the cultivation should be transferred with aseptic precautions, to a
sterile separatory funnel, preferably of the shape shown in figure 173,
and graduated if necessary. This is supported on a retort stand and
raised sufficiently above the level of the animal to be injected, so as
to secure a good "fall." A piece of sterilised rubber tubing of suitable
length, fitted with an injection needle and provided with a screw clamp,
is now attached to the nozzle of the funnel and the operation completed
according to the requirements of the particular case.
This method is quite satisfactory when the injection is made into the
pleural or abdominal cavities or directly into a vein but if the
injection has to be made into the subcutaneous tissue the "fall" may not
be sufficient to force the fluid in. In this case it will be necessary
to transfer the culture to a sterile wash-bottle and fasten a rubber
hand bellows to the air inlet tube (interposing an air filter) and
attach the tubing with the injection needle to the outlet tube (Fig.
174). By careful use sufficient force can be obtained to drive the
injection in.
(b) _Cultivations on Solid Media (e. g., Sloped Agar)._--
1. By means of a sterile graduated pipette introduce a suitable small
quantity of sterile bouillon (or sterile normal saline solution) into
the culture tube.
[Illustration: FIG. 174.--Arrangement of pressure injection apparatus.]
2. With a sterile platinum loop or spatula scrape the bacterial growth
off the surface of the medium, and emulsify it with the bouillon. It
then becomes to all intents and purposes a fluid inoculum.
3. Pour the emulsion into a sterile capsule and fill the syringe
therefrom.
(c) _Toxins._--Prepared by previously described methods (_vide_ page
318), are manipulated in a similar manner to cultivations in fluid
media.
(d) _Pathological Products._--Fluid secretions, excretions, etc., such
as serous exudation, pus, blood, etc., are treated as fluid
cultivations; but if the material is very thick or viscous, a small
quantity of sterile bouillon or normal saline solution may be used to
dilute it, and thorough incorporation effected by the help of a sterile
platinum rod.
Solid tissues, such as spleen, lymph glands, etc., may be divided into
small pieces by sterile instruments and rubbed up in a sterilised agate
mortar (using an agate pestle), with a small quantity of sterile
bouillon, and the syringe filled from the resulting emulsion.
[Illustration: FIG. 175.--Holding rabbit for shaving.]
If it is desired to inoculate tissue _en masse_, remove from the
material a small cube of 1 or 2 mm. and introduce it into a wound made
by sterile instruments in a suitable situation, and occlude the wound by
means of Michel's steel clips and a sealed dressing.
~Method of Securing Animals During Inoculation.~--
For the majority of inoculations, especially when no anaesthetic is
administered, it is customary to employ an assistant to hold the animal
(see Fig. 175).
If working single handed Voge's holder for guinea-pigs, is a useful
piece of apparatus the method of using which is readily seen from the
accompanying figures (Figs. 176, 177).
The instrument itself consists of a hollow copper cylinder, one end of
which is turned over a ring of stout copper wire, and from this open end
a slot is cut extending about half way along one side of the cylinder.
The opposite end is closed by a "pull-off" cap and is perforated around
its edge by a row of ventilating holes, which correspond with holes cut
in the rim of the cap. In the event of the animal resisting attempts to
remove it from the holder backwards, this cap is taken off and the
holder placed on the table and the guinea-pig allowed to walk out.
[Illustration: FIG. 176.--Taking guinea-pig's temperature.]
To provide for different-sized animals, two sizes of this holder will be
found useful:
1. Length, 16 cm.; breadth, 6 cm.; size of slot, 8 cm. by 2.5 cm.
2. Length, 20 cm.; breadth, 8 cm.; size of slot, 10 cm. by 2.5 cm.
A convenient holder for mice and even small rats is shown in figure 178,
the tail being securely held by the spring clip. Needless to say, the
holder should be entirely of metal, and the wire cage detachable and
easily renewed.
