XVII. EXPERIMENTAL INOCULATION OF ANIMALS.

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The use of living animals for inoculation experiments may become a necessary procedure in the Bacteriological Laboratory for some one or more of the following reasons:

A. Determination of Pathogenetic Properties of Bacteria already Isolated in Pure Culture (see page 315).

The exact study of the conditions influencing the virulence (including its maintenance, exaltation and attenuation) of an organism, and precise observations upon the pathogenic effects produced by its entrance into, and multiplication within the body tissues can obviously only be carried out by means of experimental inoculation; whilst many points relating to vitality, longevity, etc., can be most readily elucidated by such experiments.

B. Isolation of Pathogenetic Bacteria.

Certain highly parasitic bacteria (which grow with difficulty upon the artificial media of the laboratory) can only be isolated with considerable difficulty from associated saprophytic bacteria when cultural methods alone are employed; but if the mixture of parasite and saprophytes is injected into an animal susceptible to the action of the former, the pathogenic organism can readily be isolated from the tissues of the infected animal. The pneumococcus for example occurs in the sputum of patients suffering from acute lobar pneumonia, but usually in association with various saprophytes derived from the mouth and pharynx. The optimum medium for the growth of the pneumococcus, blood agar, is also an excellent pabulum for the saprophytes of the mouth, and plate cultures are rapidly overgrown by them to the destruction of the more delicate pneumococcus. But inoculate some of the sputum under the skin of a mouse and three or four days later the pneumococcus will have entered the blood stream (leaving the saprophytes at the seat of inoculation) and killed the animal. Cultivations made at the post-mortem (see page 398) from the mouse's heart blood will yield a pure growth of the pneumococcus.

C. Identification of Pathogenetic Bacteria.

The resemblances, morphological and cultural, existing between certain pathogenetic bacteria are in some cases so great as to completely overwhelm the differences; again the same bacterium may under varying conditions assume appearances so different from those regarded as typical or normal as to throw doubt on its identity. In each case a simple inoculation experiment may decide the point at once. As a concrete example may be instanced an autopsy on an animal dead from an unknown infection. Cultivations from the heart blood gave a pure growth of a typical (capsulated) pneumococcus. Cultivations from the liver gave a pure growth of what appeared to be a typical (non-capsulated) Streptococcus pyogenes longus. The latter inoculated into a rabbit caused the death of the animal from pneumococcic septicÆmia, and cultures from the rabbit's blood gave a pure growth of a typical (capsulated) pneumococcus.

D. Study of the Problems of Immunity.

It is only by a careful and elaborate study of the behaviour of the animal cell and the body fluids vis-À-vis with the infecting bacterium that it becomes possible to throw light upon the complex problem whereby the cell opposes successful resistance to the diffusion of the invading microbe, or succeeds in driving out the microbe subsequently to the occurrence of that diffusion.

At the moment, however, our attention is directed to the first of these broad headings, for it is by the application of the knowledge acquired in its pursuit that we are able to deal with problems arising under any of the remainder.

For whatever purpose the inoculation is performed, it is essential that the experiment should be planned to secure the maximum amount of information and the minimum of discomfort to the animal used. Every care therefore must be taken to ensure that the virus is introduced into the exact tissue or organ selected; and the operation itself must be carried out with skill and expedition, and under strictly aseptic conditions.

In the course of inoculation studies many instances of natural immunity, both racial and individual, will be met with; but it must be recollected that natural immunity is relative only and never absolute, and care be taken not to label an organism as non-pathogenic until many different methods of inoculation have been performed upon different species of animals, combined when necessary with various procedures calculated to overcome any apparent immunity, and have invariably given negative results.

