The Elements of Bacteriological Technique - A Laboratory Guide for Medical, Dental, and Technical Students. Second Edition Rewritten and Enlarged.

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Title: The Elements of Bacteriological Technique - A Laboratory Guide for Medical, Dental, and Technical Students. Second Edition Rewritten and Enlarged.
Author: Eyre, J. W. H. (John William Henry), 1869-
Language: English
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TECHNIQUE***

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THE ELEMENTS OF BACTERIOLOGICAL TECHNIQUE

A Laboratory Guide for Medical, Dental, and Technical Students

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

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.

JOHN W. H. EYRE.

GUY'S HOSPITAL, S. E.
_July, 1913._

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.

JOHN W. H. EYRE.

GUY'S HOSPITAL, S. E.

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.

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

[Illustration]

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._

~BACTERIOLOGICAL DEPARTMENT.~

HANDLING OF INFECTIVE MATERIALS.

The following Regulations have been drawn up in the interest
of those working in the Laboratory as well as the public at
large, and will be strictly enforced.

Their object is to avoid the dangers of infection which may
arise from neglect of necessary precautions or from
carelessness.

Everyone must note that by neglecting the general rules laid
down he not only runs grave risk himself, but is a danger to
others.

REGULATIONS.

1. Each worker must wear a gown or overall, provided at his
own expense, which must be kept in the Laboratory.

2. The hands must be disinfected with lysol 2 per cent.
solution, carbolic acid 5 per cent. solution, or corrosive
sublimate 1 per mille solution, after dealing with
infectious material, and ~before using towels~.

3. On no account must Laboratory towels or dusters be used
for wiping up infectious material, and if such towels or
dusters do become soiled, they must be immediately
sterilised by boiling.

4. Special pails containing disinfectant are provided to
receive any waste material, and nothing must be thrown on
the floor.

5. All instruments must be flamed, boiled, or otherwise
disinfected immediately after use.

6. Labels must be moistened with water, and not by the
mouth.

7. All disused cover-glasses, slides, and pipettes after use
in handling infectious material, etc., must be placed in 2
per cent. lysol solution. A vessel is supplied on each bench
for this purpose.

8. All plate and tube cultures of pathogenic organisms when
done with, must be placed for immediate disinfection in the
boxes provided for the purpose.

9. No fluids are to be discharged into sinks or drains
unless previously disinfected.

10. Animals are to be dissected only after being nailed out
on the wooden boards, and their skin thoroughly washed with
disinfectant solution.

11. Immediately after the post-mortem examination is
completed each cadaver must be placed in the zinc
animal-box--_without removing the carcase from the
post-mortem board_--and the cover of the box replaced, ready
for carriage to the destructor.

12. Dead animals, when done with, are cremated in the
destructor, and the laboratory attendant must be notified
when the bodies are ready for cremation.

13. None of the workers in the laboratory are allowed to
enter the animal houses unless accompanied by the special
attendant in charge, who must scrupulously observe the same
directions regarding personal disinfection as the workers in
the laboratories.

14. No cultures are to be taken out of the laboratory
without the permission of the head of the Department.

15. All accidents, such as spilling infected material,
cutting or pricking the fingers, must be at once reported to
the bacteriologist in charge.

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 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.

[Illustration: FIG. 1.--Bohemian flask.]

[Illustration: FIG. 2.--Pear-shaped flask.]

[Illustration: 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 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.

[Illustration: FIG. 4.--Kolle's culture flask.]

[Illustration: FIG. 5.--Roux's culture bottle.]

[Illustration: FIG. 6.--Guy's culture bottle.]

[Illustration: 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.

~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.

[Illustration: FIG. 8.--Petri dish (a), and capsules (b, c).]

[Illustration: FIG. 9.--Plate box with stirrup.]

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

1. Pipettes of 1 c.c. capacity graduated in 0.1 c.c.

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

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.

[Illustration: 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. 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.

[Illustration: 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 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.

