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National Research Council (US) Committee for the Update of the Guide for the Care and Use of Laboratory Animals. Guide for the Care and Use of Laboratory Animals. 8th edition. Washington (DC): National Academies Press (US); 2011.

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Guide for the Care and Use of Laboratory Animals. 8th edition.

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3Environment, Housing, and Management

This chapter provides guidelines for the environment, housing, and management of laboratory animals used or produced for research, testing, and teaching. These guidelines are applicable across species and are relatively general; additional information should be sought about how to apply them to meet the specific needs of any species, strain, or use (see Appendix A for references). The chapter is divided into recommendations for terrestrial (page 42) and aquatic animals (page 77), as there are fundamental differences in their environmental requirements as well as animal husbandry, housing, and care needs. Although formulated specifically for vertebrate species, the general principles of humane animal care as set out in the Guide may also apply to invertebrate species.

The design of animal facilities combined with appropriate animal housing and management are essential contributors to animal well-being, the quality of animal research and production, teaching or testing programs involving animals, and the health and safety of personnel. An appropriate Program (see Chapter 2) provides environments, housing, and management that are well suited for the species or strains of animals maintained and takes into account their physical, physiologic, and behavioral needs, allowing them to grow, mature, and reproduce normally while providing for their health and well-being.

Fish, amphibians, and reptiles are poikilothermic animals: their core temperature varies with environmental conditions and they have limited ability (compared with birds and mammals) to metabolically maintain core temperature. The majority of poikilothermic laboratory animals are aquatic species—for example, fish and most amphibians—although some, such as reptiles and certain amphibian species, are terrestrial. Personnel working with aquatic animals should be familiar with management implications, e.g., the importance of providing appropriate temperature ranges for basic physiologic function.


Terrestrial Environment

Microenvironment and Macroenvironment

The microenvironment of a terrestrial animal is the physical environment immediately surrounding it; that is, the primary enclosure such as the cage, pen, or stall. It contains all the resources with which the animals come directly in contact and also provides the limits of the animals’ immediate environment. The microenvironment is characterized by many factors, including illumination, noise, vibration, temperature, humidity, and gaseous and particulate composition of the air. The physical environment of the secondary enclosure, such as a room, a barn, or an outdoor habitat, constitutes the macroenvironment.

Microenvironment: The immediate physical environment surrounding the animal (i.e., the environment in the primary enclosure such as the cage, pen, or stall).

Although the microenvironment and the macroenvironment are generally related, the microenvironment can be appreciably different and affected by several factors, including the design of the primary enclosure and macroenvironmental conditions.

Macroenvironment: The physical environment of the secondary enclosure (e.g., a room, a barn, or an outdoor habitat).

Evaluation of the microenvironment of small enclosures can be difficult. Available data indicate that temperature, humidity, and concentrations of gases and particulate matter are often higher in the animal microenvironment than in the macroenvironment (Besch 1980; Hasenau et al. 1993; Perkins and Lipman 1995; E. Smith et al. 2004), while light levels are usually lower. Microenvironmental conditions can directly affect physiologic processes and behavior and may alter disease susceptibility (Baer et al. 1997; Broderson et al. 1976; Memarzadeh et al. 2004; Schoeb et al. 1982; Vesell et al. 1976).

Temperature and Humidity

Maintenance of body temperature within normal circadian variation is necessary for animal well-being. Animals should be housed within temperature and humidity ranges appropriate for the species, to which they can adapt with minimal stress and physiologic alteration.

The ambient temperature range in which thermoregulation occurs without the need to increase metabolic heat production or activate evaporative heat loss mechanisms is called the thermoneutral zone (TNZ) and is bounded by the lower and upper critical temperatures (LCTs and UCTs; Gordon 2005). To maintain body temperature under a given environmental temperature animals adjust physiologically (including their metabolism) and behaviorally (including their activity level and resource use). For example, the TNZ of mice ranges between 26°C and 34°C (Gordon 1993); at lower temperatures, building nests and huddling for resting and sleeping allow them to thermoregulate by behaviorally controlling their microclimate. Although mice choose temperatures below their LCT of 26°C during activity periods, they strongly prefer temperatures above their LCT for maintenance and resting behaviors (Gaskill et al. 2009; Gordon 2004; Gordon et al. 1998). Similar LCT values are found in the literature for other rodents, varying between 26–30°C for rats and 28–32°C for gerbils (Gordon 1993). The LCTs of rabbits (15–20°C; Gonzalez et al. 1971) and cats and dogs (20–25°C) are slightly lower, while those of nonhuman primates and farm animals vary depending on the species. In general, dry-bulb temperatures in animal rooms should be set below the animals’ LCT to avoid heat stress. This, in turn, means that animals should be provided with adequate resources for thermoregulation (nesting material, shelter) to avoid cold stress. Adequate resources for thermoregulation are particularly important for newborn animals whose LCT is normally considerably higher than that of their adult conspecifics.

Environmental temperature and relative humidity can be affected by husbandry and housing design and can differ considerably between primary and secondary enclosures as well as within primary enclosures. Factors that contribute to variation in temperature and humidity between and within enclosures include housing design; construction material; enrichment devices such as shelters and nesting material; use of filter tops; number, age, type, and size of the animals in each enclosure; forced ventilation of enclosures; and the type and frequency of contact bedding changes (Besch 1980).

Exposure to wide temperature and humidity fluctuations or extremes may result in behavioral, physiologic, and morphologic changes, which might negatively affect animal well-being and research performance as well as outcomes of research protocols (Garrard et al. 1974; Gordon 1990, 1993; Pennycuik 1967). These effects can be multigenerational (Barnett 1965, 1973).

The dry-bulb temperatures listed in Table 3.1 are broad and generally reflect tolerable limits for common adult laboratory animal species, provided they are housed with adequate resources for behavioral thermoregulation; temperatures should normally be selected and maintained with minimal fluctuation near the middle of these ranges. Depending on the specific housing system employed, the selection of appropriate macro- and microenvironmental temperatures will differ based on a variety of factors, including but not limited to the species or strain, age, numbers of animals in the enclosure, size and construction of the primary enclosure, and husbandry conditions (e.g., use/provision of contact bedding, nesting material and/or shelter, individually ventilated cages). Poikilotherms and young birds of some species generally require a thermal gradient in their primary enclosure to meet basic physiological processes. The temperature ranges shown may not apply to captive wild animals, wild animals maintained in their natural environment, or animals in outdoor enclosures that have the opportunity to adapt by being exposed to seasonal changes in ambient conditions.

TABLE 3.1. Recommended Dry-Bulb Macroenvironmental Temperatures for Common Laboratory Animals.


Recommended Dry-Bulb Macroenvironmental Temperatures for Common Laboratory Animals.

Some conditions require increased environmental temperatures for housing (e.g., postoperative recovery, neonatal animals, rodents with hairless phenotypes, reptiles and amphibians at certain stages of reproduction). The magnitude of the temperature increase depends on housing details; sometimes raising the temperature in the microenvironment alone (e.g., by using heating pads for postoperative recovery or radiant heat sources for reptiles) rather than raising the temperature of the macroenvironment is sufficient and preferable.

Relative humidity should also be controlled, but not nearly as narrowly as temperature for many mammals; the acceptable range of relative humidity is considered to be 30% to 70% for most mammalian species. Microenvironmental relative humidity may be of greater importance for animals housed in a primary enclosure in which the environmental conditions differ greatly from those of the macroenvironment (e.g., in static filter-top [isolator] cages).