[Illustration: FIG. 177.--Voge's holder.]
When the animal is anaesthetised, it is more convenient to secure it
firmly to some simple form of operating table, such as Tatin's (Fig.
179), which will accommodate rabbits, guinea-pigs, and rats: or to the
more elaborate table devised by the author (Fig. 180).
[Illustration: FIG. 178.--Mouse holder.]
[Illustration: FIG. 179.--Tatin's operation table.]
~Operation Table.~--This is a table of the "aseptic" type, composed of
steel tubing, nickel-plated or enamelled. The table-top frame is
sufficiently large to accommodate rabbits, dogs and monkeys; and is
supported upon telescopic uprights, so that it is adjustable as to
height; in its long axis it can be inclined (at either end) to 45 deg.
from the horizontal. Further it can be completely rotated about its long
axis. The table-top itself is composed of a sheet of copper wire gauze
loosely suspended from the long sides of the tubular frame. The
slackness of the gauze bed permits of an india rubber hot water bottle,
or an electrotherm being placed under the animal, and if during the
course of an experiment it is necessary to reverse the animal, the
table-top frame is completely rotated, the device adopted for suspending
the gauze is detached and the gauze reversed also, so that it again
supports the animal from below.
[Illustration: FIG. 180.--Author's operating table[12].]
METHODS OF INOCULATION.
The following methods of inoculation apply more particularly to the
rabbit, but from them it will readily be seen what modifications in
technique, if any, are necessary in the case of the other experimental
animals.
~1. Cutaneous Inoculation.~--(_Anaesthetic, none._)
1. Have the animal firmly held by an assistant (or secured to the
operating table).
2. Apply the liquid soap to the fur, over the area selected for
inoculation, with a wad of cotton-wool, and lather freely by the aid of
warm water; shave carefully and thoroughly; or apply the depilatory
powder.
3. Wash the denuded area of skin thoroughly with 2 per cent. lysol
solution.
4. Wash off the lysol with ether and allow the latter to evaporate.
5. Make numerous short, parallel, superficial incisions with the point
of a sterile scalpel.
6. When the oozing from the incisions has ceased, rub the inoculum into
the scarifications by means of the flat of a scalpel blade, or a sterile
platinum spatula.
7. Cover the inoculated area with a pad of sterile gauze secured _in
situ_ by strips of adhesive plaster or by sealing down the edges of the
gauze with collodion.
8. Release the animal, place it in its cage, and affix a label upon
which is written:
(a) Distinctive name or number of the animal.
(b) Its weight.
(c) Particulars as to source and dose of inoculum.
(d) Date of inoculation.
~2. Subcutaneous Inoculation.~--
(a) _Fluid Inoculum._--(_Anaesthetic, none._)
Steps 1-4. As for cutaneous inoculation.
5. Pinch up a fold of skin between the forefinger and thumb of the left
hand; take the charged hypodermic syringe in the right hand, enter the
needle into a ridge of skin raised by the left finger and thumb, and
push it steadily onward until about 2 cm. of the needle are lying in the
subcutaneous tissue. Now release the grasp of the left hand and slowly
inject the fluid contained in the syringe.
6. Withdraw the needle, and at the same moment close the puncture with a
wad of cotton wool, to prevent the escape of any of the inoculum. The
injected fluid, unless large in amount, will be absorbed within a very
short time.
7. Label, etc.
(b) _Solid Inoculum.--(Anaesthetic, none; or Ethyl chloride spray.)_
Steps 1-4. As for cutaneous inoculation.
5. Raise a small fold of skin in a pair of forceps, and make a small
incision through the skin with a pair of sharp-pointed scissors or with
the point of a scalpel.
6. Insert a probe through the opening and push it steadily onward in the
subcutaneous tissue, and by lateral movements separate the skin from the
underlying muscles to form a funnel-shaped pocket with its apex toward
the point of entrance.