[Illustration: 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 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.

[Illustration: 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 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).

[Illustration: 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 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.

[Illustration: FIG. 15.--Blood pipettes and hair-lip pin in a
test-tube.]

[Illustration: 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
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.

[Illustration: 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.

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.

[Illustration: 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 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).

[Illustration: 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).

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 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.

[Illustration: 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.

[Illustration: 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 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.

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.

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.

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).

[Illustration: 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, 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.

[Illustration: 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.

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

NOTE.--A few pieces of pipe-clay or pumice may be placed in
the beaker to prevent the "spurting" of the chromic acid.

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.

NOTE.--After once being placed in the chromic acid, the
cover-slips must on no account be touched by the fingers.

~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.

_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.

NOTE.--_Test-tubes, plates, capsules_, etc., which, from
long use, have become scratched and hazy, or which cannot be
cleaned in any other way, may be dealt with by immersing
them in an enamelled iron bath, containing water acidulated
to 1 per cent. with hydrofluoric acid, for ten minutes,
rinsing thoroughly in water, drying, and polishing.

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 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.

[Illustration: 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.

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).

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:

Lysol, 2 per cent. solution;
Perchloride of mercury, 0.1 per cent. solution;
Carbolic acid, 5 per cent. solution;
Absolute alcohol;
Ether;
Chloroform;
Camphor;
Thymol;
Toluol;
Volatile oils, such as oil of mustard, oil of garlic.

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.

NOTE.--Tubes, flasks, filter flasks, pipettes, glass tubing,
etc., may be rapidly sterilised, in case of emergency, by
washing, in turn, with distilled water, perchloride of
mercury solution, alcohol, and ether, draining, and finally
gently heating over a gas flame to completely drive off the
ether vapor. Chloroform or other volatile disinfectants may
be added to various fluids in order to effect the
destruction of contained bacteria, and when this has been
done, may be completely driven off from the fluid by the
application of gentle heat.

~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.

[Illustration: 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:

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.

NOTE.--The too rapid cooling of the candles, such as takes
place if they are removed from the muffle before it has
cooled down to the room temperature, may give rise to
microscopic cracks and flaws which will effectually destroy
their efficiency.

_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 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.

[Illustration: FIG. 26.--Hot-air oven.]

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.

To prepare a wire crate for the reception of test-tubes,
etc., cover the bottom with a layer of thick asbestos cloth;
or take some asbestos fibre, moisten it with a little water
and knead it into a paste; plaster the paste over the bottom
of the crate, working it into the meshes and smoothing the
surface by means of a pestle. When several crates have been
thus treated, place them inside the hot-air oven, close the
door, open the ventilating slide, light the gas, and run the
temperature of the interior up to about 160° C. After an
interval of ten minutes extinguish the gas, open the oven
door, and allow the contents to cool. The asbestos now forms
a smooth, dry, spongy layer over the bottom, which will last
many months before needing renewal, and will considerably
diminish the loss of tubes from breakage.

Copper cylinders and large test-tubes intended for the
reception of pipettes are prepared in a similar manner, in
order to protect the points of these articles from injury.

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.

NOTE.--Neglect of this precautionary cooling of the oven to
60° C. will result in numerous cracked and broken tubes.

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.

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).

[Illustration: 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 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.

[Illustration: FIG. 28.--Koch's steriliser.]

[Illustration: 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 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.

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.

NOTE.--After removing tubes, flasks, etc., from the steam
steriliser, they should be at once separated freely in order
to prevent moisture condensing upon the cotton-wool plugs
and soaking through into the interior of the tubes.

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., 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 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.

[Illustration: 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 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.

[Illustration: FIG. 31.--Electric signal timing clock.]

~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.

[Illustration: 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:

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.

[Illustration: 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.

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).

NOTE.--In selecting candles for use in the laboratory avoid
those with metal fittings, since during sterilisation cracks
develop at the junction of the metal and the siliceous
material owing to the unequal expansion.

