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THE TITRATION OF HAEMOLYTIC SERUM.
Apparatus Required:
Electrical centrifuge. Sterile centrifuge tubes. Water-bath regulated at 56 deg. C. Sterilised pipettes 10 c.c. graduated in tenths. Sterilised pipettes 1 c.c. graduated in tenths. Sterile test-tubes, 16 x 2 cm. Small sterile test-tubes, 9 x 1 cm. Small test-tube rack, or roll of plasticine. Capillary teat pipettes. Stout rubber band or length of small rubber tubing.
Reagents Required and Method of Preparation:
1. Normal saline solution.
2. Haemolytic serum inactivated by preliminary heating to 56 deg. C. for 15 minutes (vide supra) in test-tube labelled H. S.
3. Complement. Fresh guinea-pig serum in test-tube labelled C.
Kill a normal guinea-pig with chloroform vapour.
Open the thorax with all aseptic precautions, and collect as much blood as possible from the heart with a sterile Pasteur pipette.
Transfer it to a sterile centrifuge tube and place the tube in the incubator at 37 deg. C. Two hours later separate the clot from the sides of the tube, and centrifugalise thoroughly.
Pipette off the clear serum to a clean sterilised test-tube.
4. Erythrocyte solution, in test-tube labelled E.
Collect and wash human red blood cells (see page 388, 1-8). Measure the volume of red cells available and prepare a 2 per cent. suspension in normal saline solution.
METHOD.—
1. Take two test-tubes and number them 1 and 2, and pipette into each 9 c.c. of normal saline solution.
2. Add 1 c.c. of haemolytic rabbit serum to tube No. 1 and mix thoroughly: take up 1 c.c. of the mixture and add it to tube No. 2; mix thoroughly.
3. Set up ten small test-tubes in test-tube rack or in roll of plasticine, and number 1 to 10.
4. Pipette into tube No. 1 0.5 c.c. = 0.5 c.c.} haemolytic serum } From tube Pipette into tube No. 2 0.1 c.c. = 0.1 c.c. } H. S. haemolytic serum }
Pipette into tube No. 3 0.5 c.c. = 0.05 c.c. } haemolytic serum } Pipette into tube No. 4 0.3 c.c. = 0.03 c.c. } haemolytic serum } From Pipette into tube No. 5 0.2 c.c. = 0.02 c.c. } tube 1. haemolytic serum } pipette into tube No. 6 0.1 c.c. = 0.01 c.c. } haemolytic serum }
Pipette into tube No. 7 0.5 c.c. = 0.005 c.c. } haemolytic serum } Pipette into tube No. 8 0.3 c.c. = 0.003 c.c. } haemolytic serum } From Pipette into tube No. 9 0.2 c.c. = 0.002 c.c. } tube 2. haemolytic serum } Pipette into tube No. 10 0.1 c.c. = 0.001 c.c. } haemolytic serum }
5. To each tube add 1 c.c. of erythrocyte solution.
6. When necessary (that is to say in tubes 2, 4, 5, 6, 8, 9 and 10) add normal saline solution to the mixture in the test-tubes till the column of fluid in each reaches to the same level.
7. Shake each tube in turn, so as to thoroughly mix its contents. Plug the mouth of each tube with cotton wool, and place entire set in the incubator at 37 deg. C. for one hour.
8. Remove the tubes from the incubator and into each tube pipette 0.1 c.c. complement (guinea-pig's serum) and replace tubes in incubator at 37 deg. C. for further period of one hour.
9. Remove the tubes from the incubator, and if complete haemolysis has not taken place in every tube, stand on one side, preferably in the ice chest, for an hour.
10. Then examine the tubes.
Complete haemolysis is indicated by a clear red solution, with no deposit of red cells at the bottom of the test-tube.
Absence of haemolysis is indicated by a clear or turbid colourless fluid, with a deposit of red cells at the bottom of the test-tubes.
The smallest amount of haemolytic serum that has caused complete haemolysis is known as the minimal haemolytic dose (M. H. D.) and if haemolysis has occurred in all the tubes down to No. 7—the m. h. d. of this particular serum is .005 c.c. = 200 minimal haemolytic doses per cubic centimetre. Such a serum is strong enough for experimental work; indeed, for many purposes, complete haemolysis down to tube 6 will indicate a serum sufficiently strong(= 100 m. h. d. per cubic centimetre). If, however, only the first one or two tubes are completely haemolysed, this is an indication that the rabbit should receive further injections in order to raise the haemolytic power to a sufficiently high level.
STORAGE OF HAEMOLYSIN.
If, and when the haemolysin content of the rabbit's serum is found to be sufficient, destroy the animal by chloroform vapour.
Remove as much of its blood as possible from the heart under aseptic precautions into sterilized centrifuge tubes.
