In this article we will discuss about the Laboratory Animals:- 1. Care and Management of Laboratory Animals 2. Data’s of Laboratory Animals 3. Diseases.
Care and Management of Laboratory Animals:
Laboratory animals in most countries are protected by the Law “Cruelty to Animals Act”. One is to obtain licence from the Home Department for using them on experimental purposes.
The health and well-being of the animals depend on the care, human attitude of the animal-keepers (staff) of the animal house. To keep animals healthy, the staff has to look after cleanliness of the animal rooms and cages, provide proper food and water, to move to a cooler or warmer place and to provide air.
The general principles are outlined:
1. Fluid:
They are to be provided with plentiful supply of fresh clean drinking water from a bottle (250 ml capacity) attached to the outside of the cage. The water is led in a 6-9 mm glass tubing through a rubber bung to an accessible position inside the cage; the outlet tubing is about 3 mm.
2. Diet:
A balanced diet containing carbohydrate, fat, protein, vitamins and trace elements is to be given regularly. It is commercially in the form of cubes or pellets. Small quantities of green stuff are also to be supplied.
3. Cleanliness:
Cleanliness of room and cages is essential unless they are kept in considerable risk of epidemic diseases. Animals, when breeding, should not have their cages changed too often.
Clean cages should be used. Cages may be boiled in soapy water; alternatively, to be kept immersed in a solution of disinfectant such as 3% Lysol. Lysol, however, should not be used in cleaning the cages of rabbit because its smell distresses the animals.
4. Litter:
A layer of absorbent material (e.g. soft wood sawdust’s, sugarcane piths) should be spread to a depth of 1/2 to 1 inch (1.25-2.5 cm) on the bottom of the cages.
5. Cages:
Each species of animal require its own type of cage. It should be large enough for movement and some exercise of the animal.
6. Labelling:
Every case should be provided with a holder or socket for a small card of 6-9 cm for record of the experiment (date, identifying marks of animal, nature of experiment and specimen).
7. Ventilation:
Animal room should be air-conditioned; or at least ten changes of air in each hour are needed.
8. Humidity:
Humidity of animal house ranges between 45% for rabbits to 65% for mice.
9. Marking animals:
White or light coloured animals are marked by staining the fur with a strong dye (e.g. carbol fuchsin, eosin). Rabbits may be marked in ears with a needle dipped in India ink. Rats and mice are punctured in ear and fowls are marked by numbered metal tags on legs clipped through the loose skin of the wing.
10. Detection of disease in animals:
A routine tour of inspection of the animals should be made at least once a day with attention to the general condition of animals, amounts of food and water consumed and the nature of faeces. To see nose movements of the animals and to see any animal remaining quiet and still. Such animals may be separated and investigated for the cause of disease.
11. Recording animals’ temperature:
Clinical thermometer is liberally smeared with sterile petroleum jelly and the blunt-ended rectal thermometer is introduced into the rectum or vagina to a depth of about 3 to 3.5 cm.
12. Prevention of disease:
(i) Newly arrived animals to be kept in a special quarantine room and kept under observation for 10-14 days. Animals falling sick during the period should be kept in quarantine and necropsis must be done for finding out the cause of the illness.
(ii) Animal infected experimentally with bacteria or viruses should be kept in separate isolation rooms to prevent spread of infection to other animals.
13. Insect pest:
Bed bugs, fleas, lice, mites, ticks, flies, mosquitoes and cockroaches may all infest the animal house. These can be controlled by 0.5% insecticidal sprays or 10% DDT.
Data’s of Laboratory Animals:
Table 18.1 shows important data’s on some of the animals.
Rabbit (Fig. 18.1):
Handling:
Animals should be handled with care. The rabbit is picked up from cage with the ears by one hand in a firm grip and another hand is placed under the hind-quarters to support the weight and then lifted gently. After removing from cage, the animal is placed in a non-slippery place as it otherwise feels insecure and becomes frightened.
Breeding:
When the doe (female) is on heat, vulva and vagina become red, swollen and moist. The doe is placed in the cage of buck (male).
Guinea Pig (Fig. 18.2):
Handling:
One hand is placed across the back of the animal with the thumb behind the shoulders and the other fingers well forward on the opposite side. The animal is lifted gently supporting its weight with other hand placing the palm uppermost under the hind quarters.
Armadillo (Fig. 18.3):
When nine-banded armadillo is inoculated with lepra bacilli, generalized infection develops with extensive multiplication of the bacilli and the lesions produced resemble lepromatous leprosy.
Natural infection by a mycobacterium resembling lepra bacillus has been observed in some wild caught armadillos in Texas and Mexico. The wild armadillo, therefore, should be screened for mycobacterial infection and kept in quarantine for 3 months before inoculation. European hedgehog bred in captivity is suitable for propagation.
Mice:
Handling:
Take the help of an assistant who holds the tail of the animal with the left hand and gently raises the hind limbs from floor of the cage. The mouse placed in such position cannot turn round and bite. By the help of right finger and thumb a fold of skin close to the head is held.
Rat (Fig. 18.4):
Rat is handled in the same way as guinea pig by experienced operator.
Diseases of Laboratory Animals:
Diarrhoea due to Salmonella typhimurium is a common natural disease of rabbits, guinea pigs and mice. Coccidiosis (Protozoan disease) and pseudo tuberculosis (caused by Yersinia pseudo tuberculosis) are also common natural disease of animals. Explosive epizootics have been observed with S. typhimurium infection in which practically the whole of a colony of animals have been destroyed.
The natural diseases of the laboratory animals are given:
Rabbits:
Coccidiosis (diarrhoea), pseudo tuberculosis (caseous lymph nodes) and Taenia pisiformis (numerous cysts in omentum).
Guinea pigs:
Protozoan diseases (coccidiosis toxoplasmosis), pseudo tuberculosis, S. typhimurium, abscess in lymph node (Streptococcus group C), haemorrhagic septicaemia (P. multocida) and viral pneumonia and paralysis.
Mice:
Mouse typhoid (caused by S. typhimurium, S. enteritidis) may be responsible for severe epizootics. Other diseases include: Ectromelia (mouse pox, caused by a pox virus related to the vaccinia virus), Streptobacillus moniliformis, infection, viral infections and Taenia taeniaeformis infection (large cysts in liver).
Hamsters:
Streptococcal and staphylococcal infections, distemper (virus) and foot rot (caused by mites).
Fowls (Fig. 18.5):
Salmonellae of various types (e.g. S. typhimurium causing diarrhoea), S. pullorum (outbreak of typhoid), coccidiosis, avian leucosis (involving lymph node and bone marrow), parasitic infections due to lice and mite, virus diseases (infectious laryngotracheitis, fowl pest due to Newcastle virus).
Materials Inoculated:
1. Urine, CSF, blood and serous fluids:
These specimens are inoculated with a medium- bore needle but in case of tenacious material, like pus and sputum, is injected through a wide-bore needle.
2. Culture materials:
Liquid cultures are inoculated through a medium- bore needle. Growths on solid media are first scraped off and suspended in broth or saline, alternatively, the diluting fluid may be poured on the culture which is then emulsified with a wire loop.
3. Tissues:
Small fragments of tissues such as brain, spleen, liver and kidney are first homogenized by crushing these materials with a suitable diluent in a tissue grinder. When tissue is well-ground, more saline is added and allowed to stand for a short time.
After the tissue has settled to the bottom of the mortar, the supernatant fluid is drawn into a syringe. When intravenous injection is to be given, care must be taken that no large particles are injected. The suspension must be centrifuged at a low speed and the supernatant fluid is used for intravenous administration.
Experimental Procedure Inoculation of Material:
1. Scarification:
Procedure:
At first the animal is clipped and its flank is shaved to remove the hair. Alternatively, depilating powder (mixture) may be used to remove hair. Skin is cleansed with alcohol and allowed to evaporate. Several parallel scratches are made by a sharp sterile scalpel, just enough to draw blood. The specimen is then rubbed into the scarified area with the side of the scalpel.
Use:
This technique is mainly employed for the propagation of Vaccinia virus in rabbit.