Some species may require conditions with high relative humidity (e.g., selected species of nonhuman primates, tropical reptiles, and amphibians; Olson and Palotay 1983). In mice, both abnormally high and low humidity may increase preweaning mortality (Clough 1982). In rats, low relative humidity, especially in combination with temperature extremes, may lead to ringtail, a condition involving ischemic necrosis of the tail and sometimes toes (Crippa et al. 2000; Njaa et al. 1957; Totten 1958). For some species, elevated relative humidity may affect an animal’s ability to cope with thermal extremes. Elevated microenvironmental relative humidity in rodent isolator cages may also lead to high intracage ammonia concentrations (Corning and Lipman 1991; Hasenau et al. 1993), which can be irritating to the nasal passages and alter some biologic responses (Gordon et al. 1980; Manninen et al. 1998). In climates where it is difficult to provide a sufficient level of environmental relative humidity, animals should be closely monitored for negative effects such as excessively flaky skin, ecdysis (molting) difficulties in reptiles, and desiccation stress in semiaquatic amphibians.

Ventilation and Air Quality

The primary purpose of ventilation is to provide appropriate air quality and a stable environment. Specifically, ventilation provides an adequate oxygen supply; removes thermal loads caused by the animals, personnel, lights, and equipment; dilutes gaseous and particulate contaminants including allergens and airborne pathogens; adjusts the moisture content and temperature of room air; and, where appropriate, creates air pressure differentials (directional air flow) between adjoining spaces. Importantly, ventilating the room (i.e., the macroenvironment) does not necessarily ensure adequate ventilation of an animal’s primary enclosure (i.e., the microenvironment), that is, the air to which the animal is actually exposed. The type of primary enclosure may considerably influence the differences between these two environments—for example, differences may be negligible when animals are housed in open caging or pens, whereas they can be significant when static isolator cages are used.

The volume and physical characteristics of the air supplied to a room and its diffusion pattern influence the ventilation of an animal’s primary enclosure and are important determinants of the animal’s microenvironment. The type and location of supply air diffusers and exhaust registers in relation to the number, arrangement, location, and type of primary and secondary enclosures affect how well the microenvironments are ventilated and should therefore be considered. The use of computer modeling for assessing those factors in relation to heat loading, air diffusion patterns, and particulate movement may be helpful in optimizing ventilation of micro- and macroenvironments (Hughes and Reynolds 1995).

Direct exposure of animals to air moving at high velocity (drafts) should be avoided as the speed of air to which animals are exposed affects the rate at which heat and moisture are removed from an animal. For example, air at 20°C moving at 60 linear feet per minute (18.3 m/min) has a cooling effect of approximately 7°C (Weihe 1971). Drafts can be particularly problematic for neonatal homeotherms (which may be hairless and have poorly developed mechanisms for thermoregulatory control), for mutants lacking fur, and for semiaquatic amphibians that can desiccate.

Provision of 10 to 15 fresh air changes per hour in animal housing rooms is an acceptable guideline to maintain macroenvironmental air quality by constant volume systems and may also ensure microenvironmental air quality. Although this range is effective in many animal housing settings, it does not take into account the range of possible heat loads; the species, size, and number of animals involved; the type of primary enclosure and bedding; the frequency of cage changing; the room dimensions; or the efficiency of air distribution both in the macroenvironment and between the macro- and microenvironments. In some situations, the use of such a broad guideline might overventilate a macroenvironment containing few animals, thereby wasting energy, or underventilate a microenvironment containing many animals, allowing heat, moisture, and pollutants to accumulate.

Modern heating, ventilation, and air conditioning (HVAC) systems (e.g., variable air volume, or VAV, systems) allow ventilation rates to be set in accordance with heat load and other variables. These systems offer considerable advantages with respect to flexibility and energy conservation, but should always provide a minimum amount of air exchange, as recommended for general use laboratories (Bell 2008; DiBerardinis et al. 2009).

Individually ventilated cages (IVCs) and other types of specialized primary enclosures, that either directly ventilate the enclosure using filtered room air or are ventilated independently of the room, can effectively address animals’ ventilation requirements without the need to increase macroenvironmental ventilation. However, cautions mentioned above regarding high-velocity air should be considered (Baumans et al. 2002; Krohn et al. 2003). Nevertheless, the macroenvironment should be ventilated sufficiently to address heat loads, particulates, odors, and waste gases released from primary enclosures (Lipman 1993).

If ventilated primary enclosures have adequate filtration to address contamination risks, air exhausted from the microenvironment may be returned to the room in which animals are housed, although it is generally preferable to exhaust these systems directly into the building’s exhaust system to reduce heat load and macroenvironmental contamination.

Static isolation caging (without forced ventilation), such as that used in some types of rodent housing, restricts ventilation (Keller et al. 1989). To compensate, it may be necessary to adjust husbandry practices, including sanitation and cage change frequency, selection of contact bedding, placement of cages in a secondary enclosure, animal densities in cages, and/or decrease in macroenvironmental relative humidity to improve the microenvironment and heat dissipation.

The use of recycled air to ventilate animal rooms may save energy but entails risks. Because many animal pathogens can be airborne or travel on fomites (e.g., dust), exhaust air recycled into HVAC systems that serve multiple rooms presents a risk of cross contamination. Recycling air from nonanimal use areas (e.g., some human occupancy areas and food, bedding, and supply storage areas) may require less intensive filtration or conditioning and pose less risk of infection. The risks in some situations, however, might be too great to consider recycling (e.g., in the case of non-human primates and biohazard areas). The exhaust air to be recycled should be filtered, at minimum, with 85–95% ASHRAE efficient filters to remove airborne particles before it is recycled (NAFA 1996). Depending on the air source, composition, and proportion of recycled air used (e.g., ammonia and other gases emitted from excrement in recirculating air from animal rooms), consideration should also be given to filtering volatile substances. In areas that require filtration to ensure personnel and/or animal safety (e.g., hazardous containment holding), filter efficiency, loading, and integrity should be assessed.

The successful operation of any HVAC system requires regular preventive maintenance and evaluation, including measurement of its function at the level of the secondary enclosure. Such measurements should include supply and exhaust air volumes, fluctuation in temperature and relative humidity, and air pressure differentials between spaces as well as critical mechanical operating parameters.


Light can affect the physiology, morphology, and behavior of various animals (Azar et al. 2008; Brainard et al. 1986; Erkert and Grober 1986; Newbold et al. 1991; Tucker et al. 1984). Potential photostressors include inappropriate photoperiod, photointensity, and spectral quality of the light (Stoskopf 1983).

Numerous factors can affect animals’ needs for light and should be considered when an appropriate illumination level is being established for an animal holding room. These include light intensity and wavelength as well as the duration of the animal’s current and prior exposure to light, and the animal’s pigmentation, circadian rhythm, body temperature, hormonal status, age, species, sex, and stock or strain (Brainard 1989; Duncan and O’Steen 1985; O’Steen 1980; Saltarelli and Coppola 1979; Semple-Row-land and Dawson 1987; Wax 1977). More recent studies in rodents and primates have shown the importance of intrinsically photosensitive retinal ganglion cells (distinct from rods and cones) for neuroendocrine, circadian, and neurobehavioral regulation (Berson et al. 2002; Hanifin and Brainard 2007). These cells can respond to light wavelengths that may differ from other photoreceptors and may influence the type of lighting, light intensity, and wavelength selected for certain types of research.

In general, lighting should be diffused throughout an animal holding area and provide sufficient illumination for the animals’ well-being while permitting good housekeeping practices, adequate animal inspection including for the bottom-most cages in racks, and safe working conditions for personnel. Light in animal holding rooms should provide for both adequate vision and neuroendocrine regulation of diurnal and circadian cycles (Brainard 1989).

Photoperiod is a critical regulator of reproductive behavior in many animal species (Brainard et al. 1986; Cherry 1987), so inadvertent light exposure during the dark cycle should be minimized or avoided. Because some species, such as chickens (Apeldoorn et al. 1999), will not eat in low light or darkness, such illumination schedules should be limited to a duration that will not compromise their well-being. A time-controlled lighting system should be used to ensure a regular diurnal cycle, and system performance should be checked regularly to ensure proper cycling.