Transfer the tubes of blood to the incubator at 37 deg. C. for two hours—then centrifugalize thoroughly.
Pipette off the clear serum, and fill in quantities of 1 c.c., into small glass ampoules or pipettes, and hermetically seal in the blowpipe flame, care being taken to avoid scorching the serum.
Place the ampoules when filled with serum and sealed, in a water-bath at 56 deg. C. for 30 minutes. This destroys the complement, i. e., inactivates the serum, and at the same time, provided the various operations have been carried out under aseptic precautions, ensures its sterility. A longer exposure reduces the haemolytic power.
Place the ampoules in a closed metal box and store in the ice chest for future use.
FOOTNOTES:
[10] The quantities here given are not absolutely correct. If exactitude is essential the student must calculate the amount required by the aid of the Percentage Formula, Appendix, page 496.
[11] See Percentage Formula, Appendix, page 496.
XVII. EXPERIMENTAL INOCULATION OF ANIMALS.
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 septicaemia, 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-a-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 anaesthesia, 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 anaesthesia 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 anaesthetic. 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).
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.
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 potassae. 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, faeces, 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 Taenia 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 septicaemia due to B. cuniculicida.
The Mouse and Rat suffer from septicaemia, and from the cysticercus form of Taenia 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 (Oesophagostoma 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 Haemosporidia, 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 septicaemia—infections due to members of the haemorrhagic septicaemia 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).
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.
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.
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.
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.
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.
5. Anaesthetic.
(a) General: The safest general anaesthetic 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 deg. 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.)
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.
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.
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.
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.
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).
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.
.]
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.
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).
8. Inject the fluid from the syringe.
9. Label, etc.
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.
(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.
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.)
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.
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.
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.
9. Inhalation.—
(a) Fluid Inoculum.—(Anaesthetic, 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.—Anaesthetic, A. C. E.
1. Anaesthetise the animal and secure it firmly to the operating table.
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. (Anaesthetic 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.)
(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 deg. 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.
XVIII. THE STUDY OF EXPERIMENTAL INFECTIONS DURING LIFE.
The possession of pathogenetic properties by an organism under study is indicated by the "infection" of the experimental animal—a term which is employed to summarise the condition resulting from the successful invasion of the tissues of the experimental animal by the micro-organisms inoculated and by their multiplication therein. Infection is considered to have taken place:
1. When the death of the animal is produced as a direct consequence of the inoculation.
2. When without necessarily producing death the inoculation causes local or general changes of a pathological character.
3. When either with or without death, or local or general changes occurring, certain substances make their appearance in the body fluids, which can be shown (in vitro or in vivo) to exert some profound and specific effect when brought into contact with subcultivations of the organism originally inoculated.
The important factors in the production of infection are:
A. Seed. Virulence of organism. Dose of organism.
B. Soil. Resistance offered by the cells of the experimental animal.
The first two factors, although variable, are to a certain extent under the control of the experimenter. Thus by suitable means the virulence of an organism can be exalted or attenuated, whilst the size of the dose may be increased or diminished. The third factor also varies, not only amongst different species of animals, but also amongst different individuals of the same species. The essential causes of this variation are not so obvious, so that beyond selecting the animals intended for similar experiments with regard to such points as age, size or sex, but little can be done to standardise cell resistance.
Immediately an animal has been inoculated a period of clinical observation must be entered upon, which should only terminate with the death of the animal. The general observations should at first and if the infection is an acute one, be made daily—later, and if the animal appears to be unaffected or if the infection is chronic, both general and special observations should be carried out at weekly intervals. If the animal appears to be still unaffected, it should be killed with chloroform vapour at the end of two or three months and a complete post-mortem carried out.
A. The general observations should take cognisance of:
1. General appearance. The experimental animal should be inspected daily, not only with a view to detecting symptoms due to the experimental infection, but also to prevent any intercurrent infection, naturally acquired, from escaping notice (vide page 337).
2. The weight of the inoculated animal should be observed and recorded each day during the course of an experimental infection at precisely the same hour, preferably just before the morning feed.
3. The temperature should similarly be recorded daily, if not more frequently, during the whole period the animal is under observation, and carefully charted—individual variations will at once become apparent. It should be borne in mind that the temperature regarded as normal for man (37.5 deg. C.) is not the normal average temperature of any of the lower animals save the rat and mouse. The accompanying table of normal averages for the animals usually employed in bacteriological research may be of use in preventing the erroneous assumption that pyrexia is present in an animal, which merely shows its own normal temperature.
