Consistently delivered high-quality programs of husbandry and veterinary care provide the foundation that enables valid scientific research.
Standard operating procedures are valuable, and training and supervision are essential to ensure a consistently applied and uniformly high level of animal care. Environmental conditions must be carefully controlled within research facilities, along with conscientious application of animal care and use programs, to provide the best possible conditions for conducting research.
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. Increased emphasis has been placed on the wellness and workforce sustainability of animal care staff, with institutions implementing mental health support, workload monitoring, and retention programs to mitigate burnout in high-demand research environments.
The Guide for the Care and Use of Laboratory Animals 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 have antibodies from 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 prevent the introduction of infectious agents are effective measures to prevent disease within a colony that might confound or ruin experiments.
Continuous colony health monitoring is now achieved via real-time health status dashboards that integrate sentinel testing, environmental data, veterinary records, and microbiome profiles into centralized visualization platforms (1). These dashboards support early detection of infection or environmental deviations and enhance decision-making to maintain experimental reproducibility.
For certain studies, particularly those associated with immunity, it has become clear that animals protected from a broad array of organisms have an insufficient immune repertoire to be suitable. Adaptable management programs can maintain and segregate subjects from axenic mice (completely germ free) and gnotobiotic mice (having a specific and known microbiome) mice to those in close proximity or intentionally exposed to and potentially harboring undesirable pathogens.
Recent studies have emphasized the need for "wildling" and microbiota-transplanted models in immunology and behavioral research (2, 3), requiring husbandry systems that support variable microbiomes and controlled environmental exposures.
Although some nonhuman primate (NHP) colonies are free of most agents that cause infectious disease in these species, many NHPs used for research are of feral or wild origin. For this reason, it is critical to follow appropriate quarantine, isolation, and conditioning programs, as well as the program implemented in the importers' facilities. Biosecurity protocols for NHPs emphasize molecular surveillance, serological panels, and behavioral conditioning during quarantine to minimize stress and ensure early detection of zoonotic pathogens.
Housing for Laboratory Animals
Cages, pens, or runs for laboratory animals should provide adequate space for normal physiological 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, easy to clean and sanitize, and designed for comfort and safety. Static microisolation (filter-top) cages and individually ventilated cage (IVC) systems impede cage-to-cage transmission of infectious agents. However, infection can be transmitted horizontally or vertically from parents in breeding colonies. Naive mice introduced for crossbreeding and backcrossing can perpetuate infection, and experimental mice can be exposed to pathogens via a contaminated environment, shared watering valves, research devices, or transfer between laboratories.
Individual ventilation of cages maintains a consistent microenvironment. It also saves space in the facility and can be engineered to minimize odors, allergens, dust, and heat exhausted into the macroenvironment. Integrated cage-monitoring technologies that assess temperature, humidity, and ammonia levels in real time are used to enhance both animal welfare and early detection of housing deficiencies.
US federal law 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 NHPs must provide social and environmental enrichment to promote the animals' psychological well-being and be compatible with the experimental and practical constraints of the housing situation. Successful enrichment strategies for NHPs have included pair or group housing; variation in dietary content and method of presentation; ancillary cage equipment (eg, perches, swings, or ladders); devices to enhance visual, auditory, or tactile stimulation; and challenging, nonaversive behavioral 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 and monitored for all laboratory animals at all times. Unstable environmental conditions can profoundly affect animals' comfort, well-being, and metabolism and therefore the quality of experimental data derived from studies of those animals.
Not only must optimal temperatures be provided for the animals, but also the air temperature should be maintained within the comfort range at which human workers perform best (18–22°C [64–73°F]) and within ±1.1°C (2°F) of the set point. Ranges to consider for the optimal well-being of adapted animals are given in the table Preferred Temperature Ranges for Commonly Used Laboratory Animals.
Preferred Temperature Ranges for Commonly Used Laboratory Animals
Species | Preferred Temperature Range for Housing |
|---|---|
Rodents | 18–26°C (64–79°F) |
Rabbits | 16–22°C (61–72°F) |
Ferrets | 15–21°C (59–70°F) |
Nonhuman primates | 18–29°C (64–84°F) |
Zebrafish and other tropical fish | 27–32°C (80−90°F) |
South African clawed frog (Xenopus laevis) and axolotl (Ambystoma mexicanum) | 18–20°C (64–68°F) |
West African clawed frog (Xenopus tropicalis) | 24–26°C (75–79°F) |
Emerging evidence suggests that mice, regardless of age, presence or absence of fur, or immune status, historically have been kept under cold-stress conditions, especially in IVC systems (4, 5, 6, 7, 8, 9). This practice is to the possible detriment of colony production and many types of experiments, especially those that are reliant on the immune system or are associated with metabolism or physiology. Experience suggests, however, that mice that are 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, preferably within 10% of the set point. Updated guidance highlights thermal support strategies such as nesting materials and radiant heat zones as beneficial interventions to lessen cold stress and improve breeding outcomes in mice housed under standard IVC conditions.
Ventilation rates should be 10–15 fresh air changes per hour; however, these rates can be decreased to save energy in some situations (eg, rodent IVC systems with rack effluent directly connected to the room exhaust). Air recirculation is discouraged unless the air is treated to remove particulate and gaseous contaminants.
Evenly distributed and sufficiently intense lighting promotes animal well-being and circadian rhythmicity, and it enables personnel to observe the animals and perform all husbandry and sanitation duties safely and effectively (10).
Light cycles, as determined by species' requirements, should be controlled by automated timers to maintain circadian and neuroendocrine regulation. For example, 12 hours of light and 12 hours of dark is the most common standard used in rodent and general laboratory animal research. In breeding colonies, especially for rodents, a schedule of 14 hours of light and 10 hours of dark (eg, lights on at 6 am and off at 8 pm) is used to stimulate reproductive activity.
Although LED (light-emitting diode) technology offers many advantages that are known to consumers, the effect of LED light on circadian rhythms differs from that of the cool, white fluorescent lighting historically used. LED lighting represents arguably the most momentous environmental change influencing experiments since the removal of windows and exposure to seasonal daylight from animal research facilities (11), and it is now standard.
Refined LED lighting protocols align spectral outputs with species-specific circadian sensitivity, and tunable lighting is used in some modern facilities.
The microenvironment within certain types of caging can differ greatly from that of the macroenvironment of the room. Carefully conducted additional research is needed to more precisely define the best environmental and social conditions for each species or group of species at the cage level. Incomplete description of the environment associated with published animal experiments has been a long-standing flaw of research that has contributed to the unreliability in reproducing some animal experiment results over time and across laboratories.
Bedding for Laboratory Animals
Animal bedding is a controllable environmental factor that influences experimental data and animal well-being. Bedding materials should be nonirritating, absorbent, free of chemical contamination and pathogens, and unpalatable to discourage ingestion. Adequate quantities are vital to keep animals dry and clean between changes of bedding or caging.
Rodents are usually maintained on contact bedding consisting of either ground corncobs, hardwood chips, recycled paper, heat-treated softwood shavings, or virgin cellulose (tree pulp). Untreated softwoods are not recommended, because they contain volatile oils that can alter hepatic enzyme systems and affect certain kinds of research.
Bedding materials also vary in their phytoestrogen content and ability to suppress ammonia, which is the by-product of fecal bacterial urease catalysis of urea and also the most abundant noxious gas produced in the microenvironment.
Variation in bedding product type can introduce subtle variability to reproduction, behavior, physiology, mucosal immunity, dietary studies, tactile perception, the microbiome, and environmental microorganisms. Some institutions have adopted standardized bedding validation protocols, including periodic chemical analysis and microbiome impact studies, to support reproducibility and decrease interfacility variation.
Bedding sourced from recycled content or agricultural by-products should be monitored for pesticide residues and heavy metals. 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 the highest numbers on contact bedding, prefer softer materials and in volumes that allow burrowing and nesting.
Feeding of Laboratory Animals
Feed should be of adequate quantity, palatable, free of contaminants, nutritionally adequate, easily accessible, and provided via 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.
Routine mycotoxin screening of grain-based laboratory diets and evaluation of phytoestrogen levels in soy-containing feeds is recommended, because both mycotoxins and phytoestrogens can influence endocrine and reproductive parameters in sensitive models.
Feed should be manufactured, transported, stored, and used in ways that minimize its deterioration, contamination, or infestation. Laboratory animal diets (including those fortified with vitamin C) generally have a 9-month shelf life if stored at or below 21°C (70°F) and 50% relative humidity.
Most small animals are fed ad lib and consume food in quantities determined by their energy requirements. Consumption can be influenced by the environment and dictated by genotype. To prevent obesity and decrease wastage, rabbits, laboratory carnivores, swine, aquatic amphibians, and NHPs may receive 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 certain kinds of research.
Autoclavable or irradiated diets are available for rodents and can be used when sterilization of feed is desired. Although autoclaving of feed has been recommended to protect specific-pathogen-free colonies in cases when microbial contamination could invalidate studies, some institutions have reassessed the effect of sterilization protocols on vitamin stability and palatability in rodent diets and have adopted vacuum-packed irradiated alternatives when feasible.
Water Requirements of Laboratory Animals
For terrestrial animals in research settings, potable, uncontaminated water must be provided in adequate quantities to meet specific species requirements. Water entering an animal research facility typically comes from a local source (or sometimes from wells) and meets general standards for human consumption.
No matter what the source is, water quality can vary considerably, depending on several factors. These include geographical locale; proximity to industrial, urban, or agricultural settings; and municipal treatment approaches. Expanded monitoring of drinking water for potential contaminants, such as PFAS (per- and polyfluoroalkyl substances, known commonly as "forever chemicals"), has become a concern in animal facilities located in affected regions. Consequently, drinking water is a variable that can profoundly affect research.
Water entering an animal research facility should undergo sediment and carbon filtration, water softening, and additional purification, such as deionization or reverse osmosis filtration, before being stored in reservoirs or distributed to animals. These reservoirs should have multiple days' supply in reserve.
Water quality assurance programs that monitor pH, hardness, chemical content, and microbial load are recommended. Acidified, chlorinated, or sterile water might be required under certain experimental or husbandry conditions.
Water is usually provided to laboratory animals ad lib via bottles, plastic pouches, automated delivery, or, in certain applications, nonwetting, sterile gels.
Automated water delivery, commonly used in rodent housing, decreases operational costs, increases safety for animal care technicians, saves labor, decreases disruptions of the animals by caretakers, and provides consistently high water quality. The drawback of automated water delivery is the risk of hypothermia, drowning, or dehydration of cage inhabitants if the in-cage water delivery valve fails.
Automated water delivery systems require regular monitoring of function and periodic preventive maintenance, including full distribution system sanitization to prevent biofilm formation. Newer automated watering systems for rodents include leak sensors and telemetry-based alert mechanisms to mitigate risks of unnoticed valve failure.
Water quality is the most important environmental variable for aquatic species and a key determinant of their health. Inadequate water quality or fluctuations of water temperature are physiological stressors that affect the animals' intake, digestion, and use of food; alter their immune system; and predispose them 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 biological filtration and periodic partial replenishment with fresh water, just as with fish, helps suppress bacterial counts and prevents the buildup of toxic chemicals.
Sanitation of Laboratory Animal Facilities
A uniformly high level of animal enclosure and facility sanitation is mandatory to ensure that laboratory 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, bedding changes 1–3 times per week are usually enough; for rodents, rabbits, and NHPs in suspended cages over excreta pans and for mice in IVC 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.
Water bottles and other watering or feeding devices should be cleaned and sanitized at least weekly. Automated watering devices in cages, on racks, or in rooms should be designed and programmed to flush continuously or regularly, or they should be manually drained, rinsed, and sanitized at regular, frequent intervals.
Heating cages and other equipment to 82.2°C (180°F), or using appropriate chemical disinfection (eg, hypochlorite solutions) after removing biomatter, kills non-spore-forming pathogenic bacteria and viruses. All caging and other equipment should be rinsed thoroughly after treatment with detergents or disinfectants.
The effectiveness of sanitation programs should be evaluated regularly using appropriate microbiological, organic material detection systems or other means.
Routine audits and logs of cage wash temperatures, contact times, and chemical concentrations have become standard for confirming sanitation effectiveness. AAALAC and USDA inspections emphasize documentation and staff training related to sanitation protocols, particularly cleaning reusable enrichment items, transfer stations, and automated watering systems.
Pest Control in Laboratory Animal Facilities
Professionally directed programs to prevent, identify, and eradicate or control insects or escaped, feral, or wild rodents at research facilities must be instituted, regularly scheduled, and consistently documented. Pesticide use should be a last resort and limited to areas not used for animals or for storing feed or bedding. Researchers should be informed if these agents are used near animals or near their food or bedding.
Relatively inert substances, such as silica aerogel or boric acid powder, are recommended to help control crawling insects (eg, cockroaches). Updated guidance has emphasized integrated pest management (IPM) strategies that focus on habitat change, exclusion methods, and real-time monitoring using nontoxic traps and digital tracking systems.
During accreditation and regulatory inspections, the keeping of pest control logs, especially identifying entry points and follow-up actions, is increasingly emphasized.
For More Information
Hessler JR, Lehner NDM. Planning and Designing Research Animal Facilities. 1st ed. Elsevier/Academic Press; 2009.
Edstrom EK, Curran R. Quality assurance of animal watering systems. Lab Anim (NY).2003;32(5):32-35.
Peveler JL, Crisler R, Hickman D. Quality testing of autoclaved rodent drinking water during short-term and long-term storage. Lab Anim (NY). 2015;44(6):211-215.
Schoenberger JM, Prendergast BJ, Luchins KR, Theriault BR, Langan GP. Preference of escaped mice for live capture or glue traps and relevance to pest control programs. J Am Assoc Lab Anim Sci. 2023;62(1):38-47.
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