7. By means of a pair of fine-pointed forceps introduce a small piece of
the inoculum into this pocket and deposit it as far as possible from the
point of entrance.
[Illustration: FIG. 181.--Glass tube syringe for subcutaneous "solid"
inoculation.]
Or, improvise a syringe by sliding a piece of glass rod (to serve as a
piston) into the lumen of a slightly shorter length of glass tubing and
secure in position by a band of rubber tubing. Sterilise by boiling.
Withdraw the rod a few millimetres and deposit the piece of tissue
within the orifice of the tube, by means of sterile forceps. Now pass
the tube into the depths of the "pocket," push on the glass rod till it
projects beyond the end of the tube, and withdraw the apparatus, leaving
the tissue behind in the wound.
8. Close the wound in the skin with Michel's clips and a dressing of
gauze sealed with collodion (or Tinct. benzoin).
9. Label, etc.
~3. Intramuscular.~--
(a) _Fluid Inoculum.--(Anaesthetic, none.)_
Steps 1-4. As for cutaneous inoculation.
5. Steady the skin over the selected muscle or muscles with the slightly
separated left forefinger and thumb.
6. Thrust the needle of the injecting syringe boldly into the muscular
tissue and inject the inoculum slowly.
7. Label, etc.
(b) _Solid Inoculum.--(Anaesthetic, A. C. E.)_
1. Secure the animal to the operation table and anaesthetise.
2. Shave and disinfect the skin at the seat of operation.
3. Surround the field of operation by strips of gauze wrung out in 2 per
cent. lysol solution.
4. Incise skin, aponeurosis, and muscle in turn.
5. Deposit the inoculum in the depths of the incision.
6. Close the wound in the muscle with buried sutures and the cutaneous
wound with either continuous or interrupted sutures or with Michel's
steel clips.
7. Apply a sealed dressing of gauze and collodion.
8. Remove the animal from the operating table.
9. Label, etc.
~4. Intraperitoneal.~--
(a) _Fluid Inoculum.--(Anaesthetic, none.)_
Steps 1-4. As for cutaneous inoculation. Shave a fairly broad transverse
area, stretching from flank to flank.
5. Place the left forefinger on one flank and the thumb on the opposite,
and pinch up the entire thickness of the abdominal parietes in a
triangular fold. Now, by slipping the peritoneal surfaces (which are in
apposition) one over the other, ascertain that no coils of intestine are
included in the fold.
6. Take the syringe in the right hand and with the needle transfix the
fold near its base (Fig. 182).
7. Now release the fold, but hold the syringe steady; as the parietes
flatten out, the point of the needle is left free in the peritoneal
cavity (see Fig. 183).
[Illustration: FIG. 182.--Intraperitoneal inoculation--fluid.]
8. Inject the fluid from the syringe.
9. Label, etc.
[Illustration: FIG. 183.--Section of abdominal wall, etc., showing point
of needle lying free in the peritoneal cavity above the coils of
intestine.]
Second Method:
Steps 1-4. As in the first method.
5. Anaesthetise a small selected area of skin by spraying it with ethyl
chloride.
6. Heat platinum searing wire (0.5 mm. wire, twisted to the shape
indicated in figure 184, mounted in an aluminium handle) to redness, and
with it burn a hole through the anaesthetic area of skin and abdominal
muscle down to, but not through, the visceral peritoneum.
7. Fix a blunt-ended needle on to the charged syringe, and by pressing
the rounded end firmly against the peritoneum it can easily be pushed
through into the peritoneal cavity.
8. Inject the fluid from the syringe.
9. Label, etc.
This method is especially useful when it is desired to collect samples
of the peritoneal fluid from time to time during the period of
observation, as fluid can be removed from the peritoneal cavity, at
intervals, through this aperture in the abdominal parietes, by means of
a sterile capillary pipette.
[Illustration: FIG. 184.--Platinum wire for burning hole through
parietes.]