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.

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.

[Illustration: FIG. 34.--Porcelain filter candle.]

[Illustration: FIG. 35.--Geryk air pump.]

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

[Illustration: 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).

[Illustration: 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 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 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.

[Illustration: 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.

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.

[Illustration: 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.

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.

IV. THE MICROSCOPE.

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

[Illustration: 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 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:

[Illustration: 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
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.

[Illustration: FIG. 42.--Coarse adjustment.]

[Illustration: FIG. 43.--Fine adjustment.]

For practical purposes the tube length = distance from the
end of the nosepiece to the eyeglass of the ocular. This is
the measurement referred to in speaking of "long" or "short"
tube.

~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 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.

[Illustration: 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 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.

[Illustration: FIG. 45.--Mechanical stage.]

[Illustration: 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" 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.

[Illustration: 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 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 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.

[Illustration: 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 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:

1. A thin (one cell layer), even } { 1", 2/3", 1/2":
"blood film," stained with Jenner's } for { 1/6", 1/8"
or Romanowsky's stain. } { 1/12" oil

2. A thin cover-slip preparation }
of a young cultivation of } { 1/8" dry
_B. diphtheriæ_ (showing } for {
segmentation) stained with } { 1/12" oil
methylene-blue. }

~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 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.

[Illustration: 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.

[Illustration: FIG. 50.--Double nosepiece.]

[Illustration: 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. 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.

[Illustration: 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 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.

[Illustration: 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 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.

[Illustration: 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.

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.

[Illustration: 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.

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.

[Illustration: FIG. 56.--Eyepiece micrometer, ordinary.]

[Illustration: 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 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:

x = 10 y.

Where x = the number of included divisions of the
eyepiece micrometer.

y = the number of included divisions of the
stage micrometer.

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.

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 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.

[Illustration: FIG. 58.--Ramsden's Filar micrometer.]

[Illustration: 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.

As in the previous method, the value of each division of the micrometer
scale (i. e., the comb) must first be determined for each optical
combination. This is effected as follows:

1. Place the filar micrometer and the stage micrometer in their
respective positions.

2. Rotate the screw of the filar micrometer until the movable wire
coincides with the fixed one, and the index marks zero on the drum head.
(If when the drum head is at zero the two wires do not exactly coincide
they must be adjusted by loosening the drum screw and resetting the
drum.)

3. Focus the scale of each micrometer accurately, and make the lines on
them parallel.

4. Rotate the head of the micrometer screw until the movable line has
transversed one division of the stage micrometer. Note the number of
complete revolutions (by means of the recording comb) and the fractions
of a revolution (by means of scale on the head of the micrometer screw),
which are required to measure the 0.01 mm.

5. Make several such estimations and average the results.

6. Note the optical combination employed in this experiment and record
it carefully, together with the micrometer value in terms of µ.

7. Repeat this process for each of the different optical combinations
and record the results.

To measure an object by this method, simply note the number of
revolutions and fractions of a revolution of the screw-head required to
traverse such object from edge to edge, and express the result as
_micra_ by reference to the recorded values for that particular optical
combination.

_Microscope Illuminant._--In tropical and subtropical regions diffuse
daylight is the best illuminant. In temperate climes however daylight of
the desirable quantity is not always available, and recourse must be
had to oil lamps, gas lamps--preferably those with incandescent
mantles--and electricity; and of these the last is undoubtedly the best.
A handy lamp holder which can be manufactured in the laboratory is shown
in Fig. 60. It consists of a base board weighted with lead to which is
attached the ordinary domestic lamp holder, and behind this is fastened
a curved sheet-iron reflector. An obscured metal filament lamp of about
16 candle power gives the most suitable light, and if monochromatic
light is needed, the blue grease pencil is streaked over the side of the
lamp nearest the microscope; the current is switched on and when the
glass bulb is warm, rubbing with a wad of cotton-wool will readily
distribute the blue greasy material in an even film over the ground
glass.