2. Subcutaneous inoculation:
Procedure:
Subcutaneous inoculation is made in the loose tissue about the flank or into the abdominal wall. Hair is clipped. The skin is sterilised with iodine and alcohol and then pinched up, and the needle is inserted. Amounts up to 5 ml can be introduced. A 2 ml or 5 ml syringe is conveniently used.
Use:
This method is the same for rabbit and guinea pig and obviates superficial ulceration when tuberculous specimen is injected. In fowls, subcutaneous inoculation is done in pectoral or thigh regions.
3. Intra-cutaneous inoculation:
Procedure:
Hair is removed from the flanks of the animal either by plucking or shaving or by depilating powder. A 1 ml all-glass tuberculin syringe, fitted with short needle No. 25 or 26 gauge is used. After pinching up the skin of the animal between the thumb and forefinger, the point of the needle is inserted at the top of the fold so that the level of the needle is towards the surface of the skin.
The needle passes into the dermis only, as near as the surface. Usually, a volume of 0.2 ml is injected. When several tests are to be done, injections are to be given about one inch apart and not too near the middle line of the abdomen. No more than ten injections should be made in one animal.
Use:
This method is mainly used in testing cultures of the diphtheria bacillus for toxigenicity. Usually white guinea pigs (300-400 g) are used.
4. Intra-peritoneal inoculation:
Procedure:
Animal is held by assistant as above and the material is inoculated in the peritoneum in the midline in the lower half of the abdomen. Not more than 5 ml can be inoculated intraperitoneally.
Use:
It is done mostly in guinea pig and sometimes mice are also intraperitoneally inoculated.
5. Intravenous inoculation:
By this method, specimen is directly introduced into the circulation.
Procedure:
Intravenous inoculation is done in rabbit, guinea pig, mouse and rat as follows:
(a) Rabbit:
For this, the marginal vein of the ear is most convenient. The hair over the ear is removed by dry shaving with a sharp razor. Vein may be distended by rubbing with a piece of cotton wool or by holding the ear on an electric bulb (heat dilates blood vessels).
The rabbit is held by an assistant or placed in a special box so that only its head protrudes. The operator faces the animal and holds the ear horizontally by left hand and injects the material into the vein by a sterile syringe.
Uses:
Rabbits are mainly used for diagnostic purposes. They are also used for production of immune sera used in diagnostic purpose.
(b) Mouse and rat:
Intravenous injection may be given into a vein at the root of the tail by a fine needle. The tail vein is dilated by placing in water at about 45°C. A maximum volume of 0.5 ml can thus be inoculated.
(c) Fowl:
Fowl is placed on its side and feathers over an area in the elbow are plucked and a large brachial vein is seen on the bone beneath the skin. Intravenous administration of specimen or culture is made carefully with fine needle.
6. Intra-cerebral inoculation:
(a) Rabbit:
The animal is anaesthetized with ether, hair over the head is shaved. After disinfection of skin of head, a short incision is made by a scalpel through the scalp at a point 2 mm lateral to the sagittal suture and 1.5 mm anterior to the lambdoidal suture.
The skull is perforated with a trephine or mechanical drill. A 0.45 ml of material is introduced into the occipital lobe of brain by a needle (cut down to 5/18 inch). Rabbits may also be inoculated in frontal lobe of brain, at a point 2 mm lateral to the median plane joining the two external canthi of the eyes.
(b) Mouse and rat:
The area of skull is disinfected as above. A 0.03 ml of fluid is injected directly by a one ml syringe at a site midway between the outer canthus of the eye and the point of attachment of the pinna of the ear about 3 mm from the midline. About 1/8 to 1/6 inch point of needle is introduced through skull.
7. Intra-testicular:
Rabbit:
Testes lie in abdomen. The animal lies on the back and the testes are made to descend on scrotum by steady pressure on the belly. After fixing the testes by an assistant, skin is disinfected and then 0.2 to 0.4 ml of inoculum is directly introduced into the centre of a testis by a hypodermic needle.
8. Ophthalmic:
Rabbit:
Only one eye is inoculated by dropping a drop of material through a Pasteur pipette. Before installation of the material, one eye may be scarified by anesthesia.
9. Intranasal inoculation:
(a) Mouse:
It should preferably be kept in an inoculating chamber specially designed for the purpose so that the operator is not at a risk of inhaling the infective material. Failing this, the operator should use a mask. The animal is anaesthetized with ether and as soon as its breathing has become deep, a volume of 0.1 ml is introduced into the anterior nares one side only by an automatic pipette.
(b) Ferrets and fowls:
Under light ether anesthesia, as soon as the regular respiration is established, 0.5 to 1.5 ml material is dropped from a Pasteur pipette into the nares.
Use:
For studies of Distemper virus in ferret and New Castle virus and other virus infections in fowls.
Inoculation of infant mice:
Suckling mice of 48 hours old or less are employed for the isolation of herpes simplex, enteric and other arbo-viruses. The litters should be handled with great care and cleanliness to avoid biting from their mothers.
The following dose of injections can be used:
(i) 0.03 ml subcutaneously
(ii) 0.05 ml intraperitoneally
(iii) 0.03 ml intra-cerebrally.
Sometimes two routes — intra-peritoneal and intra-cerebral — inoculations are done together in the same animal.
Experimental Procedure:
Collection of material:
1. Ear Vein:
About 20-30 ml blood may be collected from the ear vein of a large rabbit. The ear is shaved and sterilised with sterile gauze soaked in 70% alcohol. Before operation, heated (by Bunsen burner) petroleum jelly is kept ready in a container. When the jelly becomes cool but still fluid, the jelly is painted over the vein — both on margin and underside of the ear.
The ear is held forward and the vein is made prominent by means of a small spring clip at the base of the ear. The vein is incised with a small sharp sterile scalpel. The blood is allowed to drop into a sterile flask containing glass beads. The ear vein can be dilated by holding on an electric bulb under it or by rubbing with a piece of cotton soaked xylol.
Use:
This technique is mainly used in rabbit and sometimes in guinea pig (for collection up to 0.5 ml blood).
2. Cardiac puncture:
Larger volumes of blood can be collected by cardiac puncture:
(a) Guinea pig:
After lightly anaesthetizing with ether, the animal laid on its back with its front limbs drawn forwards. An area over the fourth and fifth left intercostal spaces is plucked and the skin painted with tincture of iodine. The location of the apex beat of the heart is defined by digital palpation and thereafter, a needle (No. 20 gauge mounted on a syringe) is inserted at this point between the ribs.
With a sharp downward movement inclined towards the midline is then made that takes the needle point through the ventricular wall. As much as 15 ml of blood from the heart of guinea pig may be obtained from a 350-400 g animal, though smaller amounts are advisable if the guinea pig is to survive. For the success of this operation very sharp needles are essential.
(b) Rabbit:
The animal is anaesthetized and then fastened to a board with the body axis quite straight and the fur clipped over the left side of the chest; the area is shaved and then sterilised with alcohol and ether.
A 100 ml bulb pipette (vide Fig. 18.6) that has been cut down at both ends to 9 in length, one end being slightly tapered and the other end stoppered with cotton-wool. It is wrapped in craft paper and sterilised in the hot-air oven.
A wide-bore transfusion needle is fitted into a short length (1 ½ in) of thick rubber tubing and sterilised by boiling. When the animal is anaesthetized, the rubber tubing is attached to the tapered end of the pipette and to the other end is fitted a mouthpiece such as that used in pipetting.
After inserting the needle into the left side of the chest, suction applied. The needle should lie in the right ventricle of the heart, and blood rapidly flows into the pipette. Volume of 50 ml of blood per kg of body-weight can be obtained. Thereafter, the blood is transferred to a sterile 500 ml flask or bottle containing glass beads for defibrination.
(c) Fowls:
Method (1):
The fowl is anaesthetized and placed on its right side. The needle is inserted over the heart of the bird between the second and third ribs at a point close to the edge of the breast muscle which can be felt. At a depth of 1 ½ inch (3.75 cm) the needle enters the ventricle. This technique carries a risk of lung puncture and required experience before it can be used with confidence.
Method (2) is perhaps easier. The bird is placed on its back, ventral side uppermost, and stretch its neck over the edge of the table. Pluck the base of the neck and the skin is cleaned with alcohol as before. A 20 ml syringe with a No. 17 or 18 gauge needle 2 in long is inserted horizontally just deep to the sternum; it should be tilted very slightly downwards and will enter the heart at a depth of 1 ½ inch (3.75 cm).
Necropsy:
Post-mortem examination should be done routinely for all experimental animals, whatever may be the cause of death. When a virulent pathogen such as the bacillus of plague or of anthrax has been injected into animals, special care must be taken to avoid dissemination of infection, with danger to the operator and other workers.
The procedure in conducting a necropsy and also the method used when dealing with highly infectious organisms should be mentioned. Since the principal application of necropsy is to recover organisms previously injected into the animal, hence, the post-mortem examination must be conducted with strict aseptic precautions.
Materials required:
A suitable animal board or table, on the top of which the carcase can be fixed in the supine position should be used.
Instruments:
Instruments include three scalpels; scissors, ordinary size, four pairs; mouse-toothed forceps, four pairs; small bone forceps, if the skull is to be opened; a searing iron—a 4-oz soldering bolt is suitable for the purpose; sterile glass-wares like capillary pipettes; Petri dishes, test tubes and tubes, bottles or plates of media.
Knives are properly sterilised in strong lysol (about 20%) and then placed in a weaker solution (2%). Metal instruments are sterilised by boiling in a sterilising bath, e.g. an enameled “fish-kettle”. The tray of sterile instruments is lifted out of the sterilizer and laid on a spread towel which has previously been soaked in 1: 1,000 solution of mercuric chloride.
Where cultures are to be made, first to immerse the animal completely in weak Lysol solution (3%) for a few moments.
This technique not only destroys most of the surface organisms, but prevents the dust in the fur from getting into the air and thereby contaminating other materials. Animal is then fixed to the board and towels moistened with antiseptic solution are placed over the head and lower extremities of the animal.
Instruments are removed from the sterilizer. A long median incision is now made through the skin of the abdomen and chest and then the skin is widely dissected, exposing the abdominal and chest muscles. The peritoneal cavity is opened with second set of instruments and the abdominal wall is reflected to each side.
The spleen is removed with fresh instruments and placed in a sterile Petri dish. Other organs like the liver, kidneys and renals may be similarly removed.
The ensiform cartilage is now tightly gripped with the help of a pair of strong forceps, and by means of a sterile pair of strong scissors a cut is made on either side of the chest through the costal cartilages. Sternum is raised and pulled towards the head. The heart is now exposed.
A sterile capillary pipette, furnished with a teat, or a sterile syringe is passed through the heart wall. Blood can thus be withdrawn and inoculated into various media. When the necropsy has been properly performed, it is not necessary to scar the surface of the heart.
The lungs are then removed with fresh instruments by cutting each organ free at the hilum. Care must be taken not to open into the oesophagus if the lungs are to be used for cultivation.
After the removal of the organs these are placed in separate Petri dishes. Naked-eye appearances of the organs are studied. For culture the spleen gives the best results, but the other solid viscera may also similarly be used.
The organ is cut open with sterile instruments and a small portion which is taken up with a stiff wire and smeared on the surface of solid media. The liquid media are inoculated with a small fragment of the tissue.
While conducting post-mortem examinations, one should be familiar with various animal diseases, such as worm infestation, coccidiosis, pseudo tuberculosis, etc. which may be noticed, and the worker should have clear idea of their appearances.
When the animal is inoculated with highly pathogenic organisms the worker should always wear rubber gloves. After soaking the carcase in antiseptic solution as before it is nailed to a rough piece of board of the appropriate size.
This board is then placed in a large enameled iron tray. Necropsy should be performed carefully in the usual way. Carcase is finally covered with 10% Lysol, that flows over the board and into the tray.
The entire contents of the tray-board and carcase are then destroyed in an incinerator or furnace. Rubber gloves, instruments and tray are thoroughly sterilised. The wearing of a large overall made of waterproof material is strongly recommended in all PM examination and, in addition, the use of some form of glasses or goggles to protect the eyes is to be done.