Species and Techniques Information | Office of Scientific Affairs

The laboratory mouse (Mus musculus) is a mammal of the order Rodentia. The laboratory mouse has been domesticated by man for many generations and is in general fairly easy to handle.
Other notable biological characteristics are their very acute hearing, well developed sense of smell, poor vision, small size and short generation interval. While mice are often considered to be the most common laboratory animal used for research, studies utilizing rats are more frequently cited in the scientific literature.
"Out-bred" mice are managed to maintain genetic diversity by mating unrelated mice.
"Inbred" mice are managed to maintain genetic homozygosity by breeding siblings for more than 20 generations.
"F1 hybrid mice" involve crossbreeding two inbred strains for one generation.
"Transgenic mice" are mice in which specific genetic material has been introduced into the genome of another inbred mouse strain. "Knockout mice" are mice in which specific genetic material has been removed from the genome.
"Mutant" mice are inbred mice that have developed spontaneous genetic mutations.

B. Unique Biological Characteristics

The mouse does not have an appendix.
The urine of mice is normally clear, yellow and very concentrated. Large amounts of protein are normally excreted in the urine.
Dark pigmentation of the spleen is a non-pathogenic condition in C57Bl mice. This pigmentation may be focal.
Barbering is the removal of hair and/or whiskers and can be seen when animals are group housed. The area of hair loss is usually well demarcated and dominant mice barber subordinate mice. There are normally no skin wounds associated with barbering.
Mice are nocturnal (most active at night), however substantial activity can also be seen during the day. Mice, like most species have a circadian rhythm. Investigators should be aware that this may affect biological data and should standardize the time of day that samples/measurements are taken to avoid this affect. The standard light/dark cycle in the DLAR is 12/12. This light cycle can be modified upon the request of the investigator in special circumstances.
The adult mouse weighs approximately 20-40 grams. Their small size, and resulting large surface area/body weight ratio, makes them highly susceptible to changes in environmental conditions (i.e. hypothermia or hyperthermia). The core body temperature is easily affected by small changes in temperature that may modify the physiologic responses of the animal.
Mice have a high ratio of evaporative surface to body mass and is therefore more sensitive to water loss. The biological half time turnover for water in the mouse is 1.1 days. Mice use surface evaporation to compensate for increases in body temperature. They can easily dehydrate leading first to shock, then death.
Mice can compensate better to cold than hot ambient temperatures.
The acute hearing of mice makes them highly sensitive to ultrasounds and high pitched noises inducing a stress response that has been empirically related to cannibalism of pups by their dams.
The well developed sense of smell is used to detect pheromones used in social interactions.
The poor vision of mice makes them unable to detect color and red light is often used to observe animals during the dark cycle.

Experimental Uses

Cancer research
Monoclonal antibody production
Immunology
Molecular genetics
Dermatology
Toxicology

Sources and Ordering

Inbred, outbred, hybrid, transgenic, knockout or mutant mice are available from many commercial sources. The DLAR office staff is familiar with many of the commonly available strains and will assist in locating animals available for purchase.
Commercially available mice from Charles River and Taconic must be ordered by Tuesday at 9:00 a.m. for delivery on Thursday of that week. Mice from Jackson Laboratory must be ordered by Monday at 12:00 p.m. for delivery on Wednesday of that week.
Animals should be held for 48 hours before experimental use to allow recovery from the stress of shipping.

Behavior

The laboratory mouse is a docile animal and can be easily handled. Animals can be grouped soon after weaning usually coexist peacefully.
Some strains of mice (i.e. BALB/cJ, SJL/J, HRS/J) are prone to fighting and will begin to fight even if grouped at weaning. Housing males from one litter or pairing males from different litters prior to weaning may help to diminish fighting.
Breeding males that have been removed from breeding cages and caged together will usually fight. Wounds on the tail are a good sign of aggression between cage mates. You can normally determine which mouse is the aggressor because they will be the animal without skin lesions. This animal should be separated into another cage, however, once he is removed, another male may take on the aggressor role. In some cases when fighting is a problem, all males must be singly housed.

Basic Biological Data

Adult body weight: male	20-40gm¹
Adult body weight: female	20-40gm¹
Body surface area	10.5(wt. in grams)2/3¹
Life Span	1.5-3 years¹
Food consumption	15 gm/100 gm/day¹
Water consumption	15 ml/100 gm/day¹
Breeding onset:male	50 days¹
Breeding onset:female	50-60 days¹
Gestation Period	19-21 days^1,³
Body Temperature	36-37C²
	36.5-38.0C¹
Heart rate	500-600 beats per minute²
	325-780 beats per minute¹
Respiratory Rate	84-230 per minute²
	60-220 per minute¹

Sexing and Reproduction

Male and female mice can be differentiated by observing the distance from the anus and genital papilla which is greater in males. This difference is also present in neonatal mice.
Breeding systems can be monogamous (one male and one female) or polygamous (one male to 2-3 females). Female mice have a postpartum estrous.
The mouse is polyestrous and the female cycles every 4 to 5 days.
Evidence of mating is normally detected in the female by the presence of a copulatory plug.
Gestation in the mouse is 19-21 days, depending on strain. Litter size varies greatly between strains of mice and litter size in an individual female will increase, plateau and then dimish over her reproductive life. Genetics, environment and embryonic mortality can also influence litter size. Pups are generally weaned at 21 days of age.

Basic Husbandry

Mice can be grouped by their microbial colonization.
- Specific Pathogen Free Mice (SPF) mice are free from specified bacterial, viral, and parasitic mouse pathogens. To insure the maintenance of the SPF microbial status requires that mice be housed in more stringent conditions that prevent the introduction of rodent pathogens (i.e. maximum isolation housing).
- Conventional mice are not known to be free of pathogens. Standard housing (i.e. conventional housing) cannot prevent the introduction of mouse pathogens.
- Other less common categories are axenic mice (i.e. free from all microbial organisms) and gnotobiotic mice which have a known microbial flora. Axenic mice are normally housed within germfree isolaters and gnotobiotic animals may be housed in germfree isolaters or under maximum isolation conditions.
Choosing the proper housing conditions for mice is very important. Rodent pathogens often do not produce clinical signs in affected animals but often have immunomodulating effects. Conventionally housed animals are exposed to room air and therefore may readily contract viral, bacterial and/or parasitic diseases.
Immunocompromised animals (i.e. athymic, SCID, transgenic, knockout mice) are very sensitive to opportunistic agents, and must be housed under maximum isolation conditions to allow for their long term survival. Valuable animals and animals that may need to be transported to other institutions should also be housed under maximum isolation conditions.

Conventional Housing

Most mice are housed in shoebox cages composed of polypropylene (opaque) or polycarbonate material (clear) with a wire bar lid used to hold the water bottle and feed. Bedding is placed directly into the shoe box cage allowing the absorption of urine and the animal to burrow and/or den. This type of cage will hold 4-5 adult mice depending on the size of the cage.
When removing the lid from this type of cage it is important to remove the water bottle to prevent spillage. If the cage is to be transported, the bottle should be turned sipper tube up toprevent spillage during transport. However, you should remember to turn the bottle back over to allow access to water after transport.
It is important to replace the wire bar lid securely on the top of the cage after removing mice or changing mouse cages. Mice will readily climb out of a cage. Also, since mice climb readily, be sure not to crush the mouse when replacing the wire bar lid.
The husbandry staff changes cages once per week (singly housed) or twice per week (multiply housed), thereby providing the animal a clean cage with new bedding, food and water. Water bottles and feed hoppers are checked daily by animal caretakers. Water bottles should always be checked to insure free water flow (i.e. no air locks).

Maximum Isolation Housing

Mice housed under maximum isolation conditions are housed in sterilized, polycarbonate cages with microisolater bonnets. Bedding is sterilized within the cage and allows the absorption of urine and the opportunity for the animal to burrow. This type of cage will hold 3-4 adult mice depending on the size of the cage.
Animals are provided irradiated food and hyperfiltrated water.
Personnel are required to wear protective clothing (sterile gowns, gloves, mask, head covers and foot covers) while working in the room.
All experimental work and husbandry care must be conducted within a laminar flow hood.
If animals are removed from the housing area and taken to the investigators laboratory, they may not be returned (terminal procedure only).
Access to maximum isolation housing areas is limited to prevent inadvertent fomite transmission of rodent pathogens by personnel.
All research personnel must be trained by the DLAR Assistant Director before working in this area.

Suspended cages & conventional housing

Some mice are housed on wire mesh bottom cages to allow collection of feces and urine or to prevent contact with bedding.
This type of housing is not preferred and is used only when dictated by experimental design (i.e. toxicological research).
Most suspended cages have water bottles but are occasionally provided with automatic watering. Typically a nipple valve (i.e. lixit) is located in the back of the cage that is operated by animal contact. When replacing a suspended cage that is provided with automatic watering it is important to push the cage fully into the rack care to insure that the lixit fully extends into the cage, allowing the animal access to water.
A mouse will sometimes jam the lixit valve open, resulting in constant dripping of water, which saturates the bedding material. Upon observation the bedding will appear much darker. The cage should be immediately changed, either by research personnel or by a husbandry staff member since mice can die due to hypothermia from chilling.
Pelleted natural ingredient diets are used to feed all rodents and are composed primarily of cereal grains which are supplemented with additional protein, vitamins and mineral. Due to the nature of this type of diet the exact composition can vary substantially from lot to lot.

Environmental Parameters

Room temperature: 64-79 F
Humidity: 30-70%
Ventilation: 10-15 air changes per hour

Identification

Cage cards are utilized to identify the strain of mouse, sex, number, principal investigator, and research protocol.
Cage cards should not be removed from the cage to avoid misidentification of the animals.
Temporary identification of individual mice can be accomplished by pen marks on the tail, hair clipping or dyeing the fur. Pen marks will only last 1-2 days whereas hair clipping may last up to 14 days.
Permanent identification methods can be achieved by tail tattooing which can be performed by veterinary personnel upon request (future picture). Ear tag or ear punch identification can also be utilized but may be obliterated by fighting between individuals. Subcutaneous microchip transponders are easy to implant and work well but are expensive. Transponders can, however, serve a dual function since they can be utilized to monitor body temperature.
Tail and toe clipping are not generally recommended.

Handling

When handling mice it is advisable to wear gloves to prevent the development of allergies due to direct contact with animal allergens.
Mice are usually caught and lifted by the tail. The tail should be grasped between its midpoint and the mouse's body. The tail may be grasped with the thumb and forefinger or by the use of smooth-tipped forceps. With this simple method of holding, they may be transferred to another cage, identified, examined or sex may be determined.
Pregnant mice or very obese mice may be handled by this method but they should be supported by use of the second hand placed under their feet. However, such restraint is not sufficient for treatment and close examination.
For more effective control, the mouse may be held by the tail and placed on a table or other surface, (preferably one such as a wire cage lid that the mouse can grasp) and the loose skin over neck and shoulders grasped with thumb and fingers. It is necessary to perform this maneuver expeditiously, or the mouse may turn and bite.
Once the mouse is grasped correctly, the head is adequately controlled. Restraint is improved if the tail or the tail and rear legs are held by the third and little fingers of the same hand or with the other hand.
Mice should not be dropped into the cage, which may result in spinal fracture, but should be lowered into the cage and released upon contact with the bedding.
Mice less than two weeks of age can be handled by grasping the loose skin over the neck and shoulder with thumb and forefinger or smooth tipped forceps. Handling neonatal mice should be avoided especially during the first few days after birth to avoid cannibalism or litter abandonment by the mother. If it is necessary to handle the litter, remove the dam to a separate cage and handle the neonates using plastic gloves to avoid contamination with human scent. Multiparous females are less likely to cannibalize if they have been successful mothers and should be chosen if litter manipulation is necessary.
When handling mice remember not to squeeze the thoracic cavity because this can cause the mouse to have decreased respiratory efficiency and trauma can occur to the chest and lungs.

Restraint Devices

Numerous types of restraint devices are commercially available to restrain mice. Quality devices prevent the animal from turning around yet allow easy access to the tail or legs. Devices should also be easy to clean and provide adequate ventilation.
For tail vein injections a wire box cage top can be turned over and the tail gently passed through the wire bars preventing the mouse from turning (future picture).

Transport

Mice that are moved indoors can be transported in their cage after removing the water bottle and placing it upright in the cage lid to prevent spillage. A clean mouse cage can be obtained from the cage wash area and one half inch of bedding material added. If you cannot locate a clean cage, ask a supervisor or caretaker in the area for assistance.
A wire bar lid should be placed over the cage to secure the animal during transport. If the animal is going to be in the laboratory for more than an hour a water bottle should also be obtained.
During transport the water bottle should be placed upright in the cage lid to prevent spillage.
Transport always results in some stress to the animal, however, animals should recover from indoor transport within their own cage within an hour. No recovery time may be needed if the animals are moved with care and have become accustomed to routine transport.
It is recommended that a permeable drape be placed over the cages to darken the cage and prevent over arousal of the mice during transport.
Only DLAR husbandry or veterinary staff members can transport mice to other buildings or to other animal facilities. Investigators can request animal transportation by contacting the business office at 444-2194. Transport requests should be made 48 hours in advance to allow scheduling of staff.

Environmental Enrichment

Group housing, to allow social interaction, is a primary source of environmental enrichment. All mouse cages also have an additional shredded paper material (Enviro-dri ) added to their cage to encourage nest building.
Frequent handling by personnel is important and may increase the ease of working with these animals.
If animals cannot have environmental enrichment opportunities because of the nature of the research, please contact the DLAR Director or the Clinical Veterinarian (444-2194).

Occupational Health/Zoonosis

Zoonoses are infectious diseases that can be passed from animals to humans.
Potential zoonotic diseases associated with mice include:
- Hantavirus infection Humans that acquire this infection have severe hemorrhagic disease with involvement of the kidneys. Humans may experience fever, respiratory symptoms and myalgia. The virus is most commonly seen in wild rodents and is shed in the saliva, urine and feces. Mice show no clinical sign of disease. Mice from commercial vendors should be free of Hanatavirus, however, since the possibility of infection always exists, personnel should wear gloves and a face mask when working with mice.
- Lymphocytic choriomeningitis (LCM) Humans that acquire this disease have variable symptoms ranging from mild flu-like symptoms to central nervous system symptoms. The virus can be transmitted by direct contact with urine and feces or from a bite wound. Commercial colonies are normally free of LCM virus, however transplantation of tumors into mice may serve as a reservoir for the agent. Mice show no signs of disease. Hamsters may also carry this virus.
- Tapeworms The tapeworm Hymenolepis nana rarely infects laboratory bred mice and may infect humans.
Mice can inflict sharp, deep bites that can easily become infected. As always, it is important to clean the wound promptly and seek medical attention, when appropriate.
A tetanus vaccination is required for all personnel working with animals.
Allergy to mouse dander and urine is not uncommon and usually develops within a few years of working with the animals. Sensitive personnel should wear personal protective equipment such as face masks and/or respirators (properly fitted), gloves and a lab coat. Personnel should alert Occupational Medicine during their annual health risk assessment and if allergy is a problem, obtain advice and treatment from the Occupational Medicine physician.

Health Surveillance

A. Monitoring

Investigators should monitor their animals and animal colonies routinely for common signs of illness, such as:
- loss of appetite
- weight loss
- diarrhea
- nasal or ocular discharge
- lethargy
- unkempt appearance
- decreased reproductive efficiency
- increased neonatal mortality
- skin lesions
- changes in experimental results (i.e. immunological, behavioral)
- sudden deaths
The frequency of monitoring of the animal by the investigator is stated in their approved IACUC protocol. Early endpoints are also defined in the protocol and the investigator is responsible for euthanize their animals when these early endpoints are reached.

B. Reporting Sick Animals

If an animal is identified with either experimental or non-experimentally related illness, the investigator should notify the DLAR veterinary staff at 444-2194 between the hours of 8:00 a.m. and 5:00 p.m., Monday through Friday.
If the investigator needs to contact a veterinarian on the weekend, the veterinary on-call list is posted in the main entry area of all DLAR facilities and provides the name, home phone number and pager number of the veterinarian on duty.
When contacting the veterinarian, please provide the following information:
- Investigators name/ your name
- Species of animals and animal ID number
- Location of the animal (building, room #, rack or cage #)
- Signs of illness
- Description of the experimental manipulations performed, if any.
- Phone number where you can be reached.
In emergency situations, if an animal needs immediate veterinary care, call the main DLAR office at 444-2194. The DLAR office will send an emergency (911) page or radio the veterinarian. If the emergency occurs after hours, call the veterinarian directly.

References

Harkness JE, Wagner JE. The Biology and Medicine of Rabbits and Rodents; 3rd Edition. Lea and Febiger, Philadelphia, 1989.
The UFAW Handbook on the Care and Management of Laboratory Animals; 6th Edition; editor Poole, TB. Longman Scientific & Technical, England, 1986.
Jacoby RO, Fox JG. Laboratory Animal Medicine, Academic Press, New York, New York, 1984.
Suckow MA, Danneman P, Brayton, C. The Laboratory Mouse. CRC Press, New York, New York, 2001.

Biomethodology of the Rat

Sexing and Reproduction

Restraint Devices

Transport

Environmental Enrichment

Occupational Health/Zoonosis

Health Surveillance

References

General Biology

A. Nomenclature and Breeds

The Norway rat or laboratory rat (Rattus norvegicus), is a mammal of the order Rodentia. The laboratory rat was the first animal where the primary reason for domestication was for use in scientific endeavors.
"Out-bred" rats are bred to maintain genetic diversity by mating unrelated rats (less than 1% inbreeding per generation) and are referred to as stock. To be designated as outbred, these rats must be maintained in a closed colony for a minimum of 4 generations.
"Inbred" rats (such as Fischer 344, Wistar) are bred to maintain genetic homozygosity by utilizing a brother X sister breeding scheme for a minimum of 20 generations. Inbred animals are referred to as strains.
The "F1 hybrid rats" are produced by crossing two inbred strains for one generation.

B. Unique Biological Characteristics

The acute hearing of rats makes them sensitive to ultrasounds and high pitched sounds.
The vision of rats is very poor and they are unable to detect color and are blind to long-wave (red) light.
The tail of the rat is the principal organ for heat exchange.
Rats are nocturnal.
Rats have no tonsils, water taste receptors, sweat glands or gall bladders.
Rats do not vomit.
Malocclusion (mal-alignment of the teeth) may occur in rats due to trauma, genetics or feeding a soft diet that prevents proper wearing of teeth.

Experimental Uses

Aging studies
Oncology
Toxicology
Teratology

Sources and Ordering

Inbred, outbred, hybrid, or mutant rats are available from many commercial sources. The DLAR office staff is familiar with many of the commonly available strains and will assist in locating animals available for purchase.
Commercially available rats from Charles River or Taconic must be ordered by Tuesday at 9:00 am for delivery by Thursday of that week. Animals should be held for a minimum of 48 hours before experimental use to allow recovery from the stress of shipping.

Behavior

The rat can become accustomed to handling providing they are not upset by the experience.
Rats will bite without warning, but usually not repeatedly. Males tend to be more aggressive than females.
Unlike mice, groups of the same sex can be housed together without fighting.
Rats are active primarily during the night at which time they feed; the light hours are used primarily for rest, sleep and digestion. Handling animals during the night phase can be more difficult due to this increase in activity.
The diurnal rhythm can be changed by a 12 hour shift in the light cycle. It takes approximately two weeks for rats to adjust to this shift.
Rats are coprophagic (ingest their own feces).

Biological Data

Adult body weight: male	450-520 gm¹
	300-800 gm²
Adult body weight: female	250-320 gm¹
	250-400 gm²
Body surface area	10.5 (wt. in grams)2/3¹
	9.1 kg0.66 B.W.²
Life span	2-3.5 years¹
Food consumption	10 g/100 g/ day¹
Water consumption	10-12 ml/100 g/day¹
Breeding onset: male	65-110 days¹
Breeding onset: female	65-110 days¹
Gestation Period	21-23 days¹
Body Temperature	38-39 C²
	35.9-37.5 C¹
Heart rate	320-480 beats per minute²
	250-450 beats per minute¹
Respiratory rate	85-110 per minute²
	70-115 per minute¹
Tidal volume	0.6-2.0 ml¹
	1.6 (1.5-1.8 ml)²

Reproduction and Sexing

Breeding systems can be monogamous (one male and one female) or polygamous (one male and one to two females).
Gestation is 21-23 days. Litter size varies widely with stock/strain, ranging from 3-18 pups. Birth weights are normally 5-6 grams. Pups are weaned at 21 days of age.
Female rats have a post-partum estrus.
Male and female rats can be differentiated by observing the distance from the anus and genital papilla (anogenital distance) which is greater in males.
This difference is also present in neonatal rats.

Basic Husbandry

Rats can be grouped by their microbial colonization.
- Specific Pathogen Free (SPF) rats are free from known bacterial, viral, and parasitic rat pathogens. To insure the maintenance of the SPF microbial status requires that rats be housed in more stringent conditions that prevent the introduction of rodent pathogens (i.e. maximum isolation housing).
- Conventional rats are not known to be free of pathogens. Standard housing (i.e. conventional housing) cannot prevent the introduction of rodent pathogens
- Other less common categories are axenic rats (i.e free from organisms) and gnotobiotic rats, which have a defined microbial flora.
Choosing the proper housing conditions for rats is very important, especially those used in immunological studies. Rodent pathogens often do not produce clinical signs in affected animals but often have an immunomodulating effects. Conventionally housed animals are exposed to room air and therefore may readily contract viral, bacterial and/or parasitic diseases.
Immunocompromised animals (i.e. athymic rats) are very sensitive to opportunistic agents, and must be housed under maximum isolation conditions to allow for their long term survival. Valuable animals and animals that may need to be transported to other institutions should also be housed under maximum isolation conditions.

Conventional Housing

Most rats are housed in shoebox cages composed of polypropylene (opaque) or polycarbonate material (clear) with a wire bar lid used to hold the water bottle and feed. Bedding is placed directly into the shoe box cage allowing the absorption of urine and the animal to burrow. This type of cage will hold 1-3 adult rats depending on the size of the cage.
When removing the lid from this type of cage, it is important to remove the water bottle to prevent spillage. If the cage is to be transported the bottle should be turned sipper tube up to prevent spillage during transport. However, you should remember to turn the bottle back over to allow access to water after transport.
The husbandry staff changes cages twice per week, thereby providing the animal a clean cage with new bedding, food and water. Water bottles and feed hoppers are checked daily by animal caretakers to insure the provision of food and water.

Maximum Isolation Housing

Rats housed under maximum isolation conditions are housed in sterilized, polycarbonate cages with microisolater bonnets. Bedding is sterilized within the cage and allows the absorption of urine and the opportunity for the animal to burrow. This type of cage will hold 1-3 adult rats depending on the size of the cage.
Animals are provided irradiated food and hyperfiltrated water.
Personnel are required to wear protective clothing (sterile gowns, gloves, mask, head covers and foot covers) while working in the room.
All experimental work and husbandry care must be conducted within a laminar flow hood.
If animals are removed from the housing area and taken to the investigators laboratory, they may not be returned (terminal procedure only).
Access to maximum isolation housing areas is limited to prevent inadvertent fomite transmission of rodent pathogens by personnel.
All research personnel must be trained by the DLAR Assistant Director before working in this area.

Suspended cages & conventional housing

Some rats are housed on wire mesh bottom cages to allow collection of feces and urine or to prevent contact with bedding.
This type of housing is not preferred and is used only when dictated by experimental design (i.e. for toxocilogy studies)
Most suspended cages have water bottles but occasionally are equipped with automatic watering. Typically a nipple valve (lixit valve) is located in the back of the cage which can be operated by contact. When replacing a suspended cage that is provided with automatic watering it is important to push the cage fully into the rack to insure that the lixit valve fully extends into the cage, allowing the animal access to water.
Occasionally a rat will jam the nipple open resulting in constant dripping of water from the nipple, saturating the bedding material. Upon observation the bedding will appear much darker. This problem should be reported to husbandry personnel immediately to allow correction.
Pelleted natural ingredient diets are used to feed all rodents and are composed primarily of cereal grains which are supplemented with additional protein, vitamins and minerals. Due to the nature of this type of diet the exact composition can vary substantially from lot to lot.

Environmental Parameters

Room temperature: 64-79F
Humidity: 30-70%
Ventilation: 10-15 air changes per hour

Identification

Cage cards are utilized to identify the strain of rat, sex, number, principal investigator, and research protocol.
Cage cards should not be removed from the cage to avoid misidentification of the animals.
Temporary identification of individual rats can be accomplished by pen marks on the tail, hair clipping or dyeing the fur. Pen marks will only last 1-2 days whereas hair clipping may last up to 14 days.
Permanent identification methods can be achieved by tail tattooing which will be performed by veterinary personnel upon request. Ear tag or punch identification can be utilized but may be obliterated by fighting between individuals. Subcutaneous microchip transponders are easy to implant and work well but are expensive.
Tail and toe clipping are not recommended.

Handling

When handling rats it is advisable to wear latex gloves to prevent the development of allergies due to direct contact with animal allergens.
Rats typically become accustomed to repeated handling. In a naive animal the temperament of the animal can be determined by placing the hand into the cage to allow exploration by the animal prior to touching. Initial gentle stroking of the animal followed by gradual grasping the animal will prevent startling the animal and initiating an aggressive response.
Avoid approaching the animal from the front.
Rats are normally lifted by grasping the whole body with the palm over the back, with forefinger behind the head and the thumb and second finger under opposite axilla. This extends the rat's forelimbs so that they may be controlled.
Holding with one hand is usually adequate for control, but the tail, rear legs or lower part of body may be held by the other hand for close control, treatment or examination. The use of both hands is often necessary for rats weighing over 350 gms.
Young rats may be handled like mice when body size does not permit ease of handling within the hand.
Investigators should avoid lifting by the tail as they may strip the skin from the tail. This is particularly likely for heavy rats (>450 gms), rats that "spin," and when the tail is grasped more than a couple of centimeters from its base. However, the "base" of the tail may be grasped with the thumb and forefinger. With this simple method of holding, they may be transferred to another cage or a balance, identified, examined casually or sex may be determined.
For transporting short distances it may be helpful to support the rat with your arm or hand while holding the tail.
Rats will bite and certain strains are more aggressive than others (e.g., F344 rats tend to be more aggressive than Sprague-Dawley), so care and experience are essential to rapid handling. Various restraint devices are available for use with rats.
Neonatal rats can be handled from the day of birth but care should be taken to carefully replace the newborn in the nest with the remaining pups.

Restraint Devices

Numerous types of restraint devices are commercially available to restrain rats. Quality devices prevent the animal from turning around yet allow easy access to the tail or legs. Devices should also be easy to clean and provide adequate ventilation.
For tail vein injections in small rats, a wire box cage top can be turned over and the tail gently passed through the wire bars preventing the rat from turning.

Transport

Rats that are moved indoors can be transported in a rat cage. A clean rat cage can be obtained from the cage wash area and one half inch of bedding material added. If you cannot locate a clean cage ask a supervisor or caretaker in the area for assistance.
A wire bar lid should be placed over the cage to secure the animal during transport. If the animal is going to be in the laboratory for more than an hour a water bottle should also be obtained.
During transport the water bottle should be placed upright in the cage lid to prevent spillage.
Transport always results in some stress to the animal, however, animals should recover from indoor transport within their own cage within an hour. No recovery time may be needed if the animals are moved with care and have become accustomed to routine transport.
It is recommended that a permeable drape be placed over the cages to darken the cage and prevent over arousal of the rats during transport.
Only DLAR husbandry or veterinary staff members may transport rats to other buildings or to other animal facilities. Investigators should request animal transportation by contacting the business office at 444-2194. Transport requests should be made 48 hours in advance to allow scheduling of staff.

Environmental Enrichment

Group housing, to allow social interaction, is a primary source of environmental enrichment. All rat cages also have an additional shredded paper material (Enviro-dri ) added to their cage to encourage nest building.
Chewing items such as Nylabones can be used to augment enrichment.
Frequent handling by personnel is important and increases the ease of working with these animals.
If animals cannot have environmental enrichment opportunities because of the nature of the research please contact the DLAR Director or the Clinical Veterinarian (444-2194).

Occupational Health/Zoonosis

Zoonoses are infectious diseases that can be passed from animals to humans.
Potential zoonotic diseases are associated with rats.
- Rat Bite Fever is caused by one of two bacteria, Streptobacillus moniliformis or Spirillium minus. Transmission of these agents is by a bite wound.
- Tapeworms (Hymenolepis nana) are transmitted by the fecal-oral route.
- Ringworm, caused by Trichophyton mentagrophytes, is a fungal infection of the skin and is transmitted by physical contact.
Personnel can receive severe bites from rats and these bites can easily become infected. As always, it is important to clean the wound promptly and seek medical attention, when appropriate.
A tetanus vaccination is required for all personnel working with animals.
Allergy to rat dander and urine is not uncommon and usually develops within a few years of working with the animals. Sensitive personnel should wear face masks and/or respirators (properly fitted), gloves and a lab coat. Personnel should alert Occupational Medicine during their annual health risk assessment and if allergy is a problem, obtain advice and treatment from the Occupational Medicine physician.

Health Surveillance

A. Monitoring

Investigators should monitor their animals routinely for common signs of illness such as
- loss of appetite,
- weight loss,
- diarrhea,
- nasal discharge,
- ocular discharge (brownish-red in color; known as poryphyria),,
- lethargy,
- swelling around the jaw,
- vocalization (chattering of teeth),
- unkempt appearance, and
- subcutaneous masses
The frequency of monitoring of the animal by the investigator is stated in their approved IACUC protocol. Early endpoints are also defined in the protocol and the investigator is responsible for euthanize their animals when these early endpoints are reached.

B. Reporting Sick Animals

If an animal is identified with either experimental or non-experimentally related illness, the investigator should notify the DLAR veterinary staff at 444-2194 between the hours of 8:00 a.m. and 5:00 p.m., Monday through Friday.
If the investigator needs to contact a veterinarian on the weekend, the veterinary on-call list is posted in the main entry area of all DLAR facilities and provides the name, home phone number and pager number of the veterinarian on duty.
When contacting the veterinarian, please provide the following information:
- Investigators name/ your name
- Species of animals and animal ID number
- Location of the animal (building, room #, rack or cage #)
- Signs of illness
- Description of any experimental manipulations performed
- Phone number where you can be reached.
In emergency situations, if an animal needs immediate veterinary care, call the main DLAR office at 444-2194. The DLAR office will send an emergency (911) page or radio the veterinarian. If the emergency occurs after hours, call the veterinarian directly.

References

Harkness JE, Wagner JE.The Biology and Medicine of Rabbits and Rodents; 3rd Edition, Lea and Febiger, Philadelphia, PA, 1989.
Poole TB.The UFAW Handbook on the Care and Management of Laboratory Animals, 6th edition, Longman Scientific & Technical; England, 1987.
Sharp PE, La Regina MC. The Laboratory Rat, CRC Press, Boca Raton, NY, 1998.

Biomethodology of the Guinea Pig

Environmental Enrichment

Occupational Health /Zoonosis

Health Surveillance

References

General Biology:

Nomenclature and Breeds

The guinea pig, Cavia porcellus , is a mammal of the order Rodentia, sub-order Hystricomorpha ("porcupine-like") and family Caviidae.
Three basic breeds of guinea pigs exist. The English (short-haired), Peruvian (long-haired) and Abyssinian which has a rosette hair pattern. Pigmented guinea pigs of all three breeds are also available.
Most guinea pigs used in research are "outbred" animals of the various breeds. The common Dunkin Hartley guinea pig (future picture) is an albino outbred guinea pig of the English (short-haired) breed.
Several "inbred" guinea pig strains are also available. The "strain 2" and "strain 13" guinea pigs are the most widely used inbred guinea pig strains.

Unique Biological Characteristics

Guinea pigs are herbivores and unlike most laboratory animals, except non human primates, they require a nutritional source of Vitamin C.
Kurloff cells are unique to the guinea pig. The cell is a mononuclear leukocyte that has round to ovoid inclusions. While the origin and function of the cell is not known, their numbers increase with higher levels of estrogen and the cells migrate to the placenta in pregnant guinea pigs.
Guinea pigs have a cervical thymus making it easy to access for experimental manipulations.
Guinea pigs are considered to be a steroid resistant species.
The ECG of guinea pigs resembles that of humans with easily identifiable P,Q,R,S and T waves. Their T wave is distinctly separate from the QRS complex.
The majority of pulmonary stretch receptors in guinea pigs are located in small airways and in pulmonary parenchyma

Experimental Uses

Anaphylaxis
Histamine sensitivity in acute bronchospasm
Reproductive biology
Nutrition studies
Tuberculosis

Routine Technical Procedures

Blood Collection Sites
large volumes cranial vena cava (terminal only)
moderate volumes jugular vein, saphenous vein, dorsolateral penile vein, retro-orbital
small amount clipping toenail short

Sources and Ordering

Inbred and outbred guinea pigs are available from many commercial sources. The DLAR office staff is familiar with many of the commonly available strains and will assist in locating animals available for purchase.
Commercially bred guinea pigs from Charles River must be ordered by Tuesday at 9:00 a.m. for delivery on Thursday of that week. Animals should be held for 48 hours before experimental use to allow recovery from the stress of shipping.

Behavior

Guinea pigs are very docile and rapidly become accustomed to gentle handling.
Guinea pigs rarely bite. Aggression between females is uncommon and is more likely to occur between males in competition for a female in estrous.
Fighting is rare - even between males.
Guinea pigs are easily alarmed and will often "freeze" for extended periods (30 minutes) when startled. Group housed guinea pigs may stampede when startled which may result in injury to young guinea pigs, orthopedic injuries, and abortion in pregnant dams.
The primary interaction between guinea pigs is "huddling". Social grooming is rare. Dominant animals may "barber", or chew/clip the fur of subordinate animals.
Guinea pigs are considered crepuscular (most active in "twilight" hours) animals.
When placed in a comfortable environment, guinea pigs may be active for more than 20 hours per day.
Guinea pigs will eat as long as food is available.