In some countries experiments upon animals are only permitted under direct license from the Government, and then only within premises specially licensed for the purpose. In England this license is in the grant of the Home Secretary, and confers the permission to experiment upon animals under general anÆsthesia, provided that after the experiment is completed the animal must be destroyed before regaining consciousness. If it is intended to carry out simple hypodermic inoculations and superficial venesections, Certificate A, granting this specific permission and dispensing with the necessity for general anÆsthesia must be obtained in addition to the license; whilst if the inoculation entails more extensive operative procedures, and it is necessary to observe the subsequent course of the infection, should such occur, the license must be coupled with Certificate B—since this certificate removes the compulsion to destroy the animal whilst under the anÆsthetic. Further special certificates and combinations of certificates are required if cats, dogs, horses, asses or cattle are to be the subjects of experiment. Under every certificate it is expressly stipulated that if the animal shows signs of pain it must be destroyed immediately.

The animals generally employed in the study of the pathogenic properties of the various micro-organisms are:

Cold Blooded. Warm Blooded. Hot Blooded.
Frog. Mouse. Fowl.
Toad. Rat. Pigeon.
Lizard. Guinea pig.
Rabbit.
Monkey.

Preparation.—Before inoculation, the experimental animals should be carefully examined, to avoid the risk of employing such as are already diseased: since it must be remembered that in a state of nature, as well as in captivity, the animals employed for laboratory inoculations are subject to infection by various animal and vegetable parasites, and in some instances such infection presents no symptoms which are obvious to the casual examination; the sex should be noted, the weight recorded, and the rectal temperature taken. The remaining items of importance are the time of the inoculation, the material that is inoculated, and the method of inoculation, and finally under what authority the experiment is performed. In the author's laboratory these data are entered upon a pink card which forms part of a card index system. The card further provides space for notes on the course of the resulting infection, and carries on the reverse the weight and temperature chart (Figs. 164 and 165).

Fig. 164.—Front of inoculation card. Fig. 164.—Front of inoculation card.

Preliminary Inspection and Examination.—The preliminary examination should comprise observation of the animal at rest and in motion; the appearance of the fur, feathers or scales, inspection of the eyes, and of external orifices of the body; tactile examination of the body and limbs, and palpation of the groins and abdomen; and in many cases the microscopical examination of fresh and stained blood-films.

Some of the commoner forms of naturally acquired infection may be briefly mentioned, without however touching upon the various fleas, lice and ticks which at times infect the ordinary laboratory animals.

Fig. 165.—Back of inoculation card. Fig. 165.—Back of inoculation card.

The Rabbit, particularly in captivity, is subject to attacks of Psoric Acari, and the infection is readily transmitted to rabbits in neighbouring cages and also to guinea pigs, but not to rats and mice. One species (Sarcoptes minor var. cuniculi) gives rise to the ordinary mange. The infection first shows itself as thick yellowish scales and crusts around the nose, mouth and eyes, spreads to the bases and outer surfaces of the ears (never to the inside of the concha), to the fore and hind legs and into the groins and around the genitals. The acari can be readily demonstrated microscopically in scrapings of the skin, treated with liquor potassÆ. Another form of scabies (due to Psoroptes communis cuniculi) commences at the bottom of the concha, which is filled with whitish-yellow masses consisting of dried crusts, scales, fÆces, and dead acari. The base of the ear is hard and swollen, and lifting the animal by the ears—as is usually done—gives rise to considerable pain; indeed this symptom may be the one which first attracts attention to an infection, which causes progressive wasting and terminates in death. A mixed infection—sarcoptic plus psorotic acariasis—is sometimes seen.

If it is decided to try and save animals suffering from infection by these parasites, they must be segregated, the scabs carefully cleaned from the infected areas and the denuded surfaces washed with 5 per cent. solution of Potassium persulphate (a few drops being allowed to run into the concha), or with a preparation containing equal parts of soft paraffin and vaseline with a few drops of lysol. This treatment should be repeated daily until the acarus is destroyed and the animal has regained its normal condition. The cages should be disinfected and all neighbouring animals carefully examined, and any which show signs of infection should be treated in a similar manner. Favus also attacks the rabbit, and the typical spots are first noted around the base of the ear.

