Lesson 3a: Cultivation of viruses in animals

• Animal inoculation was the first method of virus cultivation and for many years was the only means of virus propagation.
• To confirm the infectious nature of the disease it may be necessary to reproduce the clinical illness in another member of the same species; either by placing healthy susceptible animals in direct contact with those showing symptoms of infection or by inoculating groups of healthy and immune (vaccinated) animals by a suitable route with a material obtained by a sick animal.
• If the latter procedure is adopted it is necessary to ensure that the material capable of transmitting the disease does not contain other microorganisms (for example bacteria) which produce a concurrent infection and complicate the clinical picture.
• To ensure that the inoculum is bacteriological sterile the material must be filtered or treated with antibiotics.

-Specific or natural host: sheep, goats, cows, chicken, monkeys, and horses

-Laboratory animals: mice, rabbits, guinea pigs, rats, and rodents

Advantages:

1. Used as a diagnostic procedure for isolating and identifying a virus from a clinical specimen
2. Mice provide a reliable model for studying viral replication
3. Gives unique insight into viral pathogenesis and host-virus relation
4. Used to study virus localization, spread, and exit in the host
5. Used for the study of immune responses, epidemiology, and oncogenesis

Disadvantages:

1. Expensive and difficulties in maintenance of animals
2. Difficulty in choosing animals for a particular virus
3. Some human viruses cannot be grown in animals or can be grown but do not cause disease
4. Mice do not provide models for vaccine development
5. It will lead to the generation of escape mutants
6. Issues related to animal welfare systems

The experiment animals are used for the following purposes

1. Virus isolation
2. To study pathogenicity and host immune reaction
3. To test and develop a viral vaccine
4. To raise monoclonal or polyclonal antibodies

1. Virus isolation – For diagnostic purposes the experimental animals are still used for example mice in rabies and louping ill disease diagno
2. To study pathogenicity and host immune reactions – This is studied in homologous host example pig in swine fever. The cost of using the homologous host is very high and therefore inbred experimental animals are used instead of the homologous host. Example inbred mice used in African swine fever. The laboratory animals used as models are –

•  Rabbit – the rabbits were used by the Pasteur to adopt the street virus of rabies. In malignant catarrhal fever virus, these animals react in a similar manner as the cattle.
• Guinea pig – guinea pigs react to foot and mouth disease virus when inoculated intradermally in the footpad. Primary vesical are formed on the footpad and secondary vesicles appear in the mouth following viremia.
• Ferrets – Ferrets are used in the study of the pathogenesis of the distemper virus.
• Other laboratory animals are also used in virus study or in the preparation of antisera against different viruses.

4. To test and develop viral vaccines – Mice, guinea pigs, and rabbits are used for attenuation of virus strains as well as for testing vaccines Foot and mouth disease virus vaccine is initially tested in guinea pigs and finally in cattle and pigs.
5. To raise monoclonal or polyclonal antibodies- Various routes are employed -to inoculate experimental animals with virus-infected material. The usual routes are Intracerebral, Intranasal, Intradermal, Intramuscular, intravenous, and subcutaneous, the route of inoculation largely depends upon the nature of the virus; its possible affinity for the tissue, age, and species of the experimental animal.

Preparation of inoculum

The material used for anima! inoculation may consist of filtered or unfiltered suspensions of organs or exudates. If the materials are unfiltered, it is important to add antibacterial substances such as penicillin and streptomycin, to prevent contamination or associated bacterial agents from becoming established. This is especially important in intracerebral inoculation, some bacteria which are ordinarily considered non-pathogenic may cause infection when directly introduced into the brain tissue of the living animal.

A general procedure for the Preparation of inoculum is as follows –

1. The infected tissue or exudate is removed from the animal with a sterile instrument and placed in a sterile container.
2. The tissue is cut into small pieces and placed in a sterile mortar.
3. An abrasive such as sterile alundum (90 mesh) is sprinkled over the tissue which is ground to a paste and then suspended in tryptose phosphate broth or other buffered liquid to make a 10-12% suspension.
4. Centrifugation at 3000 rpm for 3-5 minutes will clarify the material so that the supernatant can be used in a syringe for injection.
5. After centrifugation, the supernatant fluid is transferred to another tube and the antibiotics are added.
6. A period of 15-30 minutes incubation at room temperature is allowed before the mixture is injected. However, if  the material is  collected relatively free of contamination it can be inoculated immediately into animals and chicken embryo

Route of Virus Inoculation

• The usual routes commonly employed for the inoculation of the viruses into experimental animals are – intravenous, intramuscular, subcutaneous, intradermal, intraperitoneal, intracerebral, and  Intranasal.
• In addition,  other routes of inoculation viz.  intraplantar,  intradermolingual, corneal, scarification, cheek pouch, and peroral may also be employed in certain specialized cases.
• The sizes of the hypodermic needles are expressed as s.w.g. x inches.
• The abbreviation s.w.g. denotes standard wire gauge, which is the diameter of the needle.
• The larger is the s.w.g. number the smaller will be the needle diameter, thus 26 s.w.g. needle is a very fine one as compared with an 18 s.w.g needle. The length of the needle from the mount to the point is given in inches.

Intravenous

• Monkeys,  rabbits,  mice, and rats may be inoculated by this route.
• Inoculation into an ear vein may be done with a fine needle, otherwise, a vein in the leg should be used for inoculation.
• In monkeys, the femoral vein is used for inoculation.
• Mice and rats may be injected into the caudal vein.

Intramuscular

• It is a very simple inoculation for which a suitable muscle may be used and usually, thigh muscle is selected.
• In rodents, inoculation is done into the gastrocnemius muscle with a % inch 26 s.w.g needle.
• The needle is inserted about 2-3 mm into the flesh part and 0.05 ml is gently inoculated.
• In rabbits 1ml in one leg may be inoculated.

Subcutaneous

• Any area of the animal where the skin is loose may be used.
• In rabbits and mice injection may be between the shoulders.
• A fold of skin should be pinched between the under finger and the thumb and an injection made through the fold. In large animals, the loose skin along the flank may be used.

Intradermal

• Any suitable size may be chosen.
• The hair is removed to wash the site with water.
• The injection is made by inserting the needle horizontally at an angle so that the needle does not penetrate deep makes the injection.

Intraperitoneal

• All the laboratory animals are suitable for this inoculation,  the inoculation is made to one side of the midline of the lower abdomen.
• As the needle is withdrawn the skin around it should be gently pinched together and hold for a few seconds to prevent the inoculum from leaking back.
• For mice, the animal is held horizontally, the ventral surface uppermost, and the needle is inserted (1 inch 23 s.w.g.) through the abdominal wall about 5 mm lateral to the midline, at an angle of about 30° from horizontal.
• The needle is inserted 1 cm deep to avoid puncturing the viscera.
• A volume of 0.5 to 2 ml is usually injected although up to 5 ml may be inoculated.

Intracerebral

• In a suckling mouse, no assistance is required.
• The mouse is held firmly on the bench with the left hand in the sitting position of the mouse (dorsal surface uppermost) a % inch 26 s.w.g needle is inserted vertically through the left cranial wall at a point equidistant from the anterior margin of the ear, the posterior angle of the eye and the cranial midline.
• The needle is inserted to about 2 mm, 0.01 ml is inoculated and the needle is withdrawn.
• In an adult mouse and other animals, anesthesia is required. In an adult mouse, the animal is anesthetized with ether and placed on the bench.
• The remaining procedure is the same as described, above for a suckling mouse.
• The needle is inserted about 2 to 3 mm, 0.03 ml is inoculated and the needle is withdrawn.
• For rabbits and monkeys, drilling of a hole is required for inoculation to be done.

Intranasal

Mice and ferrets are usually employed for this route and inoculation is done with a V2 inch 23 s.w.g needle which should have a blunt tip. The animal is anesthetized, the animal is held keeping the head up, the needle is brought to the nostril and the required inoculum (0.05 ml in mice & 0.1 ml in ferrets) is dropped slowly.

Intraplanter

• The route is employed for such diseases as mousepox and foot and mouth disease in which case the planter pad (foot pad) is inoculated.
• For foot and mouth, disease guinea pigs are employed. A 1/ 1/2 inch 22 s.w.g needle, whose tip has been made slightly blunt by a  little grinding is used.
• The assistant should hold the guinea pig.
• The toes are held with the left hand by keeping them pressed with the thumb and the needle is inserted with the right hand. Several tunnels are made intradermally and inoculum is injected.
• The volume of inoculation is 0.1 ml in each pad. It is also called intradermal pad inoculation.

Intradermolingual

• This route is specially used for inoculating apthovirus (foot and mouth disease virus) in cattle.
• No sterilization of the inoculation area is done.
• The tongue of the animal is withdrawn and held tightly by taking the tip of the tongue in the grip of the assistant.
• A 1/ 1/2  inches 26 s.w.g. needle is inserted intradermally into the tongue by keeping the needle horizontal to the tongue surface and making tunnels.
• It starts from the inner side of the tongue making several tunnels and then coming towards the tip about five lines of the tunnels with five tunnels in each line are made.

Corneal scarification

• This route is employed in rabbits.
• No sterilization of the inoculation area is required.
• The animal is anesthetized with ether and placed on its side. One drop of inoculum is placed on the corneal surface and gently sacrificed with the scalpel blade.

Cheek pouch

• Hamsters are inoculated by this route. No sterilization of the inoculation is done.
• The animal is anesthetized with dither and its mouth open using the left thumb and the index finger the cheek pouch is held, the animal leg is suspended and the mucous membrane of the cheek pouch is exposed for inoculation.
• A V2 inch 26 s.w.g needle is introduced into the pouch tissue as superficially as possible and slowly 0.1ml inoculated.
• This route is employed for studying the oncogenic viruses for their ability to produce tumors in hamsters.

Peroral

• This route is employed in a few circumstances.
• The usual procedure is to hold the head high, open the mouth, drop the inoculum into the posterior part of the buckle cavity and keep the head high for some time and allow the animal to swallow the content.

Bleeding techniques

• The experimental animal may be bled either by vein puncture or by cardiac puncture.
• A vein puncture is preferred when a small volume of blood is to be recovered.

Monkey

• Bleeding is done from the femoral vein using a syringe with a 11/2 inch 21  s.w.g needle attached.
• The animal should be anesthetized before bleeding.

Rabbit

• For routine purposes where a  small volume of blood is required, bleeding may be done from the marginal ear vein without any anesthetic.
• To distance the vein and make it clearly visible the area is swabbed with xylene or alcohol.
• If a larger volume of blood is required, bleeding is done from the heart.
• The animal is placed in the supine position and the position of the heart is located by palpating with the finger.
• This may be done by keeping the fingers slightly over the left thoracic wall.
• The needle is inserted through the chest wall toward the heart and as soon as the needle touches the heart the heartbeat is felt.
• The needle is pushed inside the heart in one stroke and the plunger is slightly withdrawn so that blood entire the syringe freely.
• The required volume of blood is withdrawn and the needle is removed.

Rat, mouse, hamster

• For a small volume, the bleeding may be done from the retro ocular plexus of veins of the eye.
• A pasture pipette is specially prepared for this purpose.
• A pasture pipette is taken and the tip is cut with a glass cutting foil at a place so that the diameter of the tip is about 1ml.
• The tip is ground to make it blunt so that it may not cause any injury.
• The assistant will hold the animal keeping the dorsal surface uppermost.
• The eyelids are open with the left thumb and index finger and the tip of the pasture pipette is placed with the right hand in the orbital cavity without damaging the eye.
• This will result in bulging of the eye to one side.
• The pipette is pushed further so that its tip touches the reto-ocular plexus and the pasture pipette is rotated and then slightly the pressure of the pipette is loosened to allow the blood to be sucked in by capillary action.
• After the blood is sucked in, the pasture pipette is withdrawn and the blood is transferred into a small tube.
• When a large volume of blood is required, it is collected either from the heart or by sacrificing the animal.

Chicken

• Blood is withdrawn from the humeral vein with a 1-inch 21 s.w.g needle.
• The bird is placed in a supine position with the wing well spread.
• The feathers are plucked to expose the vein.
• The needle is inserted in the vein and its correct position is ascertained by withdrawing the plunger when the blood will flow in the syringe.
• The required volume of blood is collected and the needle is withdrawn.

Cattle sheep goat

• Blood is collected from the jugular vein.
• The hair is removed and the jugular vein is pressed with the left thumb so that the flow of blood is stopped and the vein becomes prominent.
• The needle is inserted and the blood will flow immediately with force if the needle is placed rightly.
• The desired volume of blood is collected and the needle is withdrawn. For bleeding large volumes of blood, the bleeding cannula may be used.

Routes of virus inoculation for different animals

1. Intracerebral- mice for rabies
2. Intravenous- calves for malignant catarrhal fever
3. Intramuscular- piglets for Teschen
4. Intradermal- guinea pigs for FMD
5. Subcutaneous- piglets for swine vesicular disease
6. Intranasal- ferret for RVF and calves for IBR and MD
7. Intraperitoneal- unweaned mice for FMD
8. Scarification- sheep for orf, rabies, or vaccinia
9. Intralingual- horses for vesicular stomatitis
10. Oral- monkeys for poliomyelitis
11. Aerosol- birds for avian influenza A
12. Intratracheal- chickens for infectious laryngotracheitis
13. Intraconjactival- mice for pseudorabies

It may be an advantage to withdraw blood samples at regular intervals from experimental animals to detect the possible development of specific antibodies (eg. Q fever) and to cull and carry out post-mortem examination of some animals during the course of the experiment. Clinical symptoms, the development of visible lesions, abnormal behavior, and all deaths, whatever the cause, should be carefully observed and recorded. At the termination of an experiment involving infectious material, all bedding utensils cages, carcasses and tissues should be removed, burned, sterilized, or thoroughly cleaned by the most appropriate method. Animals infected experimentally must be held in separate isolation rooms and all persons handling infected animals, cages or other contaminated materials must pay strict attention to personal cleanliness.

Special care must be taken when performing inclusions if the material is believed to contain virulent viruses or when specimens obtained from experimentally infected animals are being processed for further passage in animals or inoculated into eggs or cell cultures.

Viruses used for cultivation in different animal models

Detection of virus growth in the virus-infected animals

Infected animals are examined daily for:

i.Clinical signs of the disease such as feverish signs, respiratory distress, GIT involvement, CNS disorders, visible skin or membrane lesions

ii. Abnormal behaviour

iii. Defects in animal development

iv. Deaths

-Monitor the body temperatures – taken once or twice daily

Collection of samples

-Blood samples – at specific intervals to check for rising in antibody titre, presence of virus, and blood parameter changes

-Biopsy materials or tissue are collected at necropsy following deaths or culling at specific intervals, examined;

a) Macroscopically: for lesions-vesicle, pneumonia, hemorrhages

b) Histologically: pathological changes- inclusion of bodies

c) Serologically– IFAT, ELISA, for specific viral antigens

d) Microscopically using EM to observe the virus particles

e) By isolation of the infectious virus