(b) _Solid Inoculum_ (or the implantation of capsules containing fluid
cultivations).--(_Anaesthetic, A. C. E._)
1. Anaesthetise the animal and secure it to the operating table.
2. Shave a large area of the abdominal parietes.
3. Make an incision through the skin in the middle line about 2 cm. in
length, midway between the lower end of the sternum and the pubes.
4. Divide the aponeuroses between the recti upon a director.
5. Divide the peritoneum upon a director.
6. Introduce the inoculum into the peritoneal cavity.
7. Close the peritoneal cavity with Lembert's sutures.
8. Close the skin and aponeurosis incisions together with interrupted
sutures or Michel's steel clips, and apply a sealed dressing.
9. Release the animal from the operating table.
10. Label, etc.
Suitable sacs may be readily prepared by either of the following
methods:
A. ~Collodion Sacs.~
1. Dip a small test-tube (5 by 0.5 cm.), bottom downward, into a beaker
of collodion, and dry in the air; repeat this process three or four
times.
2. Dip the tube, with its coating of collodion, alternately into a
beaker of alcohol and one of water. This loosens the collodion and
allows it to be peeled off in the shape of a small test-tube.
3. Take a 20 cm. length of glass tubing, of about the diameter of the
test-tube used in forming the sac, and insert one end into the open
mouth of the sac.
4. Suspend the glass tube with attached sac, inside a larger test-tube,
by packing cotton-wool in the mouth of the test-tube around the glass
tubing, and place in the incubator at 37 deg. C. for twenty-four hours.
When removed from the incubator, the sac will be firmly adherent to the
extremity of the glass tubing.
5. Plug the open end of the glass tubing with cotton-wool, and sterilise
the test-tube and its contents in the hot-air oven.
To use the sac, remove the plug from the glass tubing, partly fill the
sac with cultivation to be inoculated, by means of a sterile capillary
pipette, and replug the tubing. When the abdominal cavity has been
opened, remove the tubing and attached sac from the protecting
test-tube, close the sac by tying a sterilised silk thread tightly
around it a little below the end of the glass tubing, and separate it
from the tubing by cutting through the collodion above the ligature, and
the sac is ready for insertion in the peritoneal cavity.
B. ~Celloidin Sacs~ (_Harris_).
_Materials Required._
Quill glass tubing.
Gelatine capsules such as pharmacists prepare for the
exhibition of bulky powders.
Various grades of celloidin, thick and thin, in wide-mouthed
bottles.
1. Take a piece of quill glass tubing some 4 cm. long by 5 mm. diameter;
heat one end in the bunsen flame.
2. Thrust the heated end of the tube just through one end of a gelatine
capsule and allow it to cool (Fig. 185).
3. Remove any gelatine from the lumen of the tube with a heated platinum
needle; paint the joint between capsule and tube with moderately thick
celloidin and allow to dry.
[Illustration: FIG. 185.--Making celloidin capsules.]
4. Dip the capsule into a beaker containing thin celloidin, beyond the
junction with the glass and after removal rotate it in front of the
blowpipe air blast to dry it evenly. Repeat these manoeuvres until a
sufficiently thick coating is obtained.
5. Apply thick celloidin to the tube-capsule joint, the opposite end of
the capsule, and the line of junction of the capsule with its cap; dry
thoroughly.
6. With a teat pipette fill the capsule (through the attached tube) with
hot water, and stand the capsule in a beaker of boiling water for a few
minutes to melt the gelatine.
7. Remove the solution of gelatine from the interior of the celloidin
case with a pipette.
8. Fill the sac with nutrient broth and place it, _glass tube downward_,
in a tube containing sufficient sterile nutrient broth to cover the sac
to the depth of 1 cm. Plug the tube and sterilise in the steamer in the
usual manner.
9. To prepare the sac for use, empty it out of the broth tube into a
sterile glass dish.
10. Grasp the tube near its junction with the sac in the jaws of sterile
forceps, and with a teat pipette remove sufficient of the contained
broth to leave a small space in the sac. Introduce the inoculum in the
form of an emulsion by means of another pipette.
11. Still holding the tube in the forceps, draw it out and seal off near
the sac in the blowpipe flame.
12. When cool wash the sac in sterile water, then transfer to a tube of
nutrient broth and incubate over night to determine its impermeability
to bacteria.
13. If the broth outside the sac remains sterile, insert the sac in the
peritoneal cavity of the experimental animal.
~5. Intracranial.~--(_Anaesthetic, A. C. E._)
[Illustration: FIG. 186.--Guarded trephine.]
_Trephines and Surgical Engine._--The most useful instrument for
intracranial operations upon animals is the small nasal trephine
(Curtis) having a tooth cutting circle of 7 mm. The addition of an
adjustable collar guard--secured by a screw--prevents accidental
laceration of the dura mater or brain substance[13] (Fig. 186). This
size is suitable for monkeys, dogs, cats and large rabbits. Other
smaller sizes which will be found useful for guinea pigs and other small
animals cut circles of 6 and 4 mm.; for very small animals--young guinea
pigs and rats--a small dental drill or screw will make a sufficiently
large hole to admit the syringe needle. The trephine can be set in
ordinary metal handles and rotated by hand, but a surgical engine of
some kind is much preferable on the score of rapidity and safety to the
animal. The Guy's electrical Dental engine[14] (Fig. 187) which can be
connected to a lamp socket or wall plug, and is operated by a foot
switch, although inexpensive is eminently satisfactory.
NOTE.--A fine dental drill attached to the dental engine
renders the manufacture of aluminium handles needles (see
page 71) quite an easy matter.
(a) _Subdural._
1. Anaesthetise the animal and secure it to the operating table, dorsum
uppermost.
2. Shave a portion of the scalp immediately in front of the ears.
[Illustration: FIG. 187.--Guy's electrical dental engine.]
3. Mark out with a sharp scalpel a crescentic flap of skin muscle, etc.,
convexity forward, commencing 0.5 cm. in front of the root of one ear
and terminating at a similar spot in front of the other ear. Reflect the
marked flap.
4. Make a corresponding incision through the periosteum and raise it
with a blunt dissector.
5. With a small trephine (diameter 6 mm.) remove a circular piece of
bone from the parietal segment. The centre of the trephine hole should
be at the intersection of the median line and a line joining the
posterior canthi (Fig. 188).
6. Introduce the inoculum by means of a hypodermic syringe, perforating
the dura mater with the needle and depositing the material immediately
below this membrane, at the same time taking care to avoid injuring the
sinuses.
7. Turn back the flap of skin and secure it in position with Michel's
steel clips.
8. Dress with sterile gauze and wool and seal the dressing with
collodion.
9. Label, etc.
(b) _Intracerebral._--This inoculation is performed precisely as for
subdural save in step 6 the needle after perforating the dura mater is
pushed onward into the substance of one or other cerebral hemispheres
before the contents are ejected.
[Illustration: FIG. 188.--Intracranial inoculation of rabbit. The circle
indicates the situation of the trephine hole.]
~6. Intraocular.~--
(a) _Fluid Inoculum._--(_Anaesthetic, cocaine._)
1. Instil a few drops of a sterile solution of cocaine, and repeat the
instillation in two minutes.
2. Five minutes later have the animal firmly held by an assistant as in
intravenous injection (see Fig. 189), the head being steadied by the
assistant's hands.
3. Select two needles to accurately fit the same syringe and sterilise.
4. Attach one needle to the syringe and take up the required dose of
inoculum and remove the needle.
5. Steady the eye with fixation forceps; then pierce the cornea with the
other syringe needle and allow the aqueous to escape through the needle.
6. Without removing the needle from the cornea attach the syringe and
make the injection into the anterior chamber.