Biological Data

Adult body weight: male	800-1200 gm¹
Adult body weight: female	250-320 gm¹
Body surface area	9.5 (wt. in grams)2/3¹
Life Span	4-5 years²
Food consumption	6 g/100 g/ day¹
Water consumption	10 ml/100 g/day¹
Puberty: male	8-10 weeks³
Puberty: female	67.8 + (21.5 SD)days³
Gestation Period	65-72 days²
Body Temperature	37.2-39.5 C¹
Heart rate	230-380 beats per minute¹
Respiratory Rate	42-104 per minute¹
Tidal volume	2.3-5.3 ml/kg¹
Blood Volume	67-92 ml/kg⁴

Basic Husbandry

Specific Pathogen Free (SPF) guinea pigs are free from certain bacterial, viral, and parasitic pathogens. To maintain the SPF microbial status animals must be housed in more stringent conditions that prevent the introduction of other pathogens. Examples of housing include isolation in separate rooms or maximum isolation housing.
Conventional guinea pigs are not known to be free of pathogens.
Most guinea pigs are gang housed in large pens. The pens have hardwood chip bedding on the bottom to collect urine and feces. Water bottles and feeders are affixed to the sides of the pens and are elevated. If feeders are not elevated, guinea pigs will sit in the feeders and urinate and defecate in them, contaminating the feed.
Some guinea pigs are singly or group housed in stainless steel cages on raised slatted floors. Wire flooring is not recommended because it can cause pododermatitis ("sore hock") which is inflammation and infection of the foot pads. This condition occurs because guinea pigs are relatively heavy in proportion to the size of their feet.
The husbandry staff change cages twice per week. Water bottles and feed hoppers are checked daily by the caretakers to insure the provision of water.
Some guinea pigs learn to play with the water bottle and will drain the bottle. Investigators should check the water bottles when they are in the room and bring any empty or low water bottles to the attention of the husbandry staff.
Feed is provided daily. Pelleted natural ingredient diets are used to feed all rodents and are composed primarily of cereal grains which are supplemented with additional protein, vitamins and minerals. Due to the nature of this type of diet the exact composition can vary substantially from each vendor.
Guinea pigs are one of the few mammals other than primates that require a nutritional source of Vitamin C. For that reason guinea pig chow is has a shorter shelf life (90 days) than standard rodent chow and is manufactured specifically for guinea pigs.

Housing parameters:

Room temperature: 64-70 F
Humidity: 30-70%
Ventilation: 10-15 air changes per hour

Identification

Cage cards are utilized to identify the strain of guinea pig, sex, number, principal investigator, and IACUC protocol number.
Cage cards should not be removed from the cage/pen to avoid misidentification of the animals.
Temporary identification of individual animals can be accomplished by dyeing the fur or clipping the hair. Various dyes such as trypan blue, picric acid, fuschein or methyl violet can be utilized. This form of identification will last only 1-2 weeks.
Permanent forms of identification can be achieved by the use of ear tags, ear punch or ear notch. Fighting between cage mates will result in the occasional loss of an ear tag.
Toe clipping is not a recommended form of identification.
Colored guinea pigs can be individually identified by noting the pattern of coloration.

Handling

When handling guinea pigs it is advisable to wear latex gloves to prevent the development of allergies.
Guinea pigs seldom bite but are timid or easily frightened and usually make determined efforts to escape when held. Guinea pigs typically become accustomed to repeated handling.
To pick up a guinea pig one hand should be gently placed dorsally over the thorax or ventrally under the thorax and the other hand should be used to support the animals hindquarters. Care should be taken not to apply to much pressure over the thorax to avoid damaging the viscera or compressing the lungs thereby compromising respiration.
Special care should be exercised in supporting the lower part of the body of pregnant females since they may become very heavy and awkward in late pregnancy.
After grasping the guinea pig secure the animal by wrapping it in a towel or holding it against your body to decrease struggling.
Do not attempt restraint by solely grasping the skin. The lack of loose skin in guinea pigs will result in hair depilation if this technique is utilized.
Neonatal guinea pigs can be handled from the day of birth.

Sexing and Reproduction

A female guinea pig is called a sow. A male guinea pig is called a boar.
Male and female guinea pigs can be differentiated by palpating the penis or extruding the penis of the male by gently applying pressure above the urethral orifice. This technique will expose the vaginal membrane, which in females closes the vagina unless the guinea pigs is in estrous or about to deliver young.
The anogenital distance is similar in males and females and cannot be used for sex determination.
Gestation in guinea pigs is 58-75 days. Labor is short. Parturition may occur at any time of the day.
Young are precocial (eyes open, fully haired) and will usually be walking, eating and drinking by one day of age.
The young are weaned at 3 weeks of age.

Transport

Guinea pigs that are moved indoors can be transported in a rat cage. A clean rat cage can be obtained from the cage wash area and filled with one half inch of bedding material. If you cannot locate a clean cage, ask a supervisor or caretaker in the area for assistance.
A wire bar lid should be placed over the cage to secure the animal during transport. If the animal is going to be in the laboratory for more than an hour a water bottle should also be obtained.
During transport the water bottle should be placed upright in the cage lid to prevent spillage.
Transport always results in some stress to the animal, however, animals should recover from indoor transport within their own cage within an hour. No recovery time may be needed if the animals are moved with care and have become accustomed to routine transport.
It is recommended that a permeable drape be placed over the cages to darken the cage and prevent over-arousal of the guinea pigs during transport.
Only DLAR husbandry or veterinary staff members can transport guinea pigs to other buildings or to other animal facilities. Investigators can request animal transportation by contacting the business office at 444-2194. Transport requests should be made 48 hours in advance to allow scheduling of staff.

Environmental Enrichment Opportunities:

Group housing is the main form of housing for guinea pigs in DLAR. This allows social interaction.
Handling by personnel is important and increases the ease of working with these animals.
Food/chewing items can be made available such as wood chew sticks and vegetable treats.
If animals cannot have environmental enrichment opportunities because of the nature of the research, please contact the DLAR Director or the Clinical Veterinarian (444-2194).

Occupational Health /Zoonosis:

Occasionally personnel may be scratched by a toenail when handling the animal. Personnel may also receive a puncture wound from the cage equipment. As always, it is important to clean the wound promptly and seek medical attention if appropriate.
A tetanus vaccination is required for all personnel working with animals.
Allergy to guinea pig dander is not uncommon. Sensitive personnel should wear face masks and/or respirators (properly fitted), gloves and a lab coat. Personnel should alert Occupational Medicine during their annual health risk assessment and if allergy is a problem, obtain advice and treatment from the Occupational Medicine physician.

Health Surveillance

A. Monitoring

Investigators should monitor their animals routinely for common signs of illness, such as:
- loss of appetite
- weight loss
- diarrhea
- nasal or ocular discharge
- lethargy
- unkempt appearance
The frequency of monitoring of the animal by the investigator is stated in their approved IACUC protocol. Early endpoints are also defined in the protocol and the investigator is responsible for euthanize their animals when these early endpoints are reached.

B. Reporting Sick Animals

Research personnel are responsible for monitoring their experimental animals as approved in their IACUC protocols.
If an animal is identified with either experimental or non-experimentally related illness, the investigator should notify the DLAR veterinary staff at 444-2194 between the hours of 8:00 a.m. and 5:00 p.m., Monday through Friday.
If the investigator needs to contact a veterinarian on the weekend, the veterinary on-call list is posted in the main entry area of all DLAR facilities and provides the name, home phone number and pager number of the veterinarian on duty.
When contacting the veterinarian, please provide the following information:
- Investigators name/ your name
- Species of animals and animal ID number
- Location of the animal (building, room #, rack or cage #)
- Signs of illness
- Description of any experimental manipulations performed
- Phone number where you can be reached
In emergency situations, if an animal needs immediate veterinary care, call the main DLAR office at 444-2194. The DLAR office will send an emergency (911) page or radio the veterinarian. If the emergency occurs after hours, call the veterinarian directly as described above.

References

1) Harkness JE, Wagner JE. The Biology and Medicine of Rabbits and Rodents ; 3rd Edition, Lea and Febiger, Philadelphia, PA, 1989.

2) The UFAW Handbook on the Care & Management of Laboratory Animals, 6th Edition; Editor Poole, TB. Longman Scientific & Technical, England, 1986.

3) Wagner JE, Manning PJ. The Biology of the Guinea Pig; Academic Press, 1976.

4) Joint Working Group on Refinements. Removal of blood from laboratory animals and birds. Lab. Anim. 27:1-22,1993.

5) Assistant Laboratory Animal Technician Manual, American Association for Laboratory Animal Science, Memphis, TN, pgs. 119-120, 1999.

6) Terril LA, Clemons DJ. The Laboratory Guinea Pig, CRC Press, Boca Raton, NY, 1998.

Biomethodology of the Rabbit

General Biology

Experimental Uses

Routine Technical Procedures

Sexing and Reproduction

Transport

Environmental Enrichment

Health Surveillance

References

General Biology

A. Nomenclature and Breeds

Of the many breeds of the domesticated European rabbit (Oryctolagus cuniculus), the albino New Zealand White is the most common breed utilized in biomedical research.
Non-albino breeds, such as the black and white Dutch belted rabbit, tend to be preferred where pigmentation is required (i.e. ophthalmological research).

B. Unique Biological Characteristics

The rabbit“s skeleton comprises only 7% of their total body weight (versus the cat skeleton which is 13%). The light skeleton leads to a greater opportunity for spontaneous vertebral fractures (broken backs) due to kicking of the rabbits strong rear legs if they are restrained improperly.
The "dewlap" is a large fold of skin under the chin and is most prominent in female rabbits.
Rabbits have hyposegmented neutrophils called Pelger-Huet cells.
The long slender ears and visibility of the peripheral vasculature in albino rabbits is advantageous for blood collection.
Rabbits like rodents have two large upper incisors and two large lower incisors. Unlike rodents, however, an additional pair of incisors(peg teeth) are located caudal to the large upper incisors. This additional pair of upper incisors is greatly reduced in size and is why rabbits are members of the order Lagomorpha rather than the order Rodentia.
Rabbit teeth continue to grow throughout life. Malocclusion occasionally occurs in rabbits preventing normal tooth wear. This results in severe overgrow of the teeth by inhibiting normal mastication.
The neutrophil of the rabbit resembles an eosinophil, due to the numerous intracytoplasmic eosinophilic granules, and are called pseudophils.
Numerous genetic mutations have been noted in the rabbit2 and several inbred rabbit strains have been produced.

Experimental Uses

Polyclonal antibody production
Orthopedics
Ophthamology
Infectious disease
Atherosclerosis
Pyrogen testing
Draize testing

Routine Technical Procedures

A. Blood Collection sites

large volume — intracardiac (terminal procedure only)
moderate volume—marginal ear vein, lateral thoracic vein, central ear artery
small volume—lateral metatarsal vein, toe nail clip

B. Polyclonal Antibody Production

Sources and Ordering

Conventional rabbits are readily available from several commercial sources. SPF rabbits and genetically unique animals are not as readily available and may take longer to acquire.
For routine orders, commercially bred rabbits from Charles River animals must be ordered by Tuesday at 9:00 a.m. for delivery by Thursday of that week.
A source list of mutant and inbred rabbits is available from the Institute of Laboratory Animal Resources.3
Conventional rabbits will be entered into a conditioning program on arrival and will be held for 7 days before experimental use to allow for prophylactic treatment for coccidia and to allow for recovery from the stress of shipping.

Behavior

Rabbits are gentle animals and, if care is taken, become accustomed to handling if they are not upset by the experience. Inappropriate handling can result in severe scratches to the handler from the toe nails on the rabbits powerful rear legs.
Rabbits are herbivores and have the unique characteristic of re-ingesting soft fecal pellets ("night feces") directly from the anus. This process of "pseudorumination" allows the acquisition of B vitamins that have been produced by microorganisms in the cecum.
Defensive behavior in rabbits includes thumping on cage floor with rear feet. Some aggressive rabbits will also charge the front of the cage.

Basic Biological Data

Adult body weight: male	2-5 kg¹
Adult body weight: female	2-6 kg¹
Life span	5-6 up to 15 years¹
Food consumption	5 g/100 g/ day¹
Water consumption	5-10 ml/100 g/day⁶
Rectal Temperature	38.5-39.5C4
Heart rate	205-235/minute⁶
Respiratory rate	30-60 per minute¹

Basic Husbandry

Rabbits can be grouped by their microbial colonization and both are available for purchase.
- Specific Pathogen Free (SPF) rabbits are free from several common pathogenic organisms such as Pasteurella, Encephalitzoonois, and Coccidiosis and are bred specifically for research use.
- Conventional rabbits tend to be colonized with the preceding organisms which can occasionally be detrimental to their health and longevity. Conventional rabbits are bred primarily for food and fur.
- Unlike SPF rabbits, conventional rabbits are widely available and if appropriate for the project can represent a substantial savings upon purchase. The investigator must weigh these considerations when choosing the type of rabbit for purchase.
Most rabbits are housed in stainless steel cages with a fenestrated floor to allow feces to drop through into a pan. Absorbent material is placed in the pan to collect urine and minimize ammonia release due to the bacterial breakdown of urea.
Caged rabbits are typically housed individually, however, some cages allow for pair housing. Rabbits less than 2 kg in weight are provided with 1.5 square feet of floor space. Rabbits weighing from 2-4 kg are maintained in 3 square foot cages.
Rabbits may also be gang housed in a large room, directly on bedding. Bedding material should be placed over cardboard liners to prevent damage to the underlying floor from urine. Care should be taken in selecting bedding materials—the ink on shredded newspaper can discolor the rabbits fur, and straw, which is dusty, can cause upper respiratory problems and conjunctivitis. A shredded, white paper material, such as Enviro-dri™, has proved to be efficacious.
Water is typically provided through either a "lixit valve" that provides water at all times or by a 1 liter bottle. The “lixit valve” on cages must be checked frequently to make sure it has not become plugged. The water bottle is attached either to the front of the cage or pen. Food is provided by an elevated “J” hopper also attached to the front of the cage or pen. Use of the “J” hopper prevents the rabbit from defecating in their food, which can occur with the use of food crocks (bowls). Water lixit valves and feed hoppers are checked daily by caretakers to insure the provision of food and water.
The drop pans are changed by the husbandry staff twice per week, and the rabbit is placed into a new sanitized cage every two weeks. Bedding in gang-housed areas is changed once weekly.
Pelleted natural ingredient diets are used to feed all rabbits and are composed primarily of cereal grains which are supplemented with additional protein, vitamins and mineral. Rabbits receive a high fiber diet which tends to minimize common gastrointestinal disorders. Due to the nature of natural ingredient diets, the exact composition can vary substantially from each vendor. Rabbits are fed a specified amount of diet daily in order to prevent obesity which tends to occur if rabbits are fed ad libitum. Newly received rabbits are placed on an incremental feeding regimen over a 4 day period to acclimate them to the new feed and prevent diarrhea.
Rabbits are provided with a 12 hours of light each day.

Housing parameters:

Room temperature: 64-70 F
Humidity: 40-70%
Ventilation: 10-15 air changes per hour

Identification

Cage cards are utilized to identify the strain of rabbit, sex, number, principal investigator, and research protocol.
Cage cards should not be removed from the cage to avoid misidentification of the animals.
Temporary identification of individual rabbits can be accomplished by pen marks on the fur, or dyeing the fur. Pen marks or dyes will only last 1-2 days.
Permanent identification methods via ear tattooing is commonly performed by the vendor before the rabbits are shipped to the facility (add picture).

Handling

When handling rabbits it is advisable to wear latex gloves to prevent the development of allergies and to provide some safety from rabbit scratches to the handler. Rabbits seldom bite but can inflict painful scratch wounds, especially with the hind feet.
Grasping the loose skin over the neck and shoulder, with the head directed away from the holder, is the best method of initial restraint. When lifting a rabbit, the lower part of the body must be supported by the other hand to prevent serious injury to the rabbit's back. If the rabbit begins to struggle, it should immediately be placed on a solid surface and calmed. Struggling frequently leads to fracture of lumbar vertebrae and injury to the spinal cord that may necessitate euthanasia.
Rabbits should never be restrained or lifted by the ears.
If properly used, commercially available rabbit restrainers help to avoid injuries.

Restraint Devices

Several types of devices are commercially available to restrain rabbits. Quality devices prevent the animal from turning around or twisting yet allow easy access to the head and ears.
Care should be taken when placing a rabbit in a restraint device since struggling may result in damage to the spine.
Struggling is reduced if the device snugly secures both the head, back and hind legs.
Rabbits should never be left unattended in restraint devices. Training in the use of these devices can be arranged through the DLAR veterinary staff.

Sexing and Reproduction

A male rabbit is called a buck. A female rabbit is called a doe.
Rabbits can be sexed by causing eversion of the penis or vulva when slight pressure is applied to the external genitalia.7
Rabbits are induced ovulators and can have pseudopregnancies.
Gestation in the rabbit is 29-35 days.
Parturition in rabbits is known as kindling.
Rabbit pups are born fully furred with their eyes closed.
Pups are weaned at 28 days of age.
Pups are normally nursed only once a day by the doe.

Transport

Rabbits that are moved indoors can be transported in a cat carrier. A paper liner should be placed in on the bottom of the carrier to catch urine and feces. Cat carriers can be obtained from the DLAR veterinary staff.
Transport always results in some stress to the animal, however, rabbits should recover from indoor transport within their own cage in an hour. No recovery time may be needed if the animals are moved with care and have become accustomed to routine transport.
Only DLAR husbandry or veterinary staff members can transport rabbits to other buildings or to other animal facilities. Investigators can request animal transportation by contacting the business office at 444-2194. Transport requests should be made 48 hours in advance to allow scheduling of staff.

Environmental Enrichment

Group housing is the main form of housing for rabbits in DLAR. This allows for social interaction.
Frequent handling by personnel is important and increases the ease of working with these animals.
Food/chewing items can be made available as rabbit enrichment including wood chew sticks and vegetable treats.
Rabbits enjoy enclosed areas to use as "burrows" such as large bore plastic tubes.
If animals cannot have environmental enrichment opportunities because of the nature of the research, please contact the DLAR Director or the Clinical Veterinarian.
Personnel can receive severe scratches by the rabbit’s hind feet. These scratches can easily become infected. As always, it is important to clean the wound promptly and seek medical attention, when appropriate.
A tetanus vaccination is required for all personnel working with animals.
Allergy to rabbit dander is not uncommon. Sensitive personnel should wear face masks and/or respirators (properly fitted), gloves and a lab coat. Personnel should alert Occupational Medicine during their annual health risk assessment and if allergy is a problem, obtain advice and treatment from the Occupational Medicine physician.

Health Surveillance

A. Monitoring

Investigators should monitor their animals routinely for common signs of illness, such as:
- loss of appetite
- weight loss
- diarrhea
- nasal or ocular discharge
- lethargy
- unkempt appearance
The frequency of monitoring of the animal by the investigator is stated in their approved IACUC protocol. Early endpoints are also defined in the protocol and the investigator is responsible for euthanize their animals when these early endpoints are reached.

B. Reporting Sick Animals

Research personnel are responsible for monitoring their experimental animals as approved in their IACUC protocols.
If an animal is identified with either experimental or non-experimentally related illness, the investigator should notify the DLAR veterinary staff at 444-2194 between the hours of 8:00 a.m. and 5:00 p.m., Monday through Friday.
If the investigator needs to contact a veterinarian on the weekend, the veterinary “on-call” list is posted in the main entry area of all DLAR facilities and provides the name, home phone number and pager number of the veterinarian on duty.
When contacting the veterinarian, please provide the following information:
- Investigators name/ your name
- Species of animals and animal ID number
- Location of the animal (building, room #, rack or cage #)
- Signs of illness
- Description of the experimental manipulations performed, if any.
- Phone number where you can be reached.
In emergency situations, if an animal needs immediate veterinary care, call the main DLAR office at 444-2194. The DLAR office will send an emergency (911) page or radio the veterinarian. If the emergency occurs after hours, call the veterinarian directly as described above.

References

Harkness JE, Wagner JE. The Biology and Medicine of Rabbits and Rodents, 3rd Edition, Lea and Febiger, Philadelphia, PA,1989.
Manning PJ, Ringer DH and Newcomer CE.The Biology of the Laboratory Rabbit, Academic Press, New York, NY, 1994.
Institute of Laboratory Animal Resources, National Academy of Sciences, National Research Council, 2101 Constitution Avenue, Washington, DC.
Ruckebusch,Y, Phaneuf LP, and Dunlop R, Physiology of Small and Large Animals. Editor Dekker, Philadelphia, PA, 1991.
The UFAW Handbook on the Care & Management of Laboratory Animals, 6th Edition; Editor Poole TB, Longman Scientific & Technical, England, 1986.
Fox RR, Crary DD. A simple technique for the sexing of newborn rabbits. Lab. Anim. Sci . 22:556-8, 1972.
Assistant Laboratory Animal Technician Manual, American Association for Laboratory Animal Science, Bookcrafters, Chelsea, MI, pg. 125-128, 1999.
Suckow MA, Douglas FA. The Laboratory Rabbit, CRC Press, Boca Raton, NY, 1997.

Rodent Survival Surgery

Introduction

The Surgical Area

Preparation of Surgical Instruments and Supplies

Preoperative Considerations/Care

Animal Preparation

Surgeon Preparation

Maintaining a Sterile Field

Monitoring anesthesia

Multiple Surgeries in a Single Session

Anesthesia Recovery

Post-Operative Care

References

Introduction

Post-operative infections in rodents can and do occur. The misconception that rodents have an innate resistance to bacterial infection has not been scientifically substantiated. Such infections, which may not be apparent on casual observation, will cause loss of vessel cannulations1, and numerous changes in physiologic parameters2 . In accordance with good scientific practice and standards set forth in the Public Health Service Guide for the Care and Use of Laboratory Animals (Guide) and the Federal Animal Welfare Act, aseptic surgical procedures must be used.

The NIH Guide provides the following guidelines for rodent surgery:

Major survival surgery on rodents does not require a special facility but should be performed using sterile instruments, surgical gloves, and aseptic procedures to prevent clinical infections.
Major survival surgery is defined as any surgical intervention that penetrates a body cavity or has the potential for producing a permanent handicap in an animal that is expected to recover.
Training in aseptic procedures should be provided for those who require it.

The Animal Welfare Act states:

“Non-major operative procedures and all surgery on rodents do not require a dedicated facility but must be performed using aseptic procedures. Operative procedures conducted at field sites need not be performed in dedicated facilities but must be performed using aseptic procedures.”

The Surgical Area

A separate room used primarily for aseptic procedures is desirable; however, it is appropriate to perform survival rodent surgical procedures in a conventional laboratory setting using aseptic technique. The investigator should adopt the following standards for aseptic procedures:

A clean, uncluttered work area and a sanitized work surface should be utilized for the surgery area.
The work area should be located to minimize laboratory traffic not related to the surgical procedure and dedicated exclusively for surgery, when in use.
Considerations should also be given to locate the surgery away from potential sources of contamination such as open windows, fans, or fume hoods which can blow dust into the area and increase desiccation of exposed tissues.
The surgery surface should be disinfected (70% alcohol or a quaternary ammonium compound) before use. It often simplifies the maintenance of asepsis if a sterile drape is then applied over the surgery surface.

Preparation of Surgical Instruments and Supplies

Survival surgical procedures on all mammalian species must be conducted using aseptic technique which requires the use of sterile instruments, supplies and wound closure materials (suture, wound clips).
Many supplies such as gloves, surgical blades, and suture material are commercially available as sterile packs. However, it is frequently necessary to sterilize, in house, items such as surgical instruments, drapes, gowns, etc.
Sterilization kills all viable microorganisms while disinfection only reduces the number of viable microorganisms. High level disinfection will not kill the more resistant bacterial spores. Commonly used disinfectants such as alcohol, iodophors, quaternary ammonium and phenolic compounds are not effective sterilants and, therefore, are not acceptable for use on items intended to be used in survival surgical procedures.
The following are approved sterilization procedures:
- High pressure/temperature steam sterilization using an autoclave and appropriate monitoring systems to assure sterility.
- High temperature dry heat systems. Since it is difficult to drape instruments prepared in this fashion they cannot be stored for future use. Typically instruments are sterilized and allowed to cool immediately prior to use by the surgeon.
- Gas sterilization with ethylene oxide using an appropriate gas sterilizer and appropriate monitoring systems to assure sterility and personnel safety.
- Cold (chemical) sterilization
- Alcohol with flaming (dulls instruments)
Effective and proper use of cold sterilization is dependent on many factors including:
- The sole use of chemicals classified as "sterilants". Those classified only as disinfectants (70% alcohol) are not adequate.
- Instruments must be relatively smooth, impervious to moisture, and be of a shape that permits all surfaces to be exposed to the sterilant. Instruments tend to degrade when exposed to sterilants which requires that their integrity be assured prior to each use.
- All surfaces, both interior and exterior, must be exposed to the sterilant. Tubing must be completely filled and the materials to be sterilized must be clean and arranged in the sterilant to assure total immersion.
- The items being sterilized must be exposed to the sterilant for the prescribed period of time.
- The sterilant solution must be clean and fresh. Most sterilants come in solutions consisting of two parts that when added together form what is referred to as an "activated" solution. The shelf life of activated solutions is indicated on the instructions for commercial products.
- Instruments, implants, and tubing (both inside and out) should be rinsed with sterile saline or sterile water prior to use to avoid tissue damage.
- Examples of commercially available sterilants:
  - Cidex®- active ingredient: 2% glutaraldehyde, requires many hours of immersion for effective sterilization
  - Clidox®- active ingredient: chlorine dioxide, minimum of 6 hours required for sterilization
Instruments used in pediatric or ophthalmic surgery are sized appropriately for rodent surgery3. These tend to be delicate instruments and the user should examine them prior to each sterilization to insure their integrity.

Preoperative Considerations/Care

Animal Health/Selection
- A healthy rodent is a prerequisite to a successful surgery. Rodents undergoing clinical or sub-clinical disease often experience anesthetic complications and are not good candidates for a successful procedure.
- It is recommended that if animals are specifically purchased for surgery that they are housed under maximum isolation conditions on arrival to insure the absence of rodent diseases.
- A minimum of 48 hours is generally required for an animal to recover from the stress of shipping; therefore, surgery should not be performed until this holding period has been completed.
Preoperative withholding of food
- It is common in larger species to withhold food prior to surgery to prevent the possibility of aspiration pneumonia after regurgitation. This practice is not necessary in rodents since they cannot vomit.
- Fasting for four hours before surgery, however, may promote the absorption of intraperitoneally administered anesthetics .
- Water should never be withheld.
Pre-surgical Medication
- To decrease tracheobronchial secretions, which may cause obstruction of the trachea, the use of atropine or glycopyrrolate should be considered.
- The investigator should be prepared to aspirate secretions from the trachea if necessary.
Pre/Postoperative antibiotics
- If proper aseptic technique is utilized, antibiotics should not be necessary.
- Antibiotics, in fact, are contraindicated in hamsters and guinea pigs due to the frequent development of fatal Clostridial enteritis.
- If the interior of the intestinal tract is exposed, antibiotics are commonly administered.
- To have the desired effect, antibiotics should be administered prior to surgery to provide adequate blood/tissue levels at the time of surgery.

Animal Preparation

Animal preparation includes preparation of the surgical site by removal of the fur by clipping, plucking or using a depilatory. An area approximately 15% larger than the area of the incision should be prepared.
A vacuum may be used to remove the majority of the fur removed and the use of an adhesive pad will often remove any extraneous fur from the surgical site.
Clean and aseptically prepare the surgical site by using an appropriate scrubbing technique (e.g. scrubbing in gradually enlarging circular pattern from the interior of the shaved area to the exterior) and an effective disinfectant (e.g. alternating Betadine™ or Nolvasan™ and alcohol scrubs X 3).
The disinfectant should be in contact for a minimum of 3 minutes before the initial incision is made.
Do not saturate other areas of the body with disinfectant since this enhances hypothermia which is a common postoperative complication in rodents.
The surgical area should be draped with sterile drapes to avoid contamination of the incision, instruments and supplies. Opaque and non-opaque materials can be used. Clear materials have the advantage of allowing the investigator to monitor respiration and perfusion through the drape. Autoclavable plastic and sterile adhesive dressings (Steri-Drapes™) are available for use.
- It is recommended that animals be placed on a water re-circulating heating blanket or pads during surgery to prevent hypothermia.
- It is recommended that ophthalmic ointment be placed in the anesthetized animals’ eyes to prevent drying of the cornea.

Surgeon Preparation

The surgeon and surgical assistant must wear a clean lab coat, mask and sterile gloves. A sterile surgical gown and head cover are recommended for major or prolonged surgeries.
The surgeon must perform a thorough surgical scrub, utilizing an appropriate disinfectant, before putting gloves on.

Maintaining a Sterile Field

The surgeon should restrict his/her contact to the surgical site and previously sterilized equipment until the incision is closed.
Only sterile solutions should be used and the tops of containers should be disinfected before use.
Use a sterilized area (surgical tray or sterile gauze) to rest materials such as catheters, implants, etc. on when not in use to avoid contamination.
Manipulation of tissues within the surgical field with gloved hands should be avoided; the ends of sterilized instruments should be used to manipulate and handle tissues.
The exteriorizing of organs should be avoided if possible, but if required, the organs should be placed on the sterile drape.
Gloves must be changed if they come in contact with a non-sterile surface.
Two separate sets of surgical instruments can be used —one set for incising and manipulating skin and another for manipulating deeper tissues.

Monitoring anesthesia

The small size of rodents precludes the use of several common methods to evaluate anesthesia used in larger species.
For rodents, periodic observation of respiration, color of mucous membranes and loss of reflected eye color (in albino animals) will provide the surgeon with a good assessment of the animal's status.
Except for guinea pigs, the absence of the pedal reflex is a good indication that a surgical plane of anesthesia has been attained in rodents.
The absence of the pinna reflex is a good indicator in guinea pigs.

Multiple Surgeries in a Single Session

It may be necessary to surgically prepare several different animals during one session using one sterile pack. This is appropriate, providing care is taken to maintain sterility of the instruments.
Preparation of animals, surgical materials and surgical area must be completed prior to putting on sterile gloves.
Instruments and gloves may be used for a series of similar surgeries provided they are used appropriately and disinfected between use.
- It may be appropriate to segregate instruments based on potential for contamination. For example, the instruments used to incise the skin could be dedicated solely for that purpose and separate instruments utilized to manipulate exposed tissues and organs.
- Manipulate the tissues with only the tips of the instruments and avoid handling the tissues directly with your hands, which tend to be more easily contaminated.
- Using a dry bead sterilizer to re-sterilize the tips of instruments between surgeries will further insure adequate aseptic technique if precautions are taken to allow cooling of the instruments before re-use.
- Alternating 2 sets of sterile instruments provide the necessary time for instruments to sit in disinfectant for the required time. (N.B. immersion times can be long (up to 6 hours) which may preclude this as a viable option). Instruments must be thoroughly rinsed with sterile saline prior to re-use.
Consider using a separate sterile pack for no more than five animals.
For major surgeries, (i.e. for surgeries that enter a major body cavity), it is highly recommended that sterile gloves be changed between animals.

Anesthesia Recovery

After surgery the animal should be placed back in a cage that is lined with an absorbent pad. Animal bedding should not be present since the unconscious animal may aspirate bedding into the nares thereby compromising respiration.
The most common complication that occurs during and after surgery is the development of hypothermia. This is often exacerbated by performing surgery directly on a heat conducting surface (stainless steel). This can be avoided by using sterile pads under the animal or utilizing a circulating water pad. Electrical heating pads cannot be appropriately regulated and should never be utilized as a heat source because they can cause severe thermal burns.
After surgery an incandescent lamp (50-75 watt) can be placed 12-14 inches away from the animal to provide supplemental heat. The lamp should be positioned so that the animal can escape the light source if desired.
Many procedures entail the loss of body fluids either through bleeding or drying during surgery. In those cases the administration of warmed sterile saline either subcutaneously or intraperitoneally will hasten the animal's recovery.
Animals recovering from anesthesia should be monitored and rotated from side to side every 15 minutes until they are able to maintain sternal recumbancy. After full recovery, animals may be returned to their home cage.
If surgery is to be performed on either lactating females or on pre-weaned pups, please contact the DLAR veterinary staff to discuss methods for re-introduction of pups to the dam.

Post-Operative Care

Animals should be monitored on a daily basis, as described in the approved IACUC protocol. The surgical site should be monitored daily to insure that the surgical wound is healing properly and that stitch abscesses, dehiscence or other complications have not occurred.
Analgesics are administered, per the IACUC approved protocol. For minor procedures it may be appropriate to administer only one dose of an analgesic. For major procedures narcotic analgesics should be administered for the first 24 hours postoperatively and continued if necessary. If unanticipated post-operative pain is observed, consult with the DLAR veterinary staff for pain management.
Non-absorbable sutures or wound clips should be removed from 7-14 days post-operatively. If not removed, the remaining suture material or clips can act as a foreign body and lead to skin infections.

References

Popp MB, Brennan MF. Long-term vascular access in the rat: importance of asepsis. Am J. Physiol. 241, H606, 1981.
Bradfield JF, Schachtman TR, McLaughlin RM, Steffen EK. Behavioral and physiologic effects of inapparent wound infection in rats. Lab Anim Sci 42(6):572-8, 1992.
Waynforth HB, Flecknell PA. Experimental and Surgical Technique in the Rat, 2nd Edition; Academic Press, 1992.
White WJ, Field KJ. Anesthesia and surgery of laboratory animals. Vet. Clin. North. Am. Small Anim. Pract. 17(5):989-1017, 1987.