(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.
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).
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).
8. Inject the fluid from the syringe.
9. Label, etc.
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.
(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.
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 manœuvres 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.)
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.
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.—(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.
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.
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 œsophageal 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 œsophagus 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 œsophageal 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.
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° 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 Temp. °C. | Pulse. | Respirations. | |
| Animal. | 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 hæmatologists, 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 HÆMOCYTOLOGY OF LABORATORY ANIMALS.
| Totals | Percentages | |||||||
| Animal | Red cells | White cells | Hb, per cent. | Lymphocytes, per cent. | Large monos, per cent. | Polymorph, per cent. | Eosinoph, per cent. | Mast cells, per 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) Normal limits. | 5,000,000 (4.5-5) millions. | 7,500 (7-9) thousands. | 100 (95-101) | 25 (20-30) | 5.5 (4-8) | 65 (55-68) | 4.0 (3-5) | 0.5 (0.5-2) |
The above table represents in each case the average of a large number of counts.
Remarks.
Frog.—The red cells are large oval nucleated (20-25µ by 12-15µ) discs, the nucleus relatively small and irregularly elongated or oval, about 10µ in length. Many primitive and developing forms are usually observed—also free nuclei and many cells in various stages of degeneration. Hæmoglobin estimation is difficult owing to turbidity of the blood after dilution with water. The polymorphonuclear leucocytes are large cells, about 20µ; 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 4.5µ 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 20µ, 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 12µ by 6µ, the nucleus being relatively small (about 4µ 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°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°, 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: