk Cultivations.—

(a) Coagulum undissolved.

(b) Coagulum finally peptonised, completely: incompletely.

Resulting solution, clear: turbid.

BY MICROSCOPICAL METHODS.

As a council of perfection preparations must be made from pure cultivations 4, 6, 8, 12, 18, and 24[Pg 273] hours; and subsequently at intervals of, say, twenty-four hours, during the entire period they are under observation, and examined—

(A) Living.—1. In hanging drop, to determine motility or non-motility.

In this connection it must be remembered that under certain conditions as to environment (e. g., when examined in an unsuitable medium, atmosphere, temperature, etc.) motile bacilli may fail to exhibit activity. No organism, therefore, should be recorded as non-motile from one observation only; a series of observations at different ages and under varying conditions should form the basis of an opinion as to the absence of true locomotion.

Size.—In the case of non-motile or sluggishly motile organisms, endeavour to measure several individuals in each hanging drop by means of the eyepiece micrometer or the eikonometer (vide page 63), and average the results.

If the organism is one which forms spores, observe—

(a) Spore Formation.—Prepare hanging-drop cultivations (vide page 78) from vegetative forms of the organism, adding a trace of magenta solution (0.5 per cent.) or other intra vitam stain (see page 77) to the drop, on the point of the platinum needle, to facilitate the observation of the phenomenon by rendering the bacilli more distinct.

Place the preparation on the stage of the microscope; if necessary, using a warm stage.

Arrange illumination, etc., and select a solitary bacillus for observation, by the help of the 1/6-inch lens.

Substitute the 1/12-inch oil-immersion lens for the sixth, and observe the formation of the spore; if possible, measure any alteration in size which may occur by means of the Ramsden micrometer.

(b) Spore Germination.—Prepare hanging-drop cultivations from old cultivations in which no living vegetative[Pg 274] forms are present, and observe the process of germination in a similar manner.

The comfort of the microscopist is largely enhanced in those cases where the period of observation is at all lengthy, by use of some form of eye screen before the unemployed eye, such as is figured on page 58 (Fig. 49).

If it is impossible to carry out the method suggested above, proceed as follows:

(a) Spore Formation.—Plant the organism in broth and incubate under optimum conditions.

At regular intervals, say every thirty minutes, remove a loopful of the cultivation and prepare a cover-slip film preparation.

Fix, while still wet, in the corrosive sublimate fixing solution.

Stain with aniline gentian violet, and partially decolourise with 2 per cent. acetic acid.

Mount and number consecutively; then examine.

(b) Spore Germination.—Expose a thick emulsion of the spores to a temperature of 80° C. for ten minutes in the differential steriliser (vide page 257).

Transfer the emulsion to a tube of sterile nutrient broth and incubate.

Remove specimens from the tube culture at intervals of, say, five minutes.

Fix, stain, etc., wet, as under (a), and examine.

(B) Fixed.—2. In stained preparations.

(a) To determine points in morphology:

Shape (vide classification, page 131).

Size:

(a) Prepare cover-slip film preparations at the various ages, and fix by exposure to a temperature of 115° C. for twenty minutes in hot-air oven.

(b) Stain the preparations by Gram's method (if applicable) or with dilute carbol-fuchsin, and mount in the usual way.[Pg 275]

(c) Measure (vide page 66) some twenty-five individuals in each film by means of the Ramsden's or the stage micrometer and average the result.

Pleomorphism; If noted, record—

(b) To demonstrate details of structure:

Flagella: If noted, record—