[Illustration: FIG. 60.--Electric microscope lamp.]

FOOTNOTES:

[1] Its importance will be realised, however, when it is stated in the
words of the late Professor Abbé: "The numerical aperture of a lens
determines all its essential qualities; the brightness of the image
increases with a given magnification and other things being equal, as
the square of the aperture; the resolving and defining powers are
directly related to it, the focal depth of differentiation of depths
varies inversely as the aperture, and so forth."

[2] Made by Mr. Otto Baumbach, 10, Lime Grove, Manchester.

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.)

[Illustration: 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.

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).

[Illustration: 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).

Another form of hanging-drop slide is made in which a
circular or oval concavity or "cell" is ground out of the
centre of a 3 by 1 slip. These are more expensive, less
convenient to work with, and are more easily contaminated by
drops of material under examination, and should be carefully
avoided.

7. Three aluminium rods (Fig. 63), each about 25 cm. long and carrying a
piece of 0.015 gauge platino-iridium wire 7.5 cm. in length. The end of
one of the wires is bent round to form an oval loop, of about 1 mm. in
its short diameter, and is termed a loop or an oese; the terminal 3 or 4
mm. of another wire is flattened out by hammering it on a smooth iron
surface to form a "spatula"; the third is left untouched or is pointed
by the aid of a file. These instruments are used for inoculating culture
tubes and preparing specimens for microscopical examination.

[Illustration: 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 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.

[Illustration: 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

Aniline water.

Gentian violet, saturated alcoholic solution.

Lugol's (Gram's) iodine.

Absolute alcohol.

Methylene-blue, }
Fuchsin, basic, } saturated alcoholic solution.

Neutral red, 1 per cent. aqueous solution.

Leishman's modified Romanowsky stain.

Carbolic acid, 5 per cent. aqueous solution.

Acetic acid, 1 per cent. solution.

Sulphuric acid, 25 per cent. solution.

Xylol.

[Illustration: FIG. 65.--Staining rack, rubber change mat and lysol
pot.]

[Illustration: FIG. 66.--Drop bottle.]

[Illustration: 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 with Canada balsam dissolved
in xylol, and sterile vaseline.

METHODS OF EXAMINATION.

Bacteria, etc., are examined microscopically.

1. In the living state, unstained, or stained.
2. In the "fixed" condition (i. e., fixed, killed,
and stained by suitable methods).

The preparation of a specimen from a tube cultivation for examination by
these methods may be described as follows:

~1. Living, Unstained.~--(a) _"Fresh" Preparation._--

1. Clean and dry a 3 by 1 glass slip and place it on one of the squares
of filter paper. Deposit a drop of water (preferably distilled) or a
drop of 1 per cent. solution of caustic potash, on the centre of the
slip, by means of the platinum loop.

[Illustration: FIG. 68.--Holding tubes for removing bacterial growth, as
seen from the front.]

TECHNIQUE OF OPENING AND CLOSING A CULTURE TUBE.

2. Remove the tube cultivation from its rack or jar with the
left hand and ignite the cotton-wool plug by holding it to
the flame of the Bunsen burner. Extinguish the flame by
blowing on the plug, whilst rotating the tube on its long
axis, its mouth directed vertically upward, between the
thumb and fingers. (This operation is termed "flaming the
plug," and is intended to destroy any micro-organisms that
may have become entangled in the loose fibres of the
cotton-wool, and which, if not thus destroyed, might fall
into the tube when the plug is removed and so accidentally
contaminate the cultivation.)

3. Hold the tube at or near its centre between the ends of
the thumb and first two fingers of the left hand, and allow
the sealed end to rest upon the back of the hand between the
thumb and forefinger, the plug pointing to the right. Keep
the tube as nearly in the horizontal position as is
consistent with safety, to diminish the risk of the
accidental entry of organisms (Fig. 68).

