Investigator and staff handbook laboratory animal program

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Cage Cards

Cage cards are utilized to identify the animal source, stock or strain of rodent, Principal Investigator, protocol number, account number, Vivarium Request Form number, sex, cage ID, and date received. Cage cards containing accurate information must be displayed for all animals. Normally, individual cards are secured to each cage. If, however, identical information is applicable to all animals on a given rack or in a room, a single card may be affixed to the rack or room door. Cage cards should not be removed and should be amended with updated information as needed.

Animal Identification Methods

  1. Multicolored animals may be identified by their natural markings.

  2. Permanent markers may be used on the tails or fur for temporary identification.

  3. Hair clipping may last up to 14 days.

  4. Ear tags may be used but can be difficult to read and may be lost.

  5. Ear punching can be utilized but may be obliterated by fighting between individuals.

  6. Tattoos: Tails are most commonly used because the lack of hair allows easy visual identification of the tattoo.

  7. Fur Dyes may be used for temporary identification.

  8. Implantable Microchips: This system uses a computer microchip which is injected subcutaneously into the animal. A portable, programmable computer scanner is then used to identify the animal.

Tail and toe clipping are not recommended.

General Information
To limit contact exposure to potent rodent urine and dander allergens, it is recommended that personnel wear disposable examination gloves when handling animals. Protective leather or chain-mesh gloves are not usually needed when handling rodents but may be used in select circumstances for handling fractious animals. If protective gloves are used, additional care should be taken to avoid inadvertently injuring the animal through overzealous handling. Rodents may become accustomed to regular handling, which can benefit both the animals and personnel.

Mice, rats and gerbils may be picked up by the base of their tails and transported short distances but should not be suspended by their tails for prolonged periods. Grasping rodents by the base the of tail may be used for removing animals from their cages, transferring between cages, cursory examinations, and sex determinations, but may not be adequate for treatments and close examination. Grasping and/or pulling the tip of the tail may strip the skin from the tail. If a degloving wound occurs, amputation of the injured tail is advisable. When returning rodents to their cage, gently lower and release them upon the floor of the cage. Dropping rodents into a cage can result in injuries.

Hand Restraint
Grasp the mouse by the tail base and place on a surface (e.g., wire bar cage top) the animal is able to grip with its feet. Then, pull gently on the tail as the mouse holds onto the surface with its forelimbs. While maintaining traction on the tail, grasp the loose skin over the neck, immediately behind the ears, with the thumb and forefinger of the opposite hand. It is necessary to perform this maneuver expeditiously to avoid being bitten. If the skin over the shoulders or an insufficient amount of loose skin is grasped, the mouse will be able to turn and bite. If the skin is grasped too tightly, the animal's respiration can be compromised. Once the loose skin over the neck is appropriately secured, the mouse may be lifted and the tail secured by the fourth and fifth fingers of the same hand. This method of restraint is adequate for most technical manipulations.
Grasp the rat by the tail base and place on a firm surface. Then, with the opposite hand, encircle the body over the shoulder and neck region. It is necessary to perform this maneuver expeditiously, or the rat may turn and bite. Do not grip the chest so tightly that respiration is hindered. Once the head and body are secured, the rat may be lifted. This method of restraint is adequate for most technical manipulations.
Hamsters are considered by some to be difficult to handle. However, if simple precautions are taken, hamsters can be routinely handled with minimal stress to the animal and handler. Nonetheless, care must be taken to avoid being bitten. Factors that affect hamster restraint are the abundant amount of loose skin, which must be gathered to immobilize the animals, the tendency to bite when startled, and the predisposition to sleep deeply and then awaken suddenly.
Hamsters can be removed from their cage or transferred between sites using a small can or cup. To do so, place a can or cup into the animal's cage and gently encourage the hamster to enter. After entering, the opening can be covered and the hamster removed from its cage. Alternatively, hamsters may be picked up by the loose skin over their back. With the hamster on a flat surface, place the palm of the hand over the hamster and press gently downward to immobilize the animal while gathering the abundant amount of loose skin over its neck and back. When an adequate amount of skin is secured, the hamster will be restrained and unable to bite.
Guinea pigs
Guinea pigs should always be restrained using both hands. One hand should gently secure the animal around the torso while the second hand supports the hindquarters. This is particularly important with large or pregnant animals to avoid internal injuries. If animals become fractious, subdued lighting and covering the eyes may have a calming effect.
Gerbils can be restrained by grasping the base of the tail with one hand and the skin on the back of the neck with the opposite hand. Extreme care must be taken not to pull on the tail because gerbils are especially prone to tail injuries from improper handling. If additional restraint is desired, gerbils may be grasped from above, securing the animal's head between the index and middle fingers and the body with the thumb and remaining fingers.
Handling neonatal rodents should be avoided especially during the first few days after birth to avoid cannibalism or litter abandonment. If it is necessary to handle immature litters, the neonates and the litter surrounding them may be scooped up with a gloved hand to avoid contamination with human scent. Individual pups may be picked up by grasping the loose skin over the neck and shoulder with thumb and forefinger.

Order: Rodentia

Suborder: Myomorpha

Family: Muridae

Mus musculus (laboratory mouse)

Rattus norvegicus (laboratory rat)

Family: Cricetidae

Mesocricetus aruatus (golden or Syrian hamster)

Cricetulus griseus (chinese or striped hamster)

Crisetus cricetus (European hamster)

Cricetulus migratorius (Armenian hamster)

Phodopus sungorus (Russian, Siberian, Dzungarian, Djungarian or

furry-footed hamster)

Meriones unguiculatus (Mongolian gerbil)

Suborder: Hystricomorpha

Family: Caviidae

Cavia porcellus (Guinea Pig)
Genetic and Microbial Categories
Rodents are characterized by genetic and microbial status. Common genetic categories are outbred stocks, inbred strains, F1 hybrids, transgenics, knockouts, and mutants. Outbred stocks maintain genetic diversity by mating unrelated individuals. Inbred strains maintain genetic homozygosity by breeding closely related relatives. F1 hybrids are a cross between two inbred strains. Transgenic animals have specific genetic material introduced into their genome. Knockouts have specific genetic material removed from their genome. Mutant animals are strains that have spontaneously developed genetic mutations.
Microbial flora of rodents can also be used to categorize rodents. Axenic rodents are free from of all microbial organisms, including both pathogens and normal commensal flora. Gnotobiotic rodents harbor only known microbial flora. Specific Pathogen-Free (SPF) rodents are free from specific bacterial, viral, and/or parasitic pathogens. Conventional rodents have an unknown microbial status. Most rodents purchased at SIUC are SPF rodents. Rodents housed in the LAP Vivarium are periodically tested for the presence of pathogens to determine their current microbial status. If rodents of different microbial statuses are worked with in the same day, the room with the fewest pathogens must always be entered first, followed by the room(s) with progressively more pathogens. If in doubt, contact personnel from the LAP for recommendations.

Anatomic and Physiologic Characteristics
Rats, mice, and hamster incisors and all guinea pig teeth grow continuously throughout life and are worn down by mastication (chewing). If teeth are not properly aligned or fail to be worn down for any reason, dental overgrowth can occur. Overgrown teeth can inhibit eating and drinking.
Glands behind the eyes of rats can produce dark red secretions that can stain the fur around the eyes and nose in response to stress or illness. The coloration is the result of porphyrin pigment and not blood, despite its appearance.
Guinea pigs differ from rats and mice by having a longer gestation period, precocious offspring, and a cellular membrane that closes over the vaginal orifice, except during estrus and parturition. Guinea pigs’ pubic symphysis separates during the latter half of gestation to allow passage of young during parturition. If guinea pigs are bred for the first time after 7-8 months of age, the symphysis separates incompletely and causes delivery difficulties.
Hamsters have highly distensible cheek pouches that extend from the oral cavity to the shoulder blades. The cheek pouches may be used to transport or store food, bedding, or other objects. Hamsters may also conceal young in their cheek pouches when threatened, however the young may suffocate if not removed in a timely manner.
Spontaneous, convulsive (epileptiform) seizures occur in approximately half of gerbils. The seizures range from mild to severe and are often elicited in unfamiliar surroundings or by a startle stimulus.
Most rodents are primarily nocturnal (during dark), but can also display diurnal (during light) activity. Unlike other rodents, guinea pigs and gerbils may have peak activity levels during the dark or light cycles.
Hamsters are not true hibernators but may become pseudohibernated in response to decreased environmental temperatures, decreased length of light cycle or other variables. During pseudohibernation, hamsters remain sensitive to touch and may become aggressive on awakening.
Rodent littermates raised together from birth usually coexist peacefully. Rodents may fight when grouped together after reaching puberty, reunited after prolonged separation, or held in crowded conditions. Fight trauma may result in abscessation, dermatitis, septicemia, or death. In most cases, males are more prone to fighting than females. However, female hamsters are dominant over males and can be aggressive. Whenever groups of rodents are assembled, or when new animals are added to a stable group, the group should be observed carefully to prevent fight injuries. When fighting occurs, the animals involved must be separated.
Rodents may also establish hierarchy status via barbering. During barbering, the dominant animal chews hair from the muzzle, body, or tail of subordinate animals.
The anogenital (between the anus and genitalia) length can be used in most rodents to determine sex in neonates and adults. Anogenital length is greater in males versus females. All rodents have open inguinal canals throughout life allowing the testicles to be retracted into the abdominal cavity and therefore may not be visible. Most adult male rodents are larger than adult females. Adult female hamsters are larger than adult males, with the exception of the Chinese or striped hamster in which the males are larger. Viewed from above, the rear margin of the male hamster is rounded because of the scrotal sacs, and the female posterior is pointed toward the tail.
Differentiation of sex in guinea pigs is slightly different from that in other rodents. The guinea pig female has a vaginal closure membrane that can be exposed by gentle digital stretching of the genital ridge that extends from the anus to the vulva. The relaxed membrane will be seen as a shallow, U-shaped crease between the anal and urethral openings. The vaginal closure membrane is intact except during estrus and parturition. Male guinea pigs (boars) are distinguished from females (sows) by a penis that can be extruded from the circular prepuce. Also, the boar has no break in the ridge between the urethral-penile opening and the anus.


Always use sterile syringes and needles. To ensure aseptic techniques and sharp needles, the one-time use of sterile disposable supplies is strongly recommended. Following usage, the syringe with the attached needle should be discarded into an appropriate puncture-proof biohazardous container. Do NOT replace caps onto needles before discarding. Recapping needles can lead to inadvertent needle sticks of personnel.

Orbital Sinus

  • Orbital sinus bleeding must only be performed on rodents under general anesthesia!

  • Retract the skin of the eyelids to cause slight protrusion of the eye.

  • Place hematocrit tube or small-bore Pasteur pipet at the inner corner of eye, adjacent to the eyeball, and displace the globe laterally.

  • Rotate the tube while pressing it caudally towards the rear of the orbit until blood enters the collection vessel by capillary action.

  • Remove the tube/pipet from the orbit, close the animal's eyelids and apply gentle pressure to ensure good hemostasis.

  • It is recommended to alternate eyes if more than one blood collection is planned.


  • Depending on the species used, blood may be collected from the central tail artery or the lateral tail veins.

  • Restrain rodent with the tail exposed.

  • Dilate the tail vessels using either a tourniquet around the base of the tail or immersion of the tail in water not exceeding 40oC.

  • With the tail under gentle traction and the needle at a very shallow angle to the tail, insert needle into the lumen of the vessel. The lateral tail veins lie immediately beneath the skin on each side of the tail. The central tail artery is midline on the ventral surface of the tail.

  • Gentle massaging of the tail towards the needle may help facilitate bleeding.

  • Remove the needle and apply gentle pressure to the site of entry to ensure good hemostasis.

Axillary Vessels

  • Brachial plexus blood collection must only be performed on rodents under general anesthesia!

  • Place anesthetized rodent on its back and secure its legs to the counter or table top.

  • Incise the skin between the thorax and the forelimb, severing the axillary vessels. Blood will pool in the area formed by the incision which may then be aspirated.

  • This procedure should be followed by euthanasia if the animal has not expired as a result of hypovolemia.

Intracardiac (for large-volume collection)

  • Due to the inherent risks associated with the procedure, cardiac bleeding should only be performed as a non-survival procedure.

  • Intracardiac blood collection must only be performed on rodents under general anesthesia!

  • Place anesthetized rodent on its back (i.e. dorsal recumbancy).

  • Personal preference will dictate whether the needle is inserted through the ventral midline or lateral thorax. For ventral midline entry, the needle is introduced just to the left of the xyphoid process (base of sternum) towards the heart. For lateral thoracic entry, the needle is introduced through the left thoracic wall.

  • After the needle tip is through the skin, gently pull back on the syringe plunger to create negative pressure and continue to advance the needle until blood can be withdrawn. If blood is not able to be collected, slowly withdraw the needle while maintaining negative pressure on the syringe until the needle is nearly, but not completely free, of the skin and redirect in a slightly different direction.

  • This procedure should be followed by euthanasia if the animal has not expired as a result of hypovolemia.

General Information
Always use sterile syringes and needles. To insure aseptic techniques and sharp needles, the one-time use of disposable supplies is strongly recommended. Following usage, the syringe with the attached needle should be discarded into an appropriate puncture-proof biohazardous container. Do NOT replace caps onto needles before discarding. Recapping needles can lead to inadvertent needle sticks of personnel.
When administering injections, select the smallest gauge needle possible to minimize tissue trauma and injection discomfort. Before injecting the solution, always check for correct placement of the needle by gently pulling back the plunger of the syringe to create a vacuum without moving the needle within the animal. Blood, urine, or fecal debris may be aspirated depending on the injection site selected. Aspiration of any material indicates improper placement of the needle. If this occurs, the needle should be withdrawn and both the needle and syringe appropriately discarded.
The manufacturer’s expiration date should be checked prior to administering drugs to animals. Only in-date drugs should be used. Expired or out-of-date drugs should be appropriately discarded or stored separately from in-date drugs to prevent inadvertent animal administration. Pharmaceutical-grade medications must be used whenever they are available, even in acute procedures. Non-pharmaceutical-grade chemical compounds should only be used in regulated species after review and approval by the IACUC for reasons such as scientific necessity or non-availability of an acceptable veterinary or human pharmaceutical-grade product. Cost savings alone is not an adequate justification for using non-pharmaceutical-grade compounds in regulated animals.
Injection Sites
Intramuscular (IM)

  • Due to the limited muscle mass of many rodents, intramuscular injections may cause discomfort and tissue irritation and are generally discouraged.

  • Insert needle into the center of the muscle of the rear leg and aspirate before injecting.

Subcutaneous (SQ)

  • Grasp and gently raise the skin over the dorsum or sides of the animal.

  • Insert the needle through the base of the raised skin at a shallow angle to the skin surface being careful not to exit the skin or enter the underlying muscle.

Intraperitoneal (IP)

  • Restrain the rodent with the ventral side up (i.e., on its back).

  • Insert the needle parallel but off midline into the lower half of the abdomen just enough to enter the peritoneal cavity and aspirate before injecting.


  • Measure the distance from the tip of rodent’s nose to the last rib to determine the appropriate length of gavage needle to be used.

  • Restrain the rodent, including its head, in one hand with the head up and straight with the body.

  • Insert the ball-tipped gavage needle at the side of the animal’s mouth.

  • Slowly and gently advance the gavage needle down the animal's esophagus and into the stomach. Forceful passage of the gavage needle risks perforation of the animal’s esophagus.

  • Signs of improper placement include: resistance of the gavage needle’s passage; inability to advance the needle to the desired depth; or severe animal struggling. If there is any doubt that the gavage needle may not be properly positioned, remove the gavage needle and reposition before administering the syringe contents. If material is inadvertantly administered into the lungs, the animal should be humanely euthanized.

Intravenous (IV) - Lateral Tail Vein

  • Restrain rodent with the tail exposed.

  • Dilate the tail veins using either a tourniquet around the base of the tail or immersion of the tail in water not exceeding 40oC.

  • With tail under gentle traction and the needle at a very shallow angle to the tail, insert needle into the lumen of the vein approximately 2-3 mm. The lateral veins lie immediately beneath the skin on each lateral side of the tail.

  • If the needle is properly positioned, aspiration may cause blood to enter the needle hub.

  • If a tourniquet is used, it should be released before injecting.

  • If a bleb appears when injecting, needle placement was not successful. If a bleb does not appear, or the blood in the lumen of the vessel proximal to the injection site is cleared during injection, needle placement was correct.

Intradermal (ID)

  • Clip the hair overlying the injection site.

  • With the needle bevel down and the needle nearly level with the skin surface, insert needle into the skin 2-3 mm.

  • A successful intradermal injection produces a distinct skin welt.

Analgesics for Laboratory Animals

Dosage Rates for Analgesic Drugs in Laboratory Animals

Drug Name and Indications





1-2mg/ml drinking water

1-2mg/ml drinking water

1-2mg/ml drinking water

Buprenorphine-mild pain; longer duration of action

0.05-2.5 mg/kg IP or SC every 6-12 hours

0.01-0.5 mg/kg SC every 8-12 hours

0.01-0.05 mg/kg SC or IV every 6-12 hours

Butorphanol-moderate pain; short duration of action

0.2-2.0 mg/kg SQ,IP every 4 hours

0.2-2.0 mg/kg SC,IP every 2-4 hours

0.1-0.5 mg/kg SC every 1-2 hours

Carprofen-Nonsteroidal anti-inflammatory drug

5 mg/kg SC or PO every 24 hours

1.5 mg/kg SC,PO, every 12 hours

4 mg/kg SC or 1.5 mg/kg per os every 6-12 hours

Flunixin meglumine


2 mg/kg SC,IM every 12 hours

0.1-0.5 mg/kg SQ every 12 hours

Ketoprofen-Nonsteroidal anti-inflammatory drug

5 mg/kg SQ every 12-24 hours

5 mg/kg SC every 12-24 hours

3 mg/kg IM or SC every 24 hours

Meloxicam-Nonsteriodal anti-inflammatory drug

1-2 mg/kg PO Every 24hours

1-2 mg/kg SC or 4 mg/kg PO

0.2-0.4 mg/kg SC or 0.2-0.6 mg/kg PO

Morphine- moderate to severe pain; short duration of action

2.5 mg/kg IP every 4 hours

2.5 mg/kg IP or IM every 4 hours

2.5 mg/kg SC,IM, every 2-4 hours

Analgesic effectiveness must be evaluated in each animal due to variations in response between individuals and strains. For more information or analgesic guidelines in other species please consult the attending veterinarian.

Adopted from University of Pennsylvania Animal Resources Program and Exotic Animal Formulary, 3rd edition, 2005, ed. Carpenter, J., Elsevier Saunders, St. Louis, Misssouri

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