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Management of Laboratory Animals


Michael J. Huerkamp

, DVM, Emory University

Last full review/revision Feb 2021 | Content last modified Feb 2021
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Consistently delivered quality programs of husbandry and veterinary care provide the foundation that enables valid scientific research. For proper management of research animals, the animal care and research staff must be responsible, sensitive to the animals’ health and well-being, well trained in the humane care and use of laboratory animals, highly motivated, experienced, and diligent in performing their duties and responsibilities. Standard operating procedures are valuable, and training and supervision are essential to assure a consistently applied and uniformly high level of animal care. Within research facilities, environmental conditions must be carefully controlled so that, along with conscientiously applied programs of animal care and use, the best possible conditions for conducting research are provided.

The Guide remains the primary reference for information on basic principles and standards for laboratory animal management. Laboratory rodents that are disease- and pathogen-free and that do not possess antibodies indicative of past infection are readily available from commercial vendors. Procuring such animals from high-quality sources, transporting them in filtered shipping containers, and maintaining them in facilities with both physical and procedural barriers to the introduction of infectious agents are effective measures to prevent disease within a colony that may confound or ruin experiments.

For certain studies, particularly those associated with immunity, it has become clear that animals protected from exposure to a broad array of organisms have an immune repertoire insufficiently developed to be suitable. This requires adaptable management programs to maintain and segregate subjects across an expanse of health profiles from axenic and gnotobiotic to those in close proximity or intentionally exposed to and potentially harboring a medley of otherwise undesirable pathogens.

Although there are colonies of some species of primates that are free of most agents that cause infectious disease in these species, many primates used are of feral or wild origin. For this reason, appropriate quarantine, isolation, and conditioning programs are critical, in addition to the program followed in the importers’ facilities.


Cages, pens, or runs should provide adequate space to allow for normal physiologic needs, permit postural adjustments, and meet requirements for species-specific behavior. When possible, compatible groups of social animals should be housed together. Primary enclosures should be constructed of durable materials, easily cleaned and sanitized, and designed for comfort and safety. Static microisolation (filter-top) cages and individually ventilated caging (IVC) systems impede cage-to-cage transmission of infectious agents. However, infection can be transmitted horizontally or vertically from parents to progeny in breeding colonies; naive mice introduced for cross-breeding and back-crossing can perpetuate infection; and experimental mice potentially can be exposed to pathogens via a contaminated environment, shared watering valves, research devices, or when taken to laboratories. Individual ventilation of cages serves to delay deterioration of the environment within the cage and maintain a more consistent and wholesome microenvironment; it also saves space in the facility and can be engineered to minimize odors, allergens, dust, and heat exhausted into the macroenvironment.

Federal law in the USA requires that laboratory dogs have an opportunity to exercise regularly and have sensory contact with other dogs unless restricted by experimental or behavioral considerations. Housing for nonhuman primates must provide social and environmental enrichment to promote their psychological well-being compatible with the experimental and practical constraints of the housing situation. Successful enrichment strategies for nonhuman primates have included pair or group housing; variation in the dietary content and method of presentation; diversification of the internal cage environment with ancillary equipment (eg, perches, swings, or ladders); provision of devices to enhance visual, auditory, or tactile stimulation; and participation in challenging, nonaversive behavioral laboratory studies. Efforts to extend and adapt environmental enrichment practices to other laboratory animal species warrant consideration.

Temperature, relative humidity, ventilation rates, lighting conditions (spectrum, intensity, and photoperiod), gaseous pollutants (eg, ammonia), and noise should be carefully controlled at all times and monitored as appropriate. Unstable environmental conditions can have a profound effect on the comfort, well-being, and metabolism of animals and therefore on the quality of experimental data derived.

In general, air temperature should take into consideration the comfort ranges where human workers perform best of 64–73°F (18–22°C) and be set at +/- 2°F of set point. Ranges to take into consideration for optimal well-being of adapted animals are given in the accompanying table. Emerging evidence suggests that mice, whether adults, neonates, hirsute or not, in particular, have been kept traditionally under cold-stress conditions, especially in IVC, to the possible detriment of colony production and many types of experiments, especially those reliant on the immune system or associated with metabolism or physiology. Experience suggests, however, that mice provided an adequate energy diet and housed socially can compensate via nest building and huddling. Relative humidity should be maintained at 30%–70% for most species and preferably within 10% of the set point.