NORMAL AVERAGES. Rectal Pulse. Respirations. Animal. Temp. deg. C. Rate per minute. Frog 8.9-17.2 80 12 Mouse 37.4 120 ... Rat 37.5 ... 210 Guinea pig 38.6 150 80 Rabbit 38.7 135 55 Cat 38.7 130 24 Dog 38.6 95 15 Goat 40.0 75 16 Ox 38.8 45 .. Horse 37.9 38 11 Monkey (Rhesus) 38.4 100 19 Pigeon 40.9 136 30 Fowl 41.6 140 12
B. Special observations comprise some or all of the following, according to the method of inoculation and the character of the virus.
1. The site of inoculation should be minutely examined at least at weekly intervals, and the neighbouring lymphatic glands palpated.
2. Any local reaction at the site of inoculation and any other readily accessible lesion should be carefully investigated. Any suppurative process which may occur, whether in the subcutaneous tissues or in joints, should be explored and the pus carefully examined both microscopically and culturally.
Fluid secretions and excretions, such as pus or serous exudates when accessible are collected direct from the body in sterile capillary pipettes (vide Fig. 13a,) in the following manner:
1. Open the case containing the pipettes, grasp one by the plugged end, remove it from the case, and replace the lid of the latter.
2. Attach a rubber teat (vide page 10) to the plugged end of the pipette and use the teat as the handle of the pipette.
3. Pass the entire length of the pipette twice or thrice through the flame of the Bunsen burner.
4. Snap off the sealed end of the pipette with a pair of sterile forceps.
5. Compress the india-rubber teat, thrust the point of the pipette into the secretion; now relax the pressure on the teat and allow the pipette to fill.
6. Remove the point of the pipette from the secretion, allow the fluid to run a short distance up the capillary stem and seal the point of the pipette in the flame. (If using a pipette with a constriction below the plugged mouthpiece (Fig 13b), this portion of the pipette may also be sealed in the flame.)
When ready to examine the morbid material snap off the sealed end of the pipette with sterile forceps and eject the contents of the pipette into a sterile capsule. The material can now be utilized for cover-slip preparations, cultivations and inoculation experiment.
3. The peripheral blood should be examined from time to time for from this tissue is often obtained the fullest information as to the course and progress of an infection.
a. The histological examination of the blood should be directed chiefly to observations on the number and kind of white cells; and since but few bacteriologists are at the same time expert comparative haematologists, some notes on the normal characters of the blood of the commoner laboratory animals, contrasted with those of man, are inserted for reference. These have been very kindly compiled for me by my friend and one time colleague Dr. Cecil Price Jones.
COMPARATIVE HAEMOCYTOLOGY OF LABORATORY ANIMALS.
Totals Percentages Animal Hb, Lympho- Large Poly- Eosin- Mast Red cells White per cytes, monos, morph, oph, cells, cells cent. per per per per per cent. cent. cent. cent. cent. Frog 490,000 8,000 58 40 10.0 22.0 15 13 Mouse 8,700,000 8,000 78 60 21.5 17.0 1.4 0.1 Rat 9,000,000 9,000 85 54 7.0 37.5 1.3 0.2 Guinea- pig 5,700,000 10,000 99 55 9.0 32.8 3.0 0.2 Rabbit 6,000,000 7,000 70 50 2.0 46.0 0.6 1.4 Rhesus 4,500,000 13,000 77 43 5.0 50.0 1.3 0.7 Goat 14,600,000 15,000 58 35 6.3 56.7 1.25 0.75 Fowl 3,500,000 30,000 100 49 3.0 42.0 1.0 5.0 Pigeon 3,500,000 20,000 101 43 9.0 43.0 3.0 2.0 Man (adult) 5,000,000 7,500 100 25 5.5 65 4.0 0.5 Normal (4.5-5) (7-9) (95- (20-30) (4-8) (55- (3-5) (0.5-2) limits. millions. thou- 101) 68) sands.
The above table represents in each case the average of a large number of counts.
REMARKS.
Frog.—The red cells are large oval nucleated (20-25 mu by 12-15 mu) discs, the nucleus relatively small and irregularly elongated or oval, about 10 mu in length. Many primitive and developing forms are usually observed—also free nuclei and many cells in various stages of degeneration. Haemoglobin estimation is difficult owing to turbidity of the blood after dilution with water. The polymorphonuclear leucocytes are large cells, about 20 mu; no definite granules can be observed. The eosinophile cells contain large deeply staining coccal-shaped granules.
Mouse.—The granules of the polymorphonuclear leucocytes are usually not stained, or only very faintly so. The nucleus of the eosinophile cell is ring-shaped or much divided, and the granules are coccal and stain oxyphile. The remarkable character of the blood is the high percentage of large mononuclear cells.
Rat.—The fine rod-shaped granules of the polymorphonuclear leucocytes are usually very faintly stained. The granules of eosinophile cells are well stained and coccal-shaped, the nucleus is often ring shaped. The basophile granular cells are few—but the granules are large, and stain deeply basophile.
Guinea-pig.—Polychromasia and punctate basophilia of red cells are very commonly observed—nucleated red cells are also frequent. The large mononuclear cells often contain vacuoles—"Kurlow cells"—possibly of a parasitic nature.
Rabbit.—It is not uncommon to find nucleated red cells in films from quite healthy animals. The granules of the polymorphonuclear leucocytes stain oxyphile. The coarse granules of the eosinophile cells appear to stain less deeply oxyphile, probably owing to the basophile staining of the cytoplasm.
Rhesus monkey.—The blood cells resemble those met with in human blood. The minute neutrophile granules of the polymorphonuclear leucocytes are often very scanty, and sometimes apparently absent. The eosinophile cells are not so densely packed with coarse oxpyhile granules as in the human eosinophile, and the nuclei of these cells are usually much divided, or polymorphous.
Goat.—The red cells are small, nonnucleated discs, only about 4.5 mu diameter, not much more than half that of the human red cell. The polymorphonuclear leucocytes have only a few very minute coccal-shaped oxyphile granules, the nucleus is polymorphous. The eosinophile cells are large cells up to 20 mu, the cytoplasm is basophile and contains coarse coccal-shaped oxyphile granules, and the nucleus is often much divided.
Fowl.—The red cells are oval nucleated discs about 12 mu by 6 mu, the nucleus being relatively small (about 4 mu long), irregularly elongated or oval; round, more deeply stained cells with round or diffuse nuclei, also free nuclei and degenerated forms of red cells are often present. The granules of the cells corresponding to the polymorphonuclear leucocytes are rod-shaped, often beaded or with clubbed ends. The nucleus is not polymorphous, but usually divided into two, though it may be single. The cells probably corresponding to eosinophile leucocytes have fine coccal-shaped granules, faintly staining eosinophile or neutrophile. The basophile granules of the "mast" cells are coccal-shaped, of various size—often quite powdery.
Pigeon.—Red cells resemble those of the fowl, and similar varieties of appearance may be noted. The granules of those cells which correspond to polymorphonuclear leucocytes are rod-shaped, but smaller and finer than in the fowl, and do not show clubbed appearances. The nucleus is not polymorphous, and only occasionally divided. The coccal-shaped granules of the eosinophile cells are stained more deeply oxyphile than those of the corresponding cells of the fowl.
The preparation of dried films for this histological examination of the blood is carried out as follows:
1. Small samples of blood for the preparation of blood films are most conveniently obtained from the veins of the ear in most of the ordinary laboratory animals, viz., monkey, goat, dog, cat, rabbit, guinea-pig; in the pigeon and fowl the axillary vein should be punctured; in the rat and mouse either a vein in the ear or preferably by wounding the tip of the tail; in the frog, the web of the foot should be selected.
2. Puncture the selected vein with a sharp needle. A flat Hagedorn needle (size No. 8) with a cutting edge is the most useful for this purpose. If the vein cannot be distended by proximal compression, vigourous friction with a piece of dry lint may have the desired effect—or a test-tube full of water at about 40 deg. C. may be placed close to the vein. Failing these methods, a drop or two of xylol may be dropped on the skin just over the vein, left on for a few seconds and then wiped off with a piece of dry lint.
3. One of the short ends of a 3 by 1 glass slip is brought into contact with the exuding drop of blood, so that it picks up a small drop.
4. The slide is then lowered transversely on to the surface of a second 3 by 1 slip, which rests on the bench near to one end at an angle of about 45 deg., and retained in this position for a few seconds, while the drop of blood spreads along the whole of the line of contact (see also Fig. 69).
5. Draw the first slide firmly and evenly along the entire length of the lower slide, leaving a thin regular film which will probably show the blood cells only one layer thick.
6. Allow the film to dry in the air.
7. Stain with one of the polychrome blood stains (see page 97).
8. Examine microscopically.
b. The bacteriological examination of the blood is directed solely to the demonstration of the presence in the circulating blood of the organisms previously injected into the animal. For this purpose several cubic centimetres of blood should be taken in an all-glass syringe from an accessible vein corresponding to one of those suggested as the site of intravenous inoculation—and under similar aseptic precautions.
1. Sterilise an all-glass syringe of suitable size, and when cool draw into the syringe some sterile sodium citrate solution and moisten the whole of the interior of the barrel; then eject all the citrate solution if less than 5 c.c. blood are to be withdrawn; if more than 5 c.c. are required retain about half a cubic centimetre of the fluid in the syringe. This prevents coagulation of the blood.
The sodium citrate solution is prepared by dissolving:
Sodium citrate 10 gramme. Sodium chloride 0.75 grammes. In distilled water 100 c.c.