Infection by Coccidium oviforme is very common, without however presenting any symptoms by which the infection may be recognised. Usually the condition is only noted post-mortem, when the liver is found to be studded with numerous cascating tubercles, which on examination prove to be cystic areas crowded with coccidia. Sometimes too the liver of a rabbit dead from some intentional or accidental bacterial infection is found at the post-mortem to be marked by fine yellowish streaks and small tubercles due to the embryos of TÆnia serrata, while the cystic form (Cysticercus pisiformis) is often noted free in the peritoneal cavity, or invading the mesentery.

Abscess formation from infection with ordinary pyogenic bacteria occurs naturally in the rabbit, and frequently the animal house of a laboratory is decimated by an infective septicÆmia due to B. cuniculicida.

The Mouse and Rat suffer from septicÆmia, and from the cysticercus form of TÆnia murina; the cystic form (Cysticercus fasciolaris) of T. crassicollis has its habitat in their livers. These small rodents are frequently infected with scabies, but if freely provided with clean straw will clean themselves by rubbing through it. The mouse is also attacked by favus, and the rat is often infected with Trypanosoma Lewisi.

The Guinea pig, like the rabbit, suffers from scabies and coccidiosis. In addition it is often naturally infected with B. tuberculosis, and it is a wise precaution to test animals as soon as they reach the laboratory by injecting Koch's Old Tuberculin—0.5 c.c. causing death in the tuberculous cavy within 48 hours.

The Monkey is naturally prone to tuberculosis, and should be injected with 1 c.c. Old Tuberculin on arrival in the laboratory. The tissues of the monkey also serve as the habitat for a Nematode worm parasitic in cattle (Œsophagostoma inflatum) resembling the Anchylostomum, and this parasite frequently bores through the intestinal wall, and provokes the formation of small cysts in the immediately adjacent mesentery. The presence of these cysts may give rise to considerable speculation at the post-mortem.

The Pigeon may be infected by HÆmosporidia, and its blood show the presence of halteridia. This bird may also be the subject of a bacterial infection known as pigeon diphtheria; while the fowl may be subject to scabies and ringworm, or suffer from fowl cholera or fowl septicÆmia—infections due to members of the hÆmorrhagic septicÆmia group.

Weighing.—The larger animals are most conveniently weighed in a decimal scale provided with a metal cage for their reception instead of the ordinary pan (Fig. 166). Mice and rats are weighed in a modification of the letter balance, weighing to 250 grammes, which has a conical wire cage, (carefully counterpoised) substituted for its original pan (Fig. 167).

Fig. 166.—Rabbit scales. Fig. 166.—Rabbit scales.

Temperature.—To take the rectal temperature of any of the laboratory animals, the animal should be carefully and firmly held by an assistant. Introduce the bulb of an ordinary clinical thermometer, well greased with vaseline, just within the sphincter ani. Allow it to remain in this position for a few seconds, and then push it on gently and steadily until the entire bulb and part of the stem, as far as the constriction, have passed into the rectum. Three to five minutes later, the time varying of course with the sensibility of the thermometer used, withdraw the instrument and take the reading. The thermometers employed for recording temperature should be verified from time to time by comparison with a standard Kew certified Thermometer kept in the laboratory for that purpose.

Fig. 167.—Mouse scales Fig. 167.—Mouse scales

Cages.—During the period which elapses between inoculation and death, or complete recovery, the experimental animals must be kept in suitable receptacles which can easily be kept clean and readily disinfected.

The mouse is usually stored in a glass jar (Fig. 168) 11 cm. high and 11 cm. in diameter, closed by a wire gauze cover which is weighted with lead or fastened to the mouth of the jar by a bayonet catch. A small oblong label, 5 cm. by 2.5 cm., sand-blasted on the side of the cylinder, is a very convenient device as notes made upon this with an ordinary lead pencil show up well and only require the use of a damp cloth to remove them (Fig. 168).

The rat is kept under observation in a glass jar similar, but larger, to that used for the mouse.

Fig. 168.—Mouse jar. Fig. 168.—Mouse jar.
Fig. 169.—Tripod. Fig. 169.—Tripod.

A layer of sawdust at the bottom of the jar absorbs any moisture and cotton-wool or paper shavings should be provided for bedding. The food should consist of bran and oats with an occasional feed of bread-and-milk sop.

The use of a metal tripod, on the platform of which are soldered two small cups for the reception of the food, inside the cage, prevents waste of food or its contamination with excreta (Fig. 169).

After use the jars and tripods are sterilised either by chemical reagents or by autoclaving.

The rabbit and the guinea-pig are confined in cages of suitable size, made entirely of metal (Fig. 170). The sides and top and bottom are of woven wire work; beneath the cage is a movable metal tray filled with sawdust, for the reception of the excreta. The cage as a whole is raised from the ground on short legs. The sides, etc., are generally hinged so that the cage packs up flat, for convenience of storing and also of sterilising.

The ordinary rat cage, a rectangular wire-work box, 30 cm. from front to back, 20 cm. wide, and 14 cm. high, makes an excellent cage for guinea-pigs if fitted with a shallow zinc tray, 35 cm. by 24 cm., for it to stand upon.

Fig. 170.—Metal rabbit rage. Fig. 170.—Metal rabbit rage.

A plentiful supply of straw should be provided for bedding and the food should consist of fresh vegetables, cabbage leaves, carrot and turnip tops and the like for the morning meal and broken animal biscuits for the evening meal. Occasionally a little water may be placed in the cage in an earthenware dish.

The tray which receives the dejecta should be cleaned out and supplied with fresh sawdust each day, and the soiled sawdust, remains of food, etc., should be cremated.

These cages are sterilised after use either by autoclaving or spraying with formalin.

As animal inoculation is purely a surgical operation, the necessary instruments will be similar to those employed by the surgeon, and, like them, must be sterile. In the performance of the inoculation strict attention must be paid to asepsis, and suitable precautions adopted to guard against accidental contamination of the material to be introduced into the animal. In addition, the hands of the operator should be carefully disinfected.

The list of apparatus used in animal inoculations given below comprises practically everything needed for any inoculation. Needless to remark, all the apparatus will never be required for any one inoculation.

Fig. 171.—Hypodermic syringe with finger rests. Fig. 171.—Hypodermic syringe with finger rests.

Apparatus Required for Animal Inoculation:

1. Water steriliser (vide page 33). It is also convenient to have a second water steriliser, similar but smaller (23 by 7 by 5 cm.), for the sterilisation of the syringes.

2. Injection syringe. The best form is one of the ordinary hypodermic pattern, 1 c.c. capacity graduated in twentieths of a cubic centimeter (0.05 c.c.), fitted with finger rests, but with the leather washers and the packing of the piston replaced by those made of asbestos (Fig. 171). The instrument must be easily taken to pieces, and spare parts should be kept on hand to replace accidental breakage or loss. Other useful syringes are those of 2 c.c., 5 c.c., 10 c.c., and 20 c.c. capacity. A good supply of needles must be kept on hand, both sharp-pointed and with blunt ends. To sterilise the syringe, fill it with water, loosen the packing of the piston and all the screw joints, place it in the steriliser and boil for at least five minutes. Disinfect the syringe after use, in a similar manner. The needles, which are exceedingly apt to rust after being boiled, should be stored in a pot of absolute alcohol when not in use.

3. Operating table.

4. Surgical instruments. Sterilise these before use by boiling, and disinfect them after use by the same means. Wipe perfectly dry immediately after the disinfection is completed.

Scissors, probe and sharp-pointed.

Dissecting forceps of various patterns.

Pressure forceps.

Retractors (small self retaining Fig. 172).

Aneurism needles, sharp and blunt.

Scalpels, } Keratomes, } with metal handles. Trephines, }

Michel's steel clips and special forceps for applying the same. These small steel clips enable the operator to easily and rapidly close skin incisions and are most satisfactory for animal operations.

Surgical needles.

Needle holder.

Soft rubber catheters, various sizes.

Gum elastic oesophageal bougies with connection to fit syringe.

Fig. 172. Small self retaining retractors. Fig. 172. Small self retaining retractors.

5. AnÆsthetic.

(a) General: The safest general anÆsthetic for animals is an A. C. E. mixture, freshly prepared, containing by volume alcohol 1 part, chloroform 2 parts, ether 6 parts, and should be administered on a "cone" formed by twisting up one corner of a towel and placing a wad of cotton-wool inside it, or from a saturated cotton-wool pad packed into the bottom of a small beaker.

(b) Local:

1. Cocaine hydrochloride, 2 per cent. in adrenalin 1 per mille solution.
2. Beta-eucaine, 2 per cent. in adrenalin, 1 per mille solution.
3. Ethyl chloride jet.

6. Sterile glass capsules of various sizes.

7. Cases of sterile pipettes { 10 c.c. (in tenths of a cubic centimetre).
{ 1 c.c. (in hundredths of a cubic centimetre).

8. Flasks (75 c.c.) containing sterilised normal saline solution (or sterile bouillon).

9. Sterilised cotton-wool. Cotton-wool (absorbent) is packed loosely in a copper cylinder similar to that used for storing capsules, and sterilised in the hot-air oven.

10. Sterilised gauze. Gauze is sterilised in the same way as cotton-wool.

11. Sterilised silk and catgut for sutures. These are sterilised, as required, by boiling for some ten minutes in the water steriliser.

12. Flexible collodion (or compound tincture of benzoin).

13. Grease pencil.

14. Tie-on celluloid labels, to affix to the cages.

15. Razor.

16. Small pot of warm water.

17. Liquid soap. Liquid soap is prepared as follows: Measure out 100 grammes of soft soap and add to 500 c.c. of 2 per cent. lysol solution in a large glass beaker; dissolve by heating in a water-bath at about 90° C. Bottle and label "Liquid Soap."

18. In place of the liquid soap and razor it is sometimes convenient to use a Depilatory powder.

Barium sulphide 1 part
Rice starch 3 parts

Dust the powder thickly over the area to be denuded of hair, sprinkle with water and mix into a thin paste in situ; allow the paste to act for three minutes, then scrape off with a bone spatula—the hair comes away with the paste and leaves a perfectly bare patch. This process is preferably carried out, the day previous to the operation.

Material Utilised for Inoculation.—The material inoculated may be either—

1. Cultures of bacteria—grown in fluid media, or on solid media.

2. Metabolic products of bacterial activity—e. g., toxins in solution.

3. Pathological products (fluid secretions and excretions, solid tissues).

The Preparation of the Inoculum.

(a) Cultivations in Fluid Media.

1. Flame the plug of the culture tube.

2. Remove the plug and flame the mouth of the tube.

3. Slightly raise the lid of a sterile capsule, insert the mouth of the culture tube into the aperture and pour some of the cultivation into the capsule.

4. Remove the mouth of the culture tube from the capsule, replace the lid of the latter, flame the mouth of the tube, and replug.

5. Remove the syringe from the steriliser, squirt out the water from its interior, and allow to cool.

6. Raise the lid of the capsule sufficiently to admit the needle of the syringe and draw the required amount of the cultivation into the barrel of the syringe.

(Or, remove a definite measured quantity of the cultivation directly from the tube or flask by means of a sterile graduated pipette, discharge the measured amount into a sterile capsule, and fill into the syringe; or take up the required quantity of the cultivation directly into the graduated syringe from the tube or flask.)

Fig. 173.—Conical separatory funnel, fitted for injection of fluid cultivations. Fig. 173.—Conical separatory funnel, fitted for injection of fluid cultivations.

If it is necessary to introduce a large bulk of fluid into the animal, 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.

Fig. 174.—Arrangement of pressure injection apparatus. 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.

Fig. 175.—Holding rabbit for shaving. 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 anÆsthetic 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.

Fig. 176.—Taking guinea-pig's temperature. 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.

Fig. 177.—Voge's holder. Fig. 177.—Voge's holder.