Preferred Temperature Ranges for Commonly Used Laboratory Animals


Preferred Temperature Range for Housing


64–79°F (18–26°C)


61–72°F (16–22°C)


59–70°F (15–21°C)


64–84°F (18–29°C)

Zebrafish and other tropical fish

80°−90°F (27°–32°C)

South African clawed frog (Xenopus laevis) and axolotl (Ambystoma mexicanum)

64°–68°F (18°–20°C)

West African clawed frog (X tropicalis)

75°–79°F (24°–26°C)

Ventilation rates should be 10–15 fresh air changes/hour but can be reduced in some situations to save energy (eg, rodent IVC with rack effluent directly connected to the room exhaust). Air recirculation is discouraged unless treated to remove particulate and gaseous contaminants.

Evenly distributed and sufficiently intense lighting promotes animal well-being, circadian rhythmicity, and allows personnel to observe the animals and perform all husbandry and sanitation duties safely and effectively. Diurnal or day-night cycles, as determined by species’ requirements, should be controlled by automatic timers to maintain circadian and neuroendocrine regulation. The introduction of LED for lighting, although offering many advantages known to consumers, influences circadian rhythms differently than cool, white fluorescent lighting traditionally in use and represents arguably the most momentous environmental change influencing experiments since the removal of windows and exposure to seasonal daylight from animal research facilities decades ago.

The microenvironment within certain types of caging may be very different from that of the macroenvironment of the room. Carefully conducted additional research is needed to more precisely define the optimal environmental and social conditions for each species or group of species at the cage level. Incomplete description of the environment in association with published animal experiments has been a longstanding flaw contributing to the contemporary crisis of unreliability in reproducing some animal experiment results over time and across laboratories.


Animal bedding is a controllable environmental factor that can influence experimental data and animal well-being. Bedding materials should be nonirritating, absorbent, free of chemical contamination and pathogens, and unpalatable, to discourage digestion. Adequate quantities are vital to keep animals dry and clean between changes of bedding or caging.

Rodents are the order of mammals maintained most commonly on contact bedding, of which ground corncobs, hardwood chips, recycled paper, heat-treated softwood shavings, or virgin cellulose are most commonly used. Untreated softwoods are not recommended because they contain volatile oils that may alter hepatic enzyme systems and affect certain kinds of research. This variation in product type has the potential to introduce subtle variability to reproduction, behavior, physiology, mucosal immunity, dietary studies, tactile perception, the microbiome, and environmental microorganisms.

Bedding materials also vary in their vitamin C and phytoestrogen content and ability to suppress ammonia, the byproduct of fecal bacterial urease catalysis of urea and the most abundant, noxious gas produced in the microenvironment. Depending on research requirements, bedding may be sterilized by autoclaving or irradiation before use or may be used as is. Mice, the species maintained in highest numbers on contact bedding, prefer softer materials and those in volumes that allow burrowing and nesting, supplemented with enrichment facilitating these and other normal behaviors. In light of these facts, those who care for and use animals in research should be cognizant that the wide variety in bedding materials may have equally broad effects upon behavior and biological processes that may influence experimental outcomes.


Feed should be of adequate quantity, palatable, free of contaminants, nutritionally adequate, easily accessible, and provided using means that meet behavioral needs according to specific species requirements. Feeds specifically manufactured for research animal use are preferred, because they are more likely to be uniformly constituted, free of contaminants, of known shelf life, and mill dated. Feed should be manufactured, transported, stored, and used in ways that minimize its deterioration, contamination, or infestation. Diets for laboratory animals generally have a 9-month shelf life if stored at or below 70°F (21°C) and 50% relative humidity. Owing to use of a stabilized type, this is the case even for diets fortified with vitamin C.

Most small animals consume food in relation to their energy requirements as influenced by the environment and dictated by their genotype and are fed ad lib; to prevent obesity and reduce wastage, rabbits, laboratory carnivores, swine, aquatic amphibians, and primates may be restricted to measured quantities of feed each day. As a general rule, laboratory animals minimally consume 4%–6% of their body weight in food daily. In addition to commercially prepared and usually pelleted natural ingredient diets of varying specification (eg, quality control and assurance of ingredients), semisynthetic or completely synthetic diets and all-liquid preparations can be formulated for use in certain kinds of research. Autoclavable or irradiated diets are available for rodents and can be used when sterilization of feed is desired.


For terrestrial animals, potable, uncontaminated water must be provided in adequate quantities to meet specific species requirements. Water entering an animal research facility will typically be supplied from a local, domestic source (although sometimes from wells), meeting general standards appropriate for human consumption. This water, however, may still be subject to considerable variation depending upon a number of factors, including geographic locale, the proximity to industrial, urban, or agricultural settings, and municipal treatment approaches. Drinking water consequently is a variable that can profoundly affect research.

Prior to presentation for consumption, water entering the animal research facility should undergo sediment and carbon filtration, water softening, and additional purification, such as deionization or reverse osmosis filtration, prior to storage in reservoirs of capacity with multiple days’ supply in reserve and distribution for animal consumption. Quality assurance programs that monitor pH, hardness, chemical content, and microbial load are recommended. Acidified, chlorinated, or sterile water may be required under certain experimental or husbandry conditions.

Water is usually provided ad lib via bottles, plastic pouches, via automated delivery, and, in certain applications, nonwetting, sterile gels. Particularly in the housing of rodents, an automated water supply enhances the advantages of ventilated caging systems and reduces operational costs/expenses, increases safety for animal care technicians, saves labor, reduces disruptions of the mice by caretakers, and provides consistently high water quality. The drawback of the use of automated drinking water supply for rodents is the risk of hypothermia, drowning, or dehydration of cage inhabitants as a consequence of failure of the in-cage water delivery valve. These systems do not operate automatically and require regular oversight of function and periodic preventive maintenance, including full distribution system sanitization to prevent biofilm formation.

Water quality is the most important environmental variable for aquatic species and a key determinant of health. Inadequate water quality or fluctuations of water temperature are physiologic stressors that impact the intake, digestion, and use of food; alter the immune system; and predispose to opportunistic infection. Water for aquatic vertebrates should be free of nitrite, ammonia, and chlorine, with total coliform counts not exceeding 200/mL. The pH should be 6.5–8.5. Although aquatic amphibians may be maintained in small containers of standing water, water recirculation with biologic filtration and periodic partial replenishment with fresh water, just as with fish, are helpful in suppressing bacterial counts and preventing the build-up of toxic chemicals.


A uniformly high level of animal enclosure and facility sanitation is mandatory to ensure that animals are clean and dry, air quality is adequate (without using masking agents), and primary enclosure surfaces and accessories are clean. Housing rooms, ancillary support spaces, and primary enclosures should be cleaned and sanitized as often as necessary to keep them free of dirt, debris, and potentially harmful contamination. For rodents in solid-bottom cages, usually 1–3 changes per week will suffice; for rodents, rabbits, and nonhuman primates in suspended cages over excreta pans and for mice in ventilated caging systems, cage changes every other week should be adequate. For larger animals, excreta and soiled bedding should be removed daily, and primary enclosures cleaned and sanitized daily, or at minimum every other week.

Water bottles and other watering or feeding devices should be cleaned and sanitized at least weekly. Automated watering devices on cages, racks, or in rooms should be designed and programmed to flush continually or regularly or they should be manually drained, rinsed, and sanitized at regular, frequent intervals.

Heating cages and other equipment to 180°F (82.2°C) or using appropriate chemical disinfection (eg, hypochlorite solutions) kills nonsporeforming pathogenic bacteria and viruses. All caging and other equipment should be rinsed thoroughly after treatment with detergents or disinfectants. The effectiveness of the programs of sanitation should be evaluated regularly using appropriate microbiologic, organic material detection systems or other means.

Vermin Control

Professionally directed programs to prevent, identify, and eradicate or control insects or escaped, feral, or wild rodents must be instituted, regularly scheduled, and consistently documented. The use of pesticides should be as a last resort and generally be confined to areas not used for animals or for storage of feed or bedding. If these agents are used in proximity to animals or their food and bedding, researchers should be informed of the use. Relatively inert substances, such as silica aerogel or boric acid powder, are recommended and are useful for control of crawling insects, eg, cockroaches.

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