Sterilise by boiling.
2. Prepare the animal as for intravenous inoculation (see page 363) and introduce the syringe needle into the lumen of the selected vein.
3. Slowly withdraw the piston of the syringe. When sufficient blood has been collected direct the assistant to release the proximal compression of the vein; and withdraw the needle.
4. Remove the needle from the nozzle of the syringe and deliver the citrated blood into a small Ehlenmeyer flask containing about 250 c.c. of nutrient broth.
5. Label, incubate and examine daily until growth occurs or until the expiration of ten days.
c. The serological examination of the blood is directed to the demonstration of the presence of certain specific antibodies in the sera of experimentally infected animals, and within certain limits to an estimation of their amounts.
The chief of these bodies are:
Antitoxin. Agglutinin. Precipitin. Opsonin. Immune body or Bacteriolysin.
None of these substances are capable of isolation in a state of purity apart from the blood serum, consequently special methods have been elaborated to permit of their recognition. In every instance the behaviour of serum from the experimental animal, which may be termed "specific" serum, is studied in comparison with that of serum from an uninoculated animal of the same species, and which is termed "normal" serum. In view of minor differences in constitution exhibited by the serum of various individuals of the same series, it is usual to employ a mixture of sera obtained from several different normal animals of the same species as the inoculated animal, under the term "pooled serum." The method of collecting blood (e. g., from the rabbit) for serological tests is as follows:
Collection of Serum.
Apparatus required:
Razor. Liquid soap. Cotton-wool. Lysol 2 per cent. solution, in drop bottle. Ether in drop bottle. Flat Hagedorn needles. Blood pipettes (Fig. 16, page 12). Centrifugal machine. Centrifuge tubes. Glass cutting knife. Bunsen flame. Writing diamond or grease pencil.
METHOD.
1. Shave the dorsal surface of the ear over the course of the posterior auricular vein (see Fig. 192).
2. Sterilise the skin by washing with lysol.
The lysol should be applied with sterile cotton-wool and the ear vigourously rubbed, not only to remove superficial scales of epithelium, but also to render the ear hyperaemic and the vein prominent.
3. Remove the lysol with ether dropped from a drop bottle, and allow the ether to evaporate.
4. Puncture the vein with a sterile Hagedorn needle.
5. Take a small blood-collecting pipette (Fig. 161) and hold it at an angle to the ear, one end touching the issuing drop of blood, the other depressed.
The blood will now enter the pipette at first by capillarity; afterward gravity will also come into play and the pipette can be two-thirds filled without difficulty.
6. Hold the tube by the end containing the blood, the clean end pointing obliquely upward—warm this end at the bunsen flame to expel some of the contained air; then seal the clean point in the flame.
7. Place the pipette down on a cool surface (e. g., a glass slide). The rapid cooling of the air in the clean end of the pipette creates a negative pressure, and the blood is sucked back into the pipette, leaving the soiled end free from blood. Seal this end in the bunsen flame.
8. Mark the distinctive title of the specimen (e. g., animal's number) upon the pipette with a writing diamond or grease pencil.
9. When the sealed ends are cold and the blood has clotted, place the pipette on the centrifuge, clean end downward; counterpoise and centrifugalise thoroughly. On removing the pipette from the centrifuge, the red cells will be collected in a firm mass at one end, and above them will appear the clear serum.
10. By marking the blood pipette above the level of the serum with the glass cutting knife and snapping the tube at that point, the blood-serum becomes readily accessible for testing purposes.
If larger quantities of blood are required, the animal, after puncturing the vein, should be inverted, an assistant holding it up by the legs. Blood to the volume of several cubic centimetres will now drop from the punctured vein, and should be caught in a tapering centrifuge tube, the tube transferred to the incubator at 37 deg. C. for two hours, then placed in the centrifugal machine, counterpoised and centrifugalised thoroughly. The three most important of the antibodies referred to which can be demonstrated with a certain amount of facility are agglutinin, opsonin and bacteriolysin; and the methods of testing for these bodies will now be considered.
AGGLUTININ.
Agglutinin is the name given to a substance present in the blood-serum of an animal that has successfully resisted inoculation with a certain micro-organism. This substance possesses the power of collecting together in clumps and masses, or agglutinating watery suspensions of that particular microbe.
Dilution of the Specific Serum:
Apparatus required:
Sterile graduated capillary pipettes to contain 10 c. mm. (Fig. 17). Sterile graduated capillary pipettes to contain 90 c. mm. (Fig. 17). Small sterile test-tubes 5 x 0.5 cm. Normal saline solution in flask or test-tube. Pipette of specific serum. Glass cutting knife, or three-square file. Glass capsule, nearly full of dry silver sand, or roll of plasticine. Grease pencil.