When the animal is anÆsthetised, 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).

Fig. 178.—Mouse holder. Fig. 178.—Mouse holder.
Fig. 179.—Tatin's operation table. 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° 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.

Fig. 180.—Author's operating table 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.—(AnÆsthetic, 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.—(AnÆsthetic, 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.—(AnÆsthetic, 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.

Fig. 181.—Glass tube syringe for subcutaneous "solid" inoculation. 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.—(AnÆsthetic, 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.—(AnÆsthetic, A. C. E.)

1. Secure the animal to the operation table and anÆsthetise.

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.—(AnÆsthetic, 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).

Fig. 182.—Intraperitoneal inoculation—fluid. Fig. 182.—Intraperitoneal inoculation—fluid.

8. Inject the fluid from the syringe.

9. Label, etc.

Fig. 183.—Section of abdominal wall, etc., showing point of needle lying free in the peritoneal cavity above the coils of intestine. 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. AnÆsthetise 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 anÆsthetic 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.

Fig. 184.—Platinum wire for burning hole through parietes. Fig. 184.—Platinum wire for burning hole through parietes.

(b) Solid Inoculum (or the implantation of capsules containing fluid cultivations).—(AnÆsthetic, A. C. E.)

1. AnÆsthetise 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° 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.

Fig. 185.—Making celloidin capsules. 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.—(AnÆsthetic, A. C. E.)

Fig. 186.—Guarded trephine. 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. AnÆsthetise the animal and secure it to the operating table, dorsum uppermost.

2. Shave a portion of the scalp immediately in front of the ears.

Fig. 187.—Guy's electrical dental engine. 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.

Fig. 188.—Intracranial inoculation of rabbit. The circle indicates the situation of the trephine hole. Fig. 188.—Intracranial inoculation of rabbit. The circle indicates the situation of the trephine hole.

6. Intraocular.

(a) Fluid Inoculum.—(AnÆsthetic, 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.—(AnÆsthetic, A. C. E.)

1. AnÆsthetise 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.—(AnÆsthetic, 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.—(AnÆsthetic, 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 anÆsthetic 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.

Fig. 189.—Intravenous inoculation. Fig. 189.—Intravenous inoculation.

9. Inhalation.

(a) Fluid Inoculum.—(AnÆsthetic, none.)

1. Place the animal in a closed metal box.

2. Through a hole in one side introduce the nozzle of some simple spraying apparatus, such as is used for nasal medicaments.

3. Fill the reservoir of the instrument (previously sterilised) with the fluid inoculum, and having attached the bellows, spray the inoculum into the interior of the box.

4. On the completion of the spraying, open the box, spray the animal thoroughly with a 10 per cent. solution of formaldehyde (to destroy any of the virus that may be adhering to fur or feathers).

5. Transfer the animal to its cage.

6. Label, etc.

7. Thoroughly disinfect the inhalation chamber.

(b) Fluid or Powdered Inoculum.AnÆsthetic, A. C. E.

1. AnÆsthetise the animal and secure it firmly to the operating table.

Fig. 190.—Gag for rabbits. Fig. 190.—Gag for rabbits.

2. Prop open the mouth by means of some form of gag; seize the tongue with a pair of forceps and draw it forward.

The most convenient form of gag for the rabbit or cat is that shown in Fig. 190. It is simply a strip of hard wood shaped at the middle and provided with a square orifice through which a tracheal or oesophageal tube can be passed.

3. Pass a previously sterilised glass tube (17 cm. long, 0.5 cm. diameter, with its terminal 2 cm. slightly curved) down through the larynx into the trachea.

4. Connect the straight portion of a Y-shaped piece of tubing to the upper end of the sterilised tube and couple one branch of the Y to a separatory funnel containing the fluid inoculum, or insufflator containing the powdered inoculum, and the other to a hand bellows.

5. Allow the fluid inoculum to run into the lungs by gravity, or blow in the powdered inoculum by means of a rubber-ball bellows.

6. Remove the intratracheal tube; release the animal from the table.

7. Label, etc.

As an alternative method in the case of fairly large animals, such as rabbits, etc., a sterile piece of glass tubing of suitable diameter may be passed through the larynx down the trachea almost to its bifurcation. Fluid cultivations may then be literally poured into the lungs, or cultivations, dried and powdered, may be blown into the lung by the aid of a small hand bellows or even a teat pipette.

10. Intragastric Inoculation.Fluid or semi-fluid inoculum. (AnÆsthetic none.)

The method of performing the operation is varied slightly according to the size of the experimental animal.

A. Monkey, Rabbit, Guinea-pig.

1. Secure the animal to the operating table ventral surface uppermost.

2. Prop the mouth open with a gag; draw the tongue forward with forceps.

3. Sterilise a soft rubber catheter (No. 10 or 8 English scale, or No. 18 or 15 French) and lubricate it with sterile glycerine.

4. Pass it to the back of the pharynx, keeping the end in the middle line.

5. Gently assist the progress of the catheter down the oesophagus until it passes the cardiac orifice of the stomach. Do not use any force.

6. Take up the required dose of inoculum into a sterilised pipette. Insert the point of the pipette into the open end of the catheter and allow the fluid to run down into the stomach. Remove the pipette and drop it into a jar of lysol.

7. With another sterile pipette run one cubic centimetre of sterile saline solution through the catheter to wash out the last traces of the inoculum.

8. Withdraw the catheter.

9. Label, etc.

B. Rats and Mice (Mark's Method).

1. Secure the animal in the vertical position.

(a) Rat.—Take a pair of catch sinus forceps about 22 cm. in length and seize the animal by the loose skin of the head as far forward as possible—fix the forceps, and holding the instrument vertically upward, transfer to the left hand of an assistant who secures the animal's tail between the fingers grasping the handle of the forceps. (See Fig. 191.)

Fig. 191.—Intragastric inoculation of rat. Fig. 191.—Intragastric inoculation of rat.

(b) Mouse.—An assistant grasps the loose skin between the ears as far forwards as possible between the forefinger and thumb of the left hand. He now grasps the tail with the right hand, draws the mouse straight and passes the tail between the fourth and little fingers of the left hand and secures it there.

2. The assistant takes a closed pair of thin-bladed forceps in his right hand, passes the ends into the animal's mouth, then allows the blades to separate. This opens the animal's jaw and serves as a gag.

3. Moisten the sterilised oesophageal tube with sterile water. (This tube is of silk rubber, 6.5 cm. in length, with the distal end rounded, the proximal end mounted in a syringe needle head, which fits the nozzles of the two sterile syringes to be used.)

4. Grasp the tube about its middle and pass it into the animal's mouth, downwards and a little to one side or the other until its length is lost in the digestive tract and mouth. Gentle guidance is alone necessary. Do not use any force.

5. Take up the required dose of inoculum into the syringe; insert the nozzle of the syringe into the needle-mount, and force the piston down.

6. Steadying the needle-mount with the left hand, detach the syringe.

7. Draw up some sterile water in the second (sterile) syringe, and inserting its nozzle into the needle-mount force a few drops of water through the tube to wash it out.

8. With one quick upward movement remove the tube from the animal's mouth.

9. Label, etc.

One other method of inoculation remains to be described, which does not require operative interference.

11. Feeding.

1. Fluid Inoculum.—Small pieces of sterilised bread or sop (sterilised in the steamer at 100° C.) are soaked in the fluid inoculum and offered to the animals in a sterile Petri dish or capsule.

2. Solid Inoculum.—Small pieces of tissue are placed in sterile vessels and offered to the animals.

FOOTNOTES:

[12] This table is made by Messrs. Down Bros., St. Thomas's Street, London, S. E.

[13] This modification is made for the author by Messrs. Down Bros., St. Thomas's Street, London, S. E.

[14] Manufactured by Messrs. Francis Lepper, 56, Great Marlborough Street, London, W.


                                                                                                                                                                                                                                                                                                           

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