Most commonly used laboratory rodents are nocturnal. Because albino rodents are more susceptible to phototoxic retinopathy than other animals (Beaumont 2002), they have been used as a basis for establishing room illumination levels (Lanum 1979). Data for room light intensities for other animals, based on scientific studies, are not available. Light levels of about 325 lux (30-ft candles) approximately 1 m (3.3 ft) above the floor appear to be sufficient for animal care and do not cause clinical signs of phototoxic retinopathy in albino rats (Bellhorn 1980). Levels up to 400 lux (37-ft candles) as measured in an empty room 1 m from the floor have been found to be satisfactory for rodents if management practices are used to prevent retinal damage in albinos (Clough 1982). However, the light experience of an individual animal can affect its sensitivity to phototoxicity; light of 130–270 lux above the light intensity under which it was raised has been reported to be near the threshold of retinal damage in some individual albino rats according to histologic, morphometric, and electrophysiologic evidence (Semple-Rowland and Dawson 1987). Some guidelines recommend a light intensity as low as 40 lux at the position of the animal in midcage (NASA 1988). Rats and mice generally prefer cages with low light intensity (Blom et al. 1996), and albino rats prefer areas with a light intensity of less than 25 lux (Schlingmann et al. 1993a). Young mice prefer much lower illumination than adults (Wax 1977). For animals that have been shown to be susceptible to phototoxic retinopathy, light should be between 130 and 325 lux in the room at cage level.

Light intensity decreases with the square of the distance from its source. Thus the location of a cage on a rack affects the intensity of light to which the animals within are exposed. Light intensity may differ as much as 80-fold in transparent cages from the top to the bottom of a rack, and differences up to 20-fold have been recorded within a cage (Schlingmann et al. 1993a,b). Management practices, such as rotating cage position relative to the light source (Greenman et al. 1982) or providing animals with ways to control their own light exposure by behavioral means (e.g., nesting or bedding material adequate for tunneling), can reduce inappropriate light stimulation. Variable-intensity lights are often used to accommodate the needs of research protocols, certain animal species, and energy conservation. However, such a system should also provide for the observation and care of the animals. Caution should be exercised as increases in daytime room illumination for maintenance purposes have been shown to change photoreceptor physiology and can alter circadian regulation (NRC 1996; Reme et al. 1991; Terman et al. 1991).

Noise and Vibration

Noise produced by animals and animal care activities is inherent in the operation of an animal facility (Pfaff and Stecker 1976) and noise control should be considered in facility design and operation (Pekrul 1991). Assessment of the potential effects of noise on an animal warrants consideration of the intensity, frequency, rapidity of onset, duration, and vibration potential of the sound and the hearing range, noise exposure history, and sound effect susceptibility of the species, stock, or strain. Similarly, occupational exposure to animal or animal care practices that generate noise may be of concern for personnel and, if of sufficient intensity, may warrant hearing protection.

Separation of human and animal areas minimizes disturbances to both human and animal occupants of the facility. Noisy animals, such as dogs, swine, goats, nonhuman primates, and some birds (e.g., zebra finches), should be housed away from quieter animals, such as rodents, rabbits, and cats. Environments should be designed to accommodate animals that make noise rather than resorting to methods of noise reduction. Exposure to sound louder than 85 dB can have both auditory and nonauditory effects (Fletcher 1976; Peterson 1980)—for example, eosinopenia, increased adrenal gland weights, and reduced fertility in rodents (Geber et al. 1966; Nayfield and Besch 1981; Rasmussen et al. 2009), and increased blood pressure in nonhuman primates (Peterson et al. 1981)—and may necessitate hearing protection for personnel (OSHA 1998). Many species can hear sound frequencies inaudible to humans (Brown and Pye 1975; Heffner and Heffner 2007); rodents, for example, are very sensitive to ultrasound (Olivier et al. 1994). The potential effects of equipment (such as video display terminals; Sales 1991; Sales et al. 1999) and materials that produce noise in the hearing range of nearby animals can thus become an uncontrolled variable for research experiments and should therefore be carefully considered (Turner et al. 2007; Willott 2007). To the greatest extent possible, activities that generate noise should be conducted in rooms or areas separate from those used for animal housing.

Because changes in patterns of sound exposure have different effects on different animals (Armario et al. 1985; Clough 1982), personnel should try to minimize the production of unnecessary noise. Excessive and intermittent noise can be minimized by training personnel in alternatives to noisy practices, the use of cushioned casters and bumpers on carts, trucks, and racks, and proper equipment maintenance (e.g., castor lubrication). Radios, alarms, and other sound generators should not be used in animal rooms unless they are part of an approved protocol or enrichment program. Any radios or sound generators used should be switched off at the end of the working day to minimize associated adverse physiologic changes (Baldwin 2007).

While some vibration is inherent to every facility and animal housing condition, excessive vibration has been associated with biochemical and reproductive changes in laboratory animals (Briese et al. 1984; Carman et al. 2007) and can become an uncontrolled variable for research experiments. The source of vibrations may be located within or outside the animal facility. In the latter case, groundborne vibration may affect both the structure and its contents, including animal racks and cages. Housing systems with moving components, such as ventilated caging system blowers, may create vibrations that could affect the animals housed within, especially if not functioning properly. Like noise, vibration varies with intensity, frequency, and duration. A variety of techniques may be used to isolate groundborne (see Chapter 5) and equipment-generated vibration (Carman et al. 2007). Attempts should be made to minimize the generation of vibration, including from humans, and excessive vibration should be avoided.

Terrestrial Housing

Microenvironment (Primary Enclosure)

All animals should be housed under conditions that provide sufficient space as well as supplementary structures and resources required to meet physical, physiologic, and behavioral needs. Environments that fail to meet the animals’ needs may result in abnormal brain development, physiologic dysfunction, and behavioral disorders (Garner 2005; van Praag et al. 2000; Würbel 2001) that may compromise both animal well-being and scientific validity. The primary enclosure or space may need to be enriched to prevent such effects (see also section on Environmental Enrichment).

An appropriate housing space or enclosure should also account for the animals’ social needs. Social animals should be housed in stable pairs or groups of compatible individuals unless they must be housed alone for experimental reasons or because of social incompatibility (see also section on Behavioral and Social Management). Structural adjustments are frequently required for social housing (e.g., perches, visual barriers, refuges), and important resources (e.g., food, water, and shelter) should be provided in such a way that they cannot be monopolized by dominant animals (see also section on Environmental Enrichment).

The primary enclosure should provide a secure environment that does not permit animal escape and should be made of durable, nontoxic materials that resist corrosion, withstand the rigors of cleaning and regular handling, and are not detrimental to the health and research use of the animals. The enclosure should be designed and manufactured to prevent accidental entrapment of animals or their appendages and should be free of sharp edges or projections that could cause injury to the animals or personnel. It should have smooth, impervious surfaces with minimal ledges, angles, corners, and overlapping surfaces so that accumulation of dirt, debris, and moisture is minimized and cleaning and disinfecting are not impaired. All enclosures should be kept in good repair to prevent escape of or injury to animals, promote physical comfort, and facilitate sanitation and servicing. Rusting or oxidized equipment, which threatens the health or safety of animals, needs to be repaired or replaced. Less durable materials, such as wood, may be appropriate in select situations, such as outdoor corrals, perches, climbing structures, resting areas, and perimeter fences for primary enclosures. Wooden items may need to be replaced periodically because of damage or difficulties with sanitation. Painting or sealing wood surfaces with nontoxic materials may improve durability in many instances.

Flooring should be solid, perforated, or slatted with a slip-resistant surface. In the case of perforated or slatted floors, the holes and slats should have smooth edges. Their size and spacing need to be commensurate with the size of the housed animal to minimize injury and the development of foot lesions. If wire-mesh flooring is used, a solid resting area may be beneficial, as this floor type can induce foot lesions in rodents and rabbits (Drescher 1993; Fullerton and Gilliatt 1967; Rommers and Meijerhof 1996). The size and weight of the animal as well as the duration of housing on wire-mesh floors may also play a role in the development of this condition (Peace et al. 2001). When given the choice, rodents prefer solid floors (with bedding) to grid or wire-mesh flooring (Blom et al. 1996; Manser et al. 1995, 1996).