METHOD.—
1. Take three sterile test-tubes and number them 1, 2 and 3.
2. Pipette 0.9 c.c. sterile normal saline solution into each tube, and stand tubes upright in the sand in the capsule, or in the plasticine block.
3. Make a scratch with the glass cutting knife on the blood pipette above the upper level of the clear serum, and snap off and discard the empty portion of the tube.
4. Remove 0.1 c.c. of the serum from the blood pipette tube, and mix it thoroughly with the fluid in tube No. 1; and label s.s., (specific serum), 10 per cent.
5. Remove 0.1 c.c. of the solution from tube No. 1 by means of a fresh pipette, and mix it with the contents of tube No. 2; and label s.s., 1 per cent.
6. Remove 0.1 c.c. of the solution from tube No. 2 by means of a fresh pipette, and mix it with the contents of tube No. 3; and label s.s., 0.1 per cent.
When the yield of serum from the specimen of blood which has been collected, or is available, is small, the above method of diluting is not practicable, and the dilution should be carried out by Wright's method in a capillary teat pipette.
Dilution of Serum by Means of a Teat Pipette.
Materials required:
Blood pipette containing sample of specific serum after centrifugalisation. Capsule of diluting fluid—normal saline solution. Supply of Pasteur pipettes (Fig. 13a). India-rubber teats. Small test-tubes. A block of plasticine to act as a test-tube stand. Grease pencil.
METHOD:
1. Mark three small test-tubes 10 per cent., 1 per cent. and 0.1 per cent. respectively, and stand them upright in the plasticine block.
2. Take a Pasteur pipette, nick the capillary stem just above the sealed end with a glass cutting knife, and snap off the sealed end with a quick movement so that the fracture is clean cut and at right angles to the long axis of the capillary stem—cut "square", in fact. Prepare several, say a dozen, in this manner.
3. Fit a rubber teat to the barrel of each of the pipettes.
4. Make a mark with the grease pencil on the stem of one of the pipettes about 2 or 3 cm. from the open extremity.
5. Compress the teat between the finger and thumb (Fig. 193) to such an extent as to drive out the greater part of the contained air.
6. Maintaining the pressure on the teat pass the stem of the pipette into the capsule holding the saline solution, until the open end of the pipette is below the level of the fluid.
7. Now cautiously relax the pressure on the teat and let the fluid enter the pipette and rise in the stem until it reaches the level of the grease pencil mark. As soon as this point is reached, check the movement of the column of fluid by maintaining the pressure on the teat, neither relaxing nor increasing it.
8. Withdraw the point of the pipette clear of the fluid, and again relax the pressure on the teat very slightly. The column of saline solution rises higher in the stem, and a column of air will now enter the pipette and serve as an index to separate the first volume of fluid drawn into the stem from the next succeeding one.
9. Again introduce the end of the pipette into the fluid and draw up a second volume of saline to the level of the grease pencil mark, and follow this with a second air index.
10. In like manner take up seven more equal volumes of saline solution and their following air bubbles. There are now nine equal volumes of normal saline in the pipette.
11. Now pass the point of the pipette into the blood tube and dip the open end below the surface of the serum. Proceeding as before, aspirate a volume of serum into the capillary stem up to the level of the pencil mark.
12. Eject the contents of the pipette into the small tube marked 10 per cent. by compressing the rubber teat between thumb and finger.
13. Mix the one volume of serum with the nine volumes of saline solution very thoroughly by repeatedly drawing up the whole of the fluid into the pipette and driving it out again into the test-tube.
14. Now take a clean pipette and proceed precisely as before, 4 to 10.
15. Having aspirated nine equal volumes of saline into this second pipette, now take up one similar volume of the fluid in the "10 per cent. tube."
16. Eject the contents of this pipette into the second tube marked 1 per cent. and mix thoroughly as before.
17. In similar fashion make the 0.1 per cent. solution and transfer to the third tube.
18. Further dilutions in multiples of ten can be prepared in the same way, and by varying the number of volumes of diluting fluid or serum any required dilution can be made (see Appendix, Dilution Tables).
NOTE.—The saline diluting fluid must always be taken into the pipette first, otherwise if the serum contains a very large amount of agglutinin the traces of this serum added to the saline solution may be sufficient to entirely vitiate the subsequent observations—whilst if more than one sample of serum is diluted from the same saline solution serious errors may be introduced into the experiments.
The Microscopical Reaction:
Apparatus Required:
Five hanging-drop slides (or preferably two slide), with two cells mounted side by side on each (Fig. 62, a), and one slide with one cell only.
Vaseline.
Cover-slips.
Platinum loop.
Grease pencil.