7. Irrigate the conjunctival sac with sterile saline solution.
8. Label, etc.
(b) _Solid Inoculum._--(_Anaesthetic, A. C. E._)
1. Anaesthetise the animal and secure it firmly to the operating table.
2. Irrigate the conjunctival sac thoroughly with sterile saline
solution.
3. Make an incision through the upper quadrant of the cornea into the
anterior chamber by means of a triangular keratome.
4. Separate the lips of the corneal wound with a flexible silver
spatula; seize the solid inoculum in a pair of iris forceps, introduce
it through the corneal wound, and deposit it on the anterior surface of
the iris; withdraw the forceps.
5. Again irrigate the sac and the surface of the cornea.
6. Release the animal from the operating table.
7. Label, etc.
~7. Intrapulmonary.~--
_Fluid Inoculum._--(_Anaesthetic, none._)
1. Have the animal firmly held by an assistant. (In this case the
foreleg of the selected side is drawn up by the assistant and held with
the ear of that side.)
2. Shave carefully in the axillary line and disinfect the denuded skin.
3. Thrust the needle of the syringe boldly through the fifth or sixth
intercostal space into the lung tissue.
4. Inject the contents of the syringe slowly.
5. Label, etc.
~8. Intravenous.~--
_Fluid Inoculum._--(_Anaesthetic, none._)
The site selected for the injection in the rabbit is the posterior
auricular vein (see Fig. 192). Although this is smaller than the median
vein, it is firmly bound down to the cartilage of the ear by dense
connective tissue, and is therefore more readily accessible. (In the
guinea-pig the jugular vein must be utilised, and in order to perform
the inoculation satisfactorily a general anaesthetic must be
administered to the animal. In the monkey or the dog, the internal
saphenous vein is the most convenient and before puncturing should be
distended or rendered prominent by compressing the vein above the
selected site.)
_Preparation of the Inoculum._--Care must be taken in preparing the
inoculum, as the injection of even small fragments may cause fatal
embolism. To obviate this risk the fluid should, if possible, be
filtered through sterile filter paper before filling into the syringe.
Air bubbles, when injected into a vein, frequently cause immediate
death. To prevent this, the syringe after being filled should be held in
the vertical position, needle uppermost. A piece of sterile filter paper
is then impaled on the needle and the piston of the syringe pressed
upward until all the air is expelled from the barrel and needle. Should
any drops of the inoculum be forced out, they will fall on the filter
paper, which should be immediately burned.
1. Have the animal firmly held by an assistant. The selected ear is
grasped at its root and stretched forward toward the operator.
2. Shave the posterior border of the dorsum of the ear.
3. Disinfect the skin over the vein, rubbing it vigourously with
cotton-wool soaked in lysol. The friction will make the vein more
conspicuous. Wash the lysol off with ether and allow the latter to
evaporate.
4. Direct the assistant to compress the vein at the root of the ear.
This will cause its peripheral portion to swell up and increase in
calibre.
5. Hold the syringe as one would a pen and thrust the point of the
needle through the skin and the wall of the vein till it enters the
lumen of the vein (Fig. 189). Now press it onward in the direction of
the blood stream--i. e., toward the body of the animal.
6. Direct the assistant to cease compressing the root of the ear, and
_slowly_ inject the inoculum. (If the fluid is being forced into the
subcutaneous tissue, a condition which is at once indicated by the
swelling that occurs, the injection must be stopped and another attempt
made at a spot closer to the root of the ear or at some point on the
corresponding vein on the opposite ear.)
7. Withdraw the needle and press a pledget of cotton-wool over the
puncture to ensure closure of the aperture in the vein wall.
8. Label, etc.
[Illustration: FIG. 189.--Intravenous inoculation.]
~9. Inhalation.~--
(a) _Fluid Inoculum._--(_Anaesthetic, none._)
1. Place the animal in a closed metal box.