Animals should have adequate bedding substrate and/or structures for resting and sleeping. For many animals (e.g., rodents) contact bedding expands the opportunities for species-typical behavior such as foraging, digging, burrowing, and nest building (Armstrong et al. 1998; Ivy et al. 2008). Moreover, it absorbs urine and feces to facilitate cleaning and sanitation. If provided in sufficient quantity to allow nest building or burrowing, bedding also facilitates thermoregulation (Gordon 2004). Breeding animals should have adequate nesting materials and/or substitute structures based on species-specific requirements (mice: Sherwin 2002; rats: Lawlor 2002; gerbils: Waiblinger 2002).

Specialized housing systems (e.g., isolation-type cages, IVCs, and gnotobiotic1 isolators) are available for rodents and certain species. These systems, designed to minimize the spread of airborne particles between cages or groups of cages, may require different husbandry practices, such as alterations in the frequency of bedding change, the use of aseptic handling techniques, and specialized cleaning, disinfecting, or sterilization regimens to prevent microbial transmission by other than airborne routes.

Appropriate housing strategies for a particular species should be developed and implemented by the animal care management, in consultation with the animal user and veterinarian, and reviewed by the IACUC. Housing should provide for the animals’ health and well-being while being consistent with the intended objectives of animal use. Expert advice should be sought when new species are housed or when there are special requirements associated with the animals or their intended use (e.g., genetically modified animals, invasive procedures, or hazardous agents). Objective assessments should be made to substantiate the adequacy of the animal’s environment, housing, and management. Whenever possible, routine procedures for maintaining animals should be documented to ensure consistency of management and care.

Environmental Enrichment

The primary aim of environmental enrichment is to enhance animal well-being by providing animals with sensory and motor stimulation, through structures and resources that facilitate the expression of species-typical behaviors and promote psychological well-being through physical exercise, manipulative activities, and cognitive challenges according to species-specific characteristics (NRC 1998a; Young 2003). Examples of enrichment include structural additions such as perches and visual barriers for nonhuman primates (Novak et al. 2007); elevated shelves for cats (Overall and Dyer 2005; van den Bos and de Cock Buning 1994) and rabbits (Stauffacher 1992); and shelters for guinea pigs (Baumans 2005), as well as manipulable resources such as novel objects and foraging devices for nonhuman primates; manipulable toys for nonhuman primates, dogs, cats, and swine; wooden chew sticks for some rodent species; and nesting material for mice (Gaskill et al. 2009; Hess et al. 2008; Hubrecht 1993; Lutz and Novak 2005; Olsson and Dahlborn 2002). Novelty of enrichment through rotation or replacement of items should be a consideration; however, changing animals’ environment too frequently may be stressful.

Well-conceived enrichment provides animals with choices and a degree of control over their environment, which allows them to better cope with environmental stressors (Newberry 1995). For example, visual barriers allow nonhuman primates to avoid social conflict; elevated shelves for rabbits and shelters for rodents allow them to retreat in case of disturbances (Baumans 1997; Chmiel and Noonan 1996; Stauffacher 1992); and nesting material and deep bedding allow mice to control their temperature and avoid cold stress during resting and sleeping (Gaskill et al. 2009; Gordon 1993, 2004).

Not every item added to the animals’ environment benefits their well-being. For example, marbles are used as a stressor in mouse anxiety studies (De Boer and Koolhaas 2003), indicating that some items may be detrimental to well-being. For nonhuman primates, novel objects can increase the risk of disease transmission (Bayne et al. 1993); foraging devices can lead to increased body weight (Brent 1995); shavings can lead to allergies and skin rashes in some individuals; and some objects can result in injury from foreign material in the intestine (Hahn et al. 2000). In some strains of mice, cage dividers and shelters have induced overt aggression in groups of males, resulting in social stress and injury (e.g., Bergmann et al. 1994; Haemisch et al. 1994). Social stress was most likely to occur when resources were monopolized by dominant animals (Bergmann et al. 1994).

Enrichment programs should be reviewed by the IACUC, researchers, and veterinarian on a regular basis to ensure that they are beneficial to animal well-being and consistent with the goals of animal use. They should be updated as needed to ensure that they reflect current knowledge. Personnel responsible for animal care and husbandry should receive training in the behavioral biology of the species they work with to appropriately monitor the effects of enrichment as well as identify the development of adverse or abnormal behaviors.

Like other environmental factors (such as space, light, noise, temperature, and animal care procedures), enrichment affects animal phenotype and may affect the experimental outcome. It should therefore be considered an independent variable and appropriately controlled.

Some scientists have raised concerns that environmental enrichment may compromise experimental standardization by introducing variability, adding not only diversity to the animals’ behavioral repertoire but also variation to their responses to experimental treatments (e.g., Bayne 2005; Eskola et al. 1999; Gärtner 1999; Tsai et al. 2003). A systematic study in mice did not find evidence to support this viewpoint (Wolfer et al. 2004), indicating that housing conditions can be enriched without compromising the precision or reproducibility of experimental results. Further research in other species may be needed to confirm this conclusion. However, it has been shown that conditions resulting in higher-stress reactivity increase variation in experimental data (e.g., Macrì et al. 2007). Because adequate environmental enrichment may reduce anxiety and stress reactivity (Chapillon et al. 1999), it may also contribute to higher test sensitivity and reduced animal use (Baumans 1997).

Sheltered or Outdoor Housing

Sheltered or outdoor housing (e.g., barns, corrals, pastures, islands) is a primary housing method for some species and is acceptable in many situations. Animals maintained in outdoor runs, pens, or other large enclosures must have protection from extremes in temperature or other harsh weather conditions and adequate opportunities for retreat (for subordinate animals). These goals can normally be achieved by providing windbreaks, species-appropriate shelters, shaded areas, areas with forced ventilation, heat-radiating structures, and/or means of retreat to conditioned spaces, such as an indoor portion of a run. Shelters should be large enough to accommodate all animals housed in the enclosure, be accessible at all times to all animals, have sufficient ventilation, and be designed to prevent buildup of waste materials and excessive moisture. Houses, dens, boxes, shelves, perches, and other furnishings should be constructed in a manner and made of materials that allow cleaning or replacement in accord with generally accepted husbandry practices.

Floors or ground-level surfaces of outdoor housing facilities may be covered with dirt, absorbent bedding, sand, gravel, grass, or similar material that can be removed or replaced when needed to ensure appropriate sanitation. Excessive buildup of animal waste and stagnant water should be avoided by, for example, using contoured or drained surfaces. Other surfaces should be able to withstand the elements and be easily maintained.

Successful management of outdoor housing relies on stable social groups of compatible animals; sufficient and species-appropriate feeding and resting places; an adequate acclimation period in advance of seasonal changes when animals are first introduced to outdoor housing; training of animals to cooperate with veterinary and investigative personnel (e.g., to enter chutes or cages for restraint or transport); and adequate security via a perimeter fence or other means.

Naturalistic Environments

Areas such as pastures and islands may provide a suitable environment for maintaining or producing animals and for some types of research. Their use results in the loss of some control over nutrition, health care and surveillance, and pedigree management. These limitations should be balanced against the benefits of having the animals live in more natural conditions. Animals should be added to, removed from, and returned to social groups in this setting with appropriate consideration of the effects on the individual animals and on the group. Adequate supplies of food, fresh water, and natural or constructed shelter should be ensured.