4. Take the handle of the loop between the thumb and
forefinger of the right hand, holding the instrument in a
position similar to that occupied by a pen or a paint-brush,
and sterilise the platinum portion by holding it in the
flame of a Bunsen burner until it is red hot. Sterilise the
adjacent portion of the aluminium handle by passing it
rapidly twice or thrice through the flame. After sterilising
it, the loop must not be allowed to leave the hand or to
touch against anything but the material it is intended to
examine, until it is finished with and has been again
sterilised.

5. Grasp the cotton-wool plug of the test-tube between the
little finger and the palm of the right hand (whilst still
holding the loop as directed in step 4), and remove it from
the mouth of the tube by a "screwing" motion of the right
hand.

6. Introduce the platinum loop into the tube and hold it in
this position until satisfied that it is quite cool. (The
cooling may be hastened by touching the loop on one of the
drops of moisture which are usually to be found condensed on
the interior of the glass tube, or by dipping it into the
condensation water at the bottom; at the same time care must
be taken in the case of cultures on solid media to avoid
touching either the medium or the growth.)

7. Remove a small portion of the growth by taking up a drop
of liquid, in the case of a fluid culture, in the loop; or
by touching the loop on the surface of the growth when the
culture is on solid medium; and withdraw the loop from the
tube without again touching the medium or the glass sides of
the tube.

8. Replace the cotton-wool plug in the mouth of the tube.

9. Replace the tube cultivation in its rack or jar.

10. Mix the contents of the loop thoroughly with the drop of water on
the 3 by 1 slide.

11. Again sterilise the loop as directed in step 4, and replace it in
its stand.

12. Remove a cover-slip from the glass capsule by means of the
cover-slip forceps, rest it for a moment on its edge, on a piece of
filter paper to remove the excess of alcohol, then pass it through the
flame of the Bunsen burner. This burns off the remainder of the alcohol,
and the cover-slip so "flamed" is now clean, dry, and sterile.

13. Lower the cover-slip, still held in the forceps, on to the surface
of the drop of fluid on the 3 by 1 slip, carefully and gently, to avoid
the inclusion of air bubbles.

14. Examine microscopically (_vide infra_).

During the microscopical examination, stains and other reagents may be
run in under a cover-slip by the simple method of placing a drop of the
reagent in contact with one edge of the cover-glass and applying the
torn edge of a piece of blotting paper to the opposite side. The reagent
may then be observed to flow across the field and come into contact with
such of the micro-organisms as lie in its path.

The non-toxic basic dyes most generally employed for the intra-vitam
staining of bacteria are

Neutral red, }
Quinoleine blue }
Methylene green } in 0.5 per cent. aqueous solutions.
Vesuvin, }

_Negative Stain_ (Burri).--By this method of demonstration the
appearances presented by dark ground illumination (by means of a
paraboloid condenser) are closely simulated, since minute particles,
bacteria, blood or pus cells etc. stand out as brilliantly white or
colourless bodies on a dark grey-brown background.

_Reagent required:_

Any one of the liquid waterproof black drawing inks (Chin-chin, Pelican,
etc.). This is prepared for use as follows:

Measure out and mix:

Liquid black ink, 25 c.c.
Tincture of iodine 1 c.c.

Allow the mixture to stand 24 hours, centrifugalise thoroughly, pipette
off the supernatant liquid to a clean bottle and then add a crystal of
thymol or one drop of formalin as a preservative.

METHOD.--

1. With the sterilised loop deposit one drop of the liquid ink close to
one end of a 3 by 1 slide.

2. With the sterilised loop deposit a drop of the fluid culture (or of
an emulsion from a solid culture) by the side of the drop of ink (Fig.
69, a); mix the two drops thoroughly by the aid of the loop.

3. Sterilise the loop.

4. Hold the slide firmly on the bench with the thumb and forefinger of
the left hand applied to the end nearest the drop of fluid.

5. Take another clean 3 by 1 slide in the right hand and lower its short
end obliquely (at an angle of about 60°) 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.

[Illustration: 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.)

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