Eighteen to twenty-four-hour-old bouillon cultivation of the organism to be tested (e. g., Bacillus typhi abdominalis)
Pipette end with the remainder of the specific serum labelled s.s.
Tubes containing the three solutions of the specific serum, 10, 1, and 0.1 per cent. respectively.
Pipette end with pooled normal serum labelled p.s.
METHOD.—
1. Make five hanging-drop preparations, thus:
(a) One loopful of bouillon cultivation + one loopful pooled serum; label "Control."
(b) One loopful culture + one loopful undiluted specific serum; label 50 per cent.
Mount these two cover-slips on a double-celled slide.
(c) One loopful bouillon culture + one loopful 10 per cent. serum; label 5 per cent.
Mount this on single-cell slide.
(d) One loopful bouillon culture + one loopful 1 per cent. serum; label 0.5 per cent.
(e) One loopful bouillon culture + one loopful 0.1 per cent. serum; label 0.05 per cent.
Mount these two cover-slips on a double-celled slide.
2. Note the time: Examine the control to determine that the bacilli are motile and uniformly scattered over the field—not collected into masses.
3. Next examine the 50 per cent. serum preparation.
If agglutinin is present and the test is giving a positive reaction, the bacilli will be collected in large clumps.
If the test is giving a negative reaction, the bacilli may be collected in large clumps owing to the viscosity of the concentrated serum.
4. Observe the 5 per cent. preparation microscopically.
If the bacilli are aggregated into clumps, positive reaction.
If the bacilli are not aggregated into clumps, observe until thirty minutes from the time of preparation before recording a negative reaction.
5. Examine the 0.5 and 0.05 per cent. preparations.
These may or may not show agglutination when the result of the examination of the 5 per cent. preparation is positive, according to the potency of the specific serum; and by the examination of a series of dilutions a quantitative comparison of the valency of specific sera from different sources, or of serum from the same animal at different periods during the course of active immunisation may be obtained.
NOTE.—The graduated pipettes supplied with Thoma's haematocytometer (intended for the collection of the specimen of blood required for the enumeration of leucocytes), giving a dilution of 1 in 10—i. e., 10 per cent.—may be substituted for the graduated capillary pipettes referred to above, if the vessel in which the serum has been separated is of sufficiently large diameter to permit of their use.
The Macroscopical Reaction:
Sterile graduated capillary pipettes to contain 90 c. mm.
Eighteen to twenty-four-hours-old bouillon cultivation of the organism to be tested.
Three test-tubes containing the 10, 1, and 0.1 per cent. solutions of specific serum (about 90 c. mm. remaining in each).
Tube containing 50 per cent. solution of pooled serum.
Sedimentation pipettes (vide page 17) or teat pipettes.
METHOD.
1. Pipette 90 c. mm. of the bouillon culture into each of the tubes containing the diluted serum; and the same quantity into the tube containing the pooled serum.
2. Fill a sedimentation tube (by aspirating) or a teat pipette from the contents of each tube. Seal off the lower ends of the sedimentation tubes in the Bunsen flame.
3. Label each tube with the dilution of serum that it contains—viz., 5, 0.5, and 0.05 per cent.
4. Place the pipettes in a vertical position, in a beaker, in the incubator at 37 deg. C., for one or two hours.
5. Observe the granular precipitate which is thrown down when the reaction is positive, and the uniform turbidity of the negative reaction as compared with the appearances in the control pooled serum.
OPSONIN.
Opsonin is the term applied by Wright to a substance, present in the serum of an inoculated animal, which is able to act upon or sensitise bacteria of the species originally injected, so as to render them an easy prey to the phagocytic activity of polymorphonuclear leucocytes. In the method for demonstrating opsonin about to be described, a comparison is made between the opsonic "power" of the pooled serum and the specific serum.
Apparatus:
Small centrifuge and tubes for same (made from the barrels of broken capillary pipettes by sealing the conical ends in the bunsen flame).
Capillary Pasteur pipettes.
India-rubber teats.
Grease pencil.
Bunsen burner with peep flame.
Electrical signal clock (see page 39) stop watch, or watch.
Rectangular glass box or tray to hold pipettes.
Incubator regulated at 37 deg. C.
3 x 1 slides.
Piece of light rubber tubing.
Rectangular block of plasticine.
Flask of normal saline solution.
Flask of sodium citrate (1.5 per cent.) in normal saline solution.
Materials required, and their preparation:
Small tube of "washed cells" (red blood discs and leucocytes); human cells are used in estimating the opsonising power of the serum of experimental animals.
Small tube of emulsion of bacteria of the species responsible for the infection of the experimental animal.
Blood pipette containing specific serum.
Blood pipette containing "pooled" serum.
Washed Cells.—
1. Take a small centrifuge tube and half fill it with sodium citrate solution. Mark with the grease pencil the upper limit of the fluid.
2. Cleanse the skin of the distal phalanx of the second finger of the left hand above the root of the nail with lint and ether. Wind the rubber tubing tightly round the second phalanx; puncture with a sterile Hagedorn needle through the cleansed area of skin.
3. Take up a sufficiency of the issuing blood (more or less according to the number of tests to be performed) with a teat pipette, transfer it to the tube of citrate solution and mix thoroughly. Make a second mark on the tube at the upper level of the mixed citrate solution and blood.
4. Place the tube in the centrifuge, counterpoise accurately and centrifugalise until the blood cells are thrown down in a compact mass occupying approximately the same volume as is included between the two pencil marks.
The column of fluid in the tube now shows clear supernatant fluid (citrate solution and blood plasma) separated from the sharp cut upper surface of the red deposit of corpuscles by a narrow greyish layer of leucocytes.
5. Remove the supernatant column of citrate solution by means of a teat pipette, fill normal saline solution into the tube up to the upper pencil mark, and distribute the blood cells throughout the saline by means of the teat pipette. Centrifugalise as before.
6. Again remove the supernatant fluid and fill in a fresh supply of saline solution and centrifugalise once more.
7. Remove the supernatant saline solution as nearly down to the level of the leucocytes as can be safely done without removing any of the leucocytes.
8. Next distribute the leucocytes evenly throughout the mass of red cells by rotating the tube between the palms of the hands—just as is done with a tube of liquefied medium prior to pouring a plate.
9. Set the tube upright in the plasticine block near to one end.
Bacterial Emulsion.—
1. Take an 18- to 24-hour culture of the required bacterium (e. g., Diplococcus pneumoniae) grown upon sloped blood agar at 37 deg. C. Pour over the surface of the medium some 5 c.c. of normal saline solution.
2. With a platinum loop emulsify the growth from the surface of the medium as evenly as possible in the saline solution.
3. Allow the tube to stand for a few minutes so that the large masses of growth may settle down; transfer the upper portion of the saline suspension to a centrifuge tube and centrifugalise thoroughly.
4. Examine a drop of the supernatant opalescent emulsion microscopically to determine its freedom from clumps and masses. If unsatisfactory prepare another emulsion, this time scraping up the surface growth with a platinum spatula, transferring it to an agate mortar and grinding it up with successive small quantities of normal saline. If satisfactory insert the tube in the plasticine block next to that containing the washed cells.
Specific Serum.—
Pooled Serum.—
These sera are collected and treated as already described (see page 379), and the portions of the blood pipettes containing them are arranged in the remaining space in plasticine block.
The plasticine block now presents the appearances shown in Fig. 194.
METHOD FOR DETERMINING THE OPSONIC INDEX.—
1. Take a capillary pipette fitted with a teat, cut the distal end square and make a pencil mark about 2 cm. from the end.
2. Aspirate into the pipette one volume of washed cells, air index, one volume of bacterial emulsion, air index, and one volume of specific serum (see Fig. 195).
3. Mix thoroughly on a 3 by 1 slide by compressing the teat and ejecting the contents of the pipette on to the surface of the slide, relaxing the pressure and so drawing the fluid up into the pipette again. These two processes should be repeated several times; finally take up the mixture in an unbroken column to the central portion of the capillary stem.
4. Seal the point of the pipette in the peep flame of the bunsen burner and remove teat.
5. Mark the pipette (with the grease pencil) with the distinctive number of the serum and place it in the glass box or tray.
6. Take another similarly prepared pipette and aspirate into it equal volumes of washed cells, bacterial emulsion and pooled serum. Treat precisely as in 3 and 4, label it "control" or "N.S." (normal serum) and place in the box by the side of the specific serum preparation.
7. Place the box with the pipettes in the incubator and set the signal clock to ring at 15 minutes (or start the stop watch).
8. At the expiration of the incubation time remove the pipettes from the incubator.
9. Cut off the sealed end of the specific serum preparation. Mix its contents thoroughly as in step 3, and then divide the mixture between two 3 by 1 slips and carefully spread a blood film (vide page 376) on each in such a way that only one-half of the surface of each slide is covered with blood—the free edge of the blood film approximating to the longitudinal axis of the slide.
Allow films to dry and label the slides with writing diamond.
10. Treat the contents of the control pipette in similar fashion.
11. Select the better film from each pair for fixing and staining.
12. Fixing and staining must be carried out under strictly comparable conditions, and to this end the slides are best handled by placing in a glass staining rack which can be lowered in turn into each of a series of glass troughs containing the various reagents (Fig. 196). Place the rack in the first trough which contains the alcoholic solution of Leishman's stain for two minutes to fix. |
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