General Considerations for All Animals An animal’s space needs are complex and consideration of only the animal’s body weight or surface area may be inadequate. Important considerations for determining space needs include the age and sex of the animal(s), the number of animals to be cohoused and the duration of the accommodation, the use for which the animals are intended (e.g., production vs. experimentation), and any special needs they may have (e.g., vertical space for arboreal species or thermal gradient for poikilotherms). In many cases, for example, adolescent animals, which usually weigh less than adults but are more active, may require more space relative to body weight (Ikemoto and Panksepp 1992). Group-housed, social animals can share space such that the amount of space required per animal may decrease with increasing group size; thus larger groups may be housed at slightly higher stocking densities than smaller groups or individual animals. Socially housed animals should have sufficient space and structural complexity to allow them to escape aggression or hide from other animals in the pair or group. Breeding animals will require more space, particularly if neonatal animals will be raised together with their mother or as a breeding group until weaning age. Space quality also affects its usability. Enclosures that are complex and environmentally enriched may increase activity and facilitate the expression of species-specific behaviors, thereby increasing space needs. Thus there is no ideal formula for calculating an animal’s space needs based only on body size or weight and readers should take the performance indices discussed in this section into consideration when utilizing the species-specific guidelines presented in the following pages.

Consideration of floor area alone may not be sufficient in determining adequate cage size; with some species, cage volume and spatial arrangement may be of greater importance. In this regard, the Guide may differ from the US Animal Welfare Regulations (AWRs) or other guidelines. The height of an enclosure can be important to allow for expression of species-specific behaviors and postural adjustments. Cage height should take into account the animal’s typical posture and provide adequate clearance for the animal from cage structures, such as feeders and water devices. Some species—for example, nonhuman primates, cats, and arboreal animals—use the vertical dimensions of the cage to a greater extent than the floor. For these animals, the ability to stand or to perch with adequate vertical space to keep their body, including their tail, above the cage floor can improve their well-being (Clarence et al. 2006; MacLean et al. 2009).

Space allocations should be assessed, reviewed, and modified as necessary by the IACUC considering the performance indices (e.g., health, reproduction, growth, behavior, activity, and use of space) and special needs determined by the characteristics of the animal strain or species (e.g., obese, hyperactive, or arboreal animals) and experimental use (e.g., animals in long-term studies may require greater and more complex space). At a minimum, animals must have enough space to express their natural postures and postural adjustments without touching the enclosure walls or ceiling, be able to turn around, and have ready access to food and water. In addition, there must be sufficient space to comfortably rest away from areas soiled by urine and feces. Floor space taken up by food bowls, water containers, litter boxes, and enrichment devices (e.g., novel objects, toys, foraging devices) should not be considered part of the floor space.

The space recommendations presented here are based on professional judgment and experience. They should be considered the minimum for animals housed under conditions commonly found in laboratory animal housing facilities. Adjustments to the amount and arrangement of space recommended in the following tables should be reviewed and approved by the IACUC and should be based on performance indices related to animal well-being and research quality as described in the preceding paragraphs, with due consideration of the AWRs and PHS Policy and other applicable regulations and standards.

It is not within the scope of the Guide to discuss the housing requirements of all species used in research. For species not specifically indicated, advice should be sought from the scientific literature and from species-relevant experts.

Laboratory Rodents Table 3.2 lists recommended minimum space for commonly used laboratory rodents housed in groups. If they are housed singly or in small groups or exceed the weights in the table, more space per animal may be required, while larger groups may be housed at slightly higher densities.

TABLE 3.2. Recommended Minimum Space for Commonly Used Laboratory Rodents Housed in Groups.


Recommended Minimum Space for Commonly Used Laboratory Rodents Housed in Groups.

Studies have recently evaluated space needs and the effects of social housing, group size, and density (Andrade and Guimaraes 2003; Bartolomucci et al. 2002, 2003; Georgsson et al. 2001; Gonder and Laber 2007; Perez et al. 1997;A.L. Smith et al. 2004), age (Arakawa 2005; Davidson et al. 2007; Yildiz et al. 2007), and housing conditions (Gordon et al. 1998; Van Loo et al. 2004) for many different species and strains of rodents, and have reported varying effects on behavior (such as aggression) and experimental outcomes (Karolewicz and Paul 2001; Laber et al. 2008; McGlone et al. 2001; Rock et al. 1997; Smith et al. 2005; Van Loo et al. 2001). However, it is difficult to compare these studies due to the study design and experimental variables that have been measured. For example, variables that may affect the animals’ response to different cage sizes and housing densities include, but are not limited to, species, strain (and social behavior of the strain), phenotype, age, gender, quality of the space (e.g., vertical access), and structures placed in the cage. These issues remain complex and should be carefully considered when housing rodents.

Other Common Laboratory Animals Tables 3.3 and 3.4 list recommended minimum space for other common laboratory animals and for avian species. These allocations are based, in general, on the needs of pair- or group-housed animals. Space allocations should be reevaluated to provide for enrichment or to accommodate animals that exceed the weights in the tables, and should be based on species characteristics, behavior, compatibility of the animals, number of animals, and goals of the housing situation (Held et al. 1995; Lupo et al. 2000; Raje 1997; Turner et al. 1997). Singly housed animals may require more space per animal than that recommended for group-housed animals, while larger groups may be housed at slightly higher densities. For cats, dogs, and some rabbits, housing enclosures that allow greater freedom of movement and less restricted vertical space are preferred (e.g., kennels, runs, or pens instead of cages). Dogs and cats, especially when housed individually or in smaller enclosures (Bayne 2002), should be allowed to exercise and provided with positive human interaction. Species-specific plans for housing and management should be developed. Such plans should also include strategies for environmental enrichment.

TABLE 3.3. Recommended Minimum Space for Rabbits, Cats, and Dogs Housed in Pairs or Groups.


Recommended Minimum Space for Rabbits, Cats, and Dogs Housed in Pairs or Groups.

TABLE 3.4. Recommended Minimum Space for Avian Species Housed in Pairs or Groups.


Recommended Minimum Space for Avian Species Housed in Pairs or Groups.

Nonhuman Primates The recommended minimum space for nonhuman primates detailed in Table 3.5 is based on the needs of pair- or group-housed animals. Like all social animals, nonhuman primates should normally have social housing (i.e., in compatible pairs or in larger groups of compatible animals) (Hotchkiss and Paule 2003; NRC 1998a; Weed and Watson 1998; Wolfensohn 2004). Group composition is critical and numerous species-specific factors such as age, behavioral repertoire, sex, natural social organization, breeding requirements, and health status should be taken into consideration when forming a group. In addition, due to conformational differences of animals within groups, more space or height may be required to meet the animals’ physical and behavioral needs. Therefore, determination of the appropriate cage size is not based on body weight alone, and professional judgment is paramount in making such determinations (Kaufman et al. 2004; Williams et al. 2000).

TABLE 3.5. Recommended Minimum Space for Nonhuman Primates Housed in Pairs or Groups.


Recommended Minimum Space for Nonhuman Primates Housed in Pairs or Groups.

If it is necessary to house animals singly—for example, when justified for experimental purposes, for provision of veterinary care, or for incompatible animals—this arrangement should be for the shortest duration possible. If single animals are housed in small enclosures, an opportunity for periodic release into larger enclosures with additional enrichment items should be considered, particularly for animals housed singly for extended periods of time. Singly housed animals may require more space per animal than recommended for pair- or group-housed animals, while larger groups may be housed at slightly higher densities. Because of the many physical and behavioral characteristics of nonhuman primate species and the many factors to consider when using these animals in a biomedical research setting, species-specific plans for housing and management should be developed. Such plans should include strategies for environmental and psychological enrichment.

Agricultural Animals Table 3.6 lists recommended minimum space for agricultural animals commonly used in a laboratory setting. As social animals, they should be housed in compatible pairs or larger groups of compatible animals. When animals exceed the weights in the table, more space is required. For larger animals (particularly swine) it is important that the configuration of the space allow the animals to turn around and move freely (Becker et al. 1989; Bracke et al. 2002). Food troughs and water devices should be provided in sufficient numbers to allow ready access for all animals. Singly housed animals may require more space than recommended in the table to enable them to turn around and move freely without touching food or water troughs, have ready access to food and water, and have sufficient space to comfortably rest away from areas soiled by urine and feces.

TABLE 3.6. Recommended Minimum Space for Agricultural Animals.


Recommended Minimum Space for Agricultural Animals.

Terrestrial Management

Behavioral and Social Management

Activity Animal Activity typically implies motor activity but also includes cognitive activity and social interaction. Animals’ natural behavior and activity profile should be considered during evaluation of suitable housing or behavioral assessment.

Animals maintained in a laboratory environment are generally restricted in their activities compared to free-ranging animals. Forced activity for reasons other than attempts to meet therapeutic or approved protocol objectives should be avoided. High levels of repetitive, unvarying behavior (stereotypies, compulsive behaviors) may reflect disruptions of normal behavioral control mechanisms due to housing conditions or management practices (Garner 2005; NRC 1998a).

Dogs, cats, rabbits, and many other animals benefit from positive human interaction (Augustsson et al. 2002; Bayne et al. 1993; McCune 1997; Poole 1998; Rennie and Buchanan-Smith 2006; Rollin 1990). Dogs can be given additional opportunities for activity by being walked on a leash, having access to a run, or being moved into areas for social contact, play, or exploration (Wolff and Rupert 1991). Loafing areas, exercise lots, and pastures are suitable for large farm animals, such as sheep, horses, and cattle.

Social Environment Appropriate social interactions among members of the same species (conspecifics) are essential to normal development and well-being (Bayne et al. 1995; Hall 1998; Novak et al. 2006). When selecting a suitable social environment, attention should be given to whether the animals are naturally territorial or communal and whether they should be housed singly, in pairs, or in groups. An understanding of species-typical natural social behavior (e.g., natural social composition, population density, ability to disperse, familiarity, and social ranking) is key to successful social housing.

Not all members of a social species are necessarily socially compatible. Social housing of incompatible animals can induce chronic stress, injury, and even death. In some species, social incompatibility may be sex biased; for example, male mice are generally more prone to aggression than female mice, and female hamsters are generally more aggressive than male hamsters. Risks of social incompatibility are greatly reduced if the animals to be grouped are raised together from a young age, if group composition remains stable, and if the design of the animals’ enclosure and their environmental enrichment facilitate the avoidance of social conflicts. Social stability should be carefully monitored; in cases of severe or prolonged aggression, incompatible individuals need to be separated.

For some species, developing a stable social hierarchy will entail antagonistic interactions between pair or group members, particularly for animals introduced as adults. Animals may have to be introduced to each other over a period of time and should be monitored closely during this introductory period and thereafter to ensure compatibility.

Single housing of social species should be the exception and justified based on experimental requirements or veterinary-related concerns about animal well-being. In these cases, it should be limited to the minimum period necessary, and where possible, visual, auditory, olfactory, and tactile contact with compatible conspecifics should be provided. In the absence of other animals, enrichment should be offered such as positive interaction with the animal care staff and additional enrichment items or addition of a companion animal in the room or housing area. The need for single housing should be reviewed on a regular basis by the IACUC and veterinarian.

Procedural Habituation and Training of Animals Habituating animals to routine husbandry or experimental procedures should be encouraged whenever possible as it may assist the animal to better cope with a captive environment by reducing stress associated with novel procedures or people. The type and duration of habituation needed will be determined by the complexity of the procedure. In most cases, principles of operant conditioning may be employed during training sessions, using progressive behavioral shaping, to induce voluntary cooperation with procedures (Bloomsmith et al. 1998; Laule et al. 2003; NRC 2006a; Reinhardt 1997).


Food Animals should be fed palatable, uncontaminated diets that meet their nutritional and behavioral needs at least daily, or according to their particular requirements, unless the protocol in which they are being used requires otherwise. Subcommittees of the National Research Council Committee on Animal Nutrition have prepared comprehensive reports of the nutrient requirements of laboratory animals (NRC 1977, 1982, 1993, 1994, 1995a, 1998b, 2000, 2001, 2003a, 2006b, 2007); these publications consider issues of quality assurance, freedom from chemical or microbial contaminants and natural toxicants in feedstuffs, bioavailability of nutrients in feeds, and palatability.

There are several types of diets classified by the degree of refinement of their ingredients. Natural-ingredient diets are formulated with agricultural products and byproducts and are commercially available for all species commonly used in the laboratory. Although not a significant factor in most instances, the nutrient composition of ingredients varies, and natural ingredients may contain low levels of naturally occurring or artificial contaminants (Ames et al. 1993; Knapka 1983; Newberne 1975; NRC 1996; Thigpen et al. 1999, 2004). Contaminants such as pesticide residues, heavy metals, toxins, carcinogens, and phytoestrogens may be at levels that induce few or no health sequelae yet may have subtle effects on experimental results (Thigpen et al. 2004). Certified diets that have been assayed for contaminants are commercially available for use in select studies, such as preclinical toxicology, conducted in compliance with FDA Good Laboratory Practice standards (CFR 2009). Purified diets are refined such that each ingredient contains a single nutrient or nutrient class; they have less nutrient concentration variability and the potential for chemical contamination is lower. Chemically defined diets contain the most elemental ingredients available, such as individual amino acids and specific sugars (NRC 1996). The latter two types of diet are more likely to be used for specific types of studies in rodents but are not commonly used because of cost, lower palatability, and a reduced shelf life.

Animal colony managers should be judicious when purchasing, transporting, storing, and handling food to minimize the introduction of diseases, parasites, potential disease vectors (e.g., insects and other vermin), and chemical contaminants in animal colonies. Purchasers are encouraged to consider manufacturers’ and suppliers’ procedures and practices (e.g., storage, vermin control, and handling) for protecting and ensuring diet quality. Institutions should urge feed vendors to periodically provide data from laboratory-based feed analyses for critical nutrients. The user should know the date of manufacture and other factors that affect the food’s shelf life. Stale food or food transported and stored inappropriately can become deficient in nutrients. Upon receipt, bags of feed should be examined to ensure that they are intact and unstained to help ensure that their contents have not been potentially exposed to vermin, penetrated by liquids, or contaminated. Careful attention should be paid to quantities received in each shipment, and stock should be rotated so that the oldest food is used first.

Areas in which diets and diet ingredients are processed or stored should be kept clean and enclosed to prevent the entry of pests. Food stocks should be stored off the floor on pallets, racks, or carts in a manner that facilitates sanitation. Opened bags of food should be stored in vermin-proof containers to minimize contamination and to avoid the potential spread of pathogens. Exposure to elevated storage room temperatures, extremes in relative humidity, unsanitary conditions, and insects and other vermin hastens food deterioration. Storage of natural-ingredient diets at less than 21°C (70°F) and below 50% relative humidity is recommended. Precautions should be taken if perishable items—such as meats, fruits, and vegetables and some specialty diets (e.g., select medicated or high-fat diets)—are fed, because storage conditions may lead to variation in food quality.

Most natural-ingredient, dry laboratory animal diets stored properly can be used up to 6 months after manufacture. Nonstabilized vitamin C in manufactured feeds generally has a shelf life of only 3 months, but commonly used stabilized forms can extend the shelf life of feed. Refrigeration preserves nutritional quality and lengthens shelf life, but food storage time should be reduced to the lowest practical period and the manufacturers’ recommendations considered. Purified and chemically defined diets are often less stable than natural-ingredient diets and their shelf life is usually less than 6 months (Fullerton et al. 1982); they should be stored at 4°C (39°F) or lower.

Irradiated and fortified autoclavable diets are commercially available and are commonly used for axenic and microbiologically defined rodents, and immunodeficient animals (NRC 1996). The use of commercially fortified autoclavable diets ensures that labile vitamin content is not compromised by steam and/or heat (Caulfield et al. 2008; NRC 1996). But consideration should be given to the impact of autoclaving on pellets as it may affect their hardness and thus palatability and also lead to chemical alteration of ingredients (Thigpen et al. 2004; Twaddle et al. 2004). The date of sterilization should be recorded and the diet used quickly.

Feeders should be designed and placed to allow easy access to food and to minimize contamination with urine and feces, and maintained in good condition. When animals are housed in groups, there should be enough space and enough feeding points to minimize competition for food and ensure access to food for all animals, especially if feed is restricted as part of the protocol or management routine. Food storage containers should not be transferred between areas that pose different risks of contamination without appropriate treatment, and they should be cleaned and sanitized regularly.

Management of caloric intake is an accepted practice for long-term housing of some species, such as some rodents, rabbits, and nonhuman primates, and as an adjunct to some clinical, experimental, and surgical procedures (afor more discussion of food and fluid regulation as an experimental tool see Chapter 2 and NRC 2003a). Benefits of moderate caloric restriction in some species may include increased longevity and reproduction, and decreased obesity, cancer rates, and neurogenerative disorders (Ames et al. 1993; Colman et al. 2009; Keenan et al. 1994, 1996; Lawler et al. 2008; Weindruch and Walford 1988).

Under standard housing conditions, changes in biologic needs commensurate with aging should be taken into consideration. For example, there is good evidence that mice and rats with continuous access to food can become obese, with attendant metabolic and cardiovascular changes such as insulin resistance and higher blood pressure (Martin et al. 2010). These and other changes along with a more sedentary lifestyle and lack of exercise increase the risk of premature death (ibid.). Caloric management, which may affect physiologic adaptations and alter metabolic responses in a species-specific manner (Leveille and Hanson 1966), can be achieved by reducing food intake or by stimulating exercise.

In some species (e.g., nonhuman primates) and on some occasions, varying nutritionally balanced diets and providing “treats,” including fresh fruit and vegetables, can be appropriate and improve well-being. Scattering food in the bedding or presenting part of the diet in ways that require the animals to work for it (e.g., puzzle feeders for nonhuman primates) gives the animals the opportunity to forage, which, in nature, normally accounts for a large proportion of their daily activity. A diet should be nutritionally balanced; it is well documented that many animals offered a choice of unbalanced or balanced foods do not select a balanced diet and become malnourished or obese through selection of high-energy, low-protein foods (Moore 1987). Abrupt changes in diet, which can be difficult to avoid at weaning, should be minimized because they can lead to digestive and metabolic disturbances; these changes occur in omnivores and carnivores, but herbivores (Eadie and Mann 1970) are especially sensitive.

Water Animals should have access to potable, uncontaminated drinking water according to their particular requirements. Water quality and the definition of potable water can vary with locality (Homberger et al. 1993). Periodic monitoring for pH, hardness, and microbial or chemical contamination may be necessary to ensure that water quality is acceptable, particularly for use in studies in which normal components of water in a given locality can influence the results. Water can be treated or purified to minimize or eliminate contamination when protocols require highly purified water. The selection of water treatments should be carefully considered because many forms of water treatment have the potential to cause physiologic alterations, reduction in water consumption, changes in microflora, or effects on experimental results (Fidler 1977; Hall et al. 1980; Hermann et al. 1982; Homberger et al. 1993; NRC 1996).

Watering devices, such as drinking tubes and automated water delivery systems, should be checked frequently to ensure appropriate maintenance, cleanliness, and operation. Animals sometimes have to be trained to use automated watering devices and should be observed regularly until regular usage has been established to prevent dehydration. It is better to replace water bottles than to refill them, because of the potential for microbiologic cross contamination; if bottles are refilled, care should be taken to return each bottle to the cage from which it was removed. Automated watering distribution systems should be flushed or disinfected regularly. Animals housed in outdoor facilities may have access to water in addition to that provided in watering devices, such as that available in streams or in puddles after a heavy rainfall. Care should be taken to ensure that such accessory sources of water do not constitute a hazard, but their availability need not routinely be prevented. In cold weather, steps should be taken to prevent freezing of outdoor water sources.

Bedding and Nesting Materials Animal bedding and nesting materials are controllable environmental factors that can influence experimental data and improve animal well-being in most terrestrial species. Bedding is used to absorb moisture, minimize the growth of microorganisms, and dilute and limit animals’ contact with excreta, and specific bedding materials have been shown to reduce the accumulation of intracage ammonia (Perkins and Lipman 1995;E. Smith et al. 2004). Various materials are used as both contact and noncontact bedding; the desirable characteristics and methods of evaluating bedding have been described (Gibson et al. 1987; Jones 1977; Kraft 1980; Thigpen et al. 1989; Weichbrod et al. 1986). The veterinarian or facility manager, in consultation with investigators, should select the most appropriate bedding and nesting materials. A number of species, most notably rodents, exhibit a clear preference for specific materials (Blom et al. 1996; Manser et al. 1997, 1998; Ras et al. 2002), and mice provided with appropriate nesting material build better nests (Hess et al. 2008). Bedding that enables burrowing is encouraged for some species, such as mice and hamsters.

No type of bedding is ideal for all species under all management and experimental conditions. For example, in nude or hairless mice that lack eyelashes, some forms of paper bedding with fines (i.e., very small particles found in certain types of bedding) can result in periorbital abscesses (White et al. 2008), while cotton nestlets may lead to conjunctivitis (Bazille et al. 2001). Bedding can also influence mucosal immunity (Sanford et al. 2002) and endocytosis (Buddaraju and Van Dyke 2003).

Softwood beddings have been used, but the use of untreated softwood shavings and chips is contraindicated for some protocols because they can affect metabolism (Vesell 1967; Vesell et al. 1973, 1976). Cedar shavings are not recommended because they emit aromatic hydrocarbons that induce hepatic microsomal enzymes and cytotoxicity (Torronen et al. 1989; Weichbrod et al. 1986, 1988) and have been reported to increase the incidence of cancer (Jacobs and Dieter 1978; Vlahakis 1977). Prior treatment with high heat (kiln drying or autoclaving) may, depending on the material and the concentration of aromatic hydrocarbon constituents, reduce the concentration of volatile organic compounds, but the amounts remaining may be sufficient to affect specific protocols (Cunliffe-Beamer et al. 1981; Nevalainen and Vartiainen 1996).

The purchase of bedding products should take into consideration vendors’ manufacturing, monitoring, and storage methods. Bedding may be contaminated with toxins and other substances, bacteria, fungi, and vermin. It should be transported and stored off the floor on pallets, racks, or carts in a fashion consistent with maintenance of quality and avoidance of contamination. Bags should be stored sufficiently away from walls to facilitate cleaning. During autoclaving, bedding can absorb moisture and as a result lose absorbency and support the growth of microorganisms. Therefore, appropriate drying times and storage conditions should be used or, alternatively, gamma-irradiated materials if sterile bedding is indicated.

Bedding should be used in amounts sufficient to keep animals dry between cage changes, and, in the case of small laboratory animals, it should be kept from coming into contact with sipper tubes as such contact could cause leakage of water into the cage.

Sanitation Sanitation—the maintenance of environmental conditions conducive to health and well-being—involves bedding change (as appropriate), cleaning, and disinfection. Cleaning removes excessive amounts of excrement, dirt, and debris, and disinfection reduces or eliminates unacceptable concentrations of microorganisms. The goal of any sanitation program is to maintain sufficiently clean and dry bedding, adequate air quality, and clean cage surfaces and accessories.

The frequency and intensity of cleaning and disinfection should depend on what is necessary to provide a healthy environment for an animal. Methods and frequencies of sanitation will vary with many factors, including the normal physiologic and behavioral characteristics of the animals; the type, physical characteristics, and size of the enclosure; the type, number, size, age, and reproductive status of the animals; the use and type of bedding materials; temperature and relative humidity; the nature of the materials that create the need for sanitation; and the rate of soiling of the surfaces of the enclosure. Some housing systems or experimental protocols may require specific husbandry techniques, such as aseptic handling or modification in the frequency of bedding change.

Agents designed to mask animal odors should not be used in animal housing facilities. They cannot substitute for good sanitation practices or for the provision of adequate ventilation, and they expose animals to volatile compounds that might alter basic physiologic and metabolic processes.

Bedding/Substrate Change Soiled bedding should be removed and replaced with fresh materials as often as necessary to keep the animals clean and dry and to keep pollutants, such as ammonia, at a concentration below levels irritating to mucous membranes. The frequency of bedding change depends on multiple factors, such as species, number, and size of the animals in the primary enclosure; type and size of the enclosure; macro- and microenvironmental temperature, relative humidity, and direct ventilation of the enclosure; urinary and fecal output and the appearance and wetness of bedding; and experimental conditions, such as those of surgery or debilitation, that might limit an animal’s movement or access to clean bedding. There is no absolute minimal frequency of bedding changes; the choice is a matter of professional judgment and consultation between the investigator and animal care personnel. It typically varies from daily to weekly. In some instances frequent bedding changes are contraindicated; examples include portions of the pre- or postpartum period, research objectives that will be affected, and species in which scent marking is critical and successful reproduction is pheromone dependent.

Cleaning and Disinfection of the Microenvironment The frequency of sanitation of cages, cage racks, and associated equipment (e.g., feeders and watering devices) is governed to some extent by the types of caging and husbandry practices used, including the use of regularly changed contact or noncontact bedding, regular flushing of suspended catch pans, and the use of wire-bottom or perforated-bottom cages. In general, enclosures and accessories, such as tops, should be sanitized at least once every 2 weeks. Solid-bottom caging, bottles, and sipper tubes usually require sanitation at least once a week. Some types of cages and housing systems may require less frequent cleaning or disinfection; such housing may include large cages with very low animal density and frequent bedding changes, cages containing animals in gnotobiotic conditions with frequent bedding changes, individually ventilated cages, and cages used for special situations. Other circumstances, such as filter-topped cages without forced-air ventilation, animals that urinate excessively (e.g., diabetic or renal patients), or densely populated enclosures, may require more frequent sanitation.

The increased use of individually ventilated cages (IVCs) for rodents has led to investigations of the maintenance of a suitable microenvironment with extended cage sanitation intervals and/or increased housing densities (Carissimi et al. 2000; Reeb-Whitaker et al. 2001; Schondelmeyer et al. 2006). By design, ventilated caging systems provide direct continuous exchange of air, compared to static caging systems that depend on passive ventilation from the macroenvironment. As noted above, decreased sanitation frequency may be justified if the microenvironment in the cages, under the conditions of use (e.g., cage type and manufacturer, bedding, species, strain, age, sex, density, and experimental considerations), is not compromised (Reeb et al. 1998). Verification of microenvironmental conditions may include measurement of pollutants such as ammonia and CO2, microbiologic load, observation of the animals’ behavior and appearance, and the condition of bedding and cage surfaces.

Primary enclosures can be disinfected with chemicals, hot water, or a combination of both.2 Washing times and conditions and postwashing processing procedures (e.g., sterilization) should be sufficient to reduce levels or eliminate vegetative forms of opportunistic and pathogenic bacteria, adventitious viruses, and other organisms that are presumed to be controllable by the sanitation program. Disinfection from the use of hot water alone is the result of the combined effect of the temperature and the length of time that a given temperature (cumulative heat factor) is applied to the surface of the item. The same cumulative heat factor can be obtained by exposing organisms either to very high temperatures for short periods or to lower temperatures for longer periods (Wardrip et al. 1994, 2000). Effective disinfection can be achieved with wash and rinse water at 143–180°F or more. The traditional 82.2°C (180°F) temperature requirement for rinse water refers to the water in the tank or in the sprayer manifold. Detergents and chemical disinfectants enhance the effectiveness of hot water but should be thoroughly rinsed from surfaces before reuse of the equipment. Their use may be contraindicated for some aquatic species, as residue may be highly deleterious. Mechanical washers (e.g., cage and rack, tunnel, and bottle washers) are recommended for cleaning quantities of caging and movable equipment.

Sanitation of cages and equipment by hand with hot water and detergents or disinfectants can also be effective but requires considerable attention to detail. It is particularly important to ensure that surfaces are rinsed free of residual chemicals and that personnel have appropriate equipment to protect themselves from exposure to hot water or chemical agents used in the process.

Water bottles, sipper tubes, stoppers, feeders, and other small pieces of equipment should be washed with detergents and/or hot water and, where appropriate, chemical agents to destroy microorganisms. Cleaning with ultrasound may be a useful method for small pieces of equipment.

If automated watering systems are used, some mechanism to ensure that microorganisms and debris do not build up in the watering devices is recommended (Meier et al. 2008); the mechanism can be periodic flushing with large volumes of water or appropriate chemical agents followed by a thorough rinsing. Constant recirculation loops that use properly maintained filters, ultraviolet lights, or other devices to disinfect recirculated water are also effective. Attention should be given to the routine sanitation of automatic water delivery valves (i.e., lixits) during primary enclosure cleaning.

Conventional methods of cleaning and disinfection are adequate for most animal care equipment. However, it may be necessary to also sterilize caging and associated equipment to ensure that pathogenic or opportunistic microorganisms are not introduced into specific-pathogen-free or immuno-compromised animals, or that experimental biologic hazards are destroyed before cleaning. Sterilizers should be regularly evaluated and monitored to ensure their safety and effectiveness.

For pens or runs, frequent flushing with water and periodic use of detergents or disinfectants are usually appropriate to maintain sufficiently clean surfaces. If animal waste is to be removed by flushing, this will need to be done at least once a day. During flushing, animals should be kept dry. The timing of pen or run cleaning should take into account the normal behavioral and physiologic processes of the animals; for example, the gastrocolic reflex in meal-fed animals results in defecation shortly after food consumption.

Cleaning and Disinfection of the Macroenvironment All components of the animal facility, including animal rooms and support spaces (e.g., storage areas, cage-washing facilities, corridors, and procedure rooms) should be regularly cleaned and disinfected as appropriate to the circumstances and at a frequency based on the use of the area and the nature of likely contamination. Vaporized hydrogen peroxide or chlorine dioxide are effective compounds for room decontamination, particularly following completion of studies with highly infectious agents (Krause et al. 2001) or contamination with adventitious microbial agents.

Cleaning implements should be made of materials that resist corrosion and withstand regular sanitation. They should be assigned to specific areas and should not be transported between areas with different risks of contamination without prior disinfection. Worn items should be replaced regularly. The implements should be stored in a neat, organized fashion that facilitates drying and minimizes contamination or harborage of vermin.

Assessing the Effectiveness of Sanitation Monitoring of sanitation practices should fit the process and materials being cleaned and may include visual inspection and microbiologic and water temperature monitoring (Compton et al. 2004a,b; Ednie et al. 1998; Parker et al. 2003). The intensity of animal odors, particularly that of ammonia, should not be used as the sole means of assessing the effectiveness of the sanitation program. A decision to alter the frequency of cage bedding changes or cage washing should be based on such factors as ammonia concentration, bedding condition, appearance of the cage and animals, and the number and size of animals housed in the cage.

Mechanical washer function should be evaluated regularly and include examination of mechanical components such as spray arms and moving headers as well as spray nozzles to ensure that they are functioning appropriately. If sanitation is temperature dependent, the use of temperature-sensing devices (e.g., thermometers, probes, or temperature-sensitive indicator strips) is recommended to ensure that the equipment being sanitized is exposed to the desired conditions.

Whether the sanitation process is automated or manual, regular evaluation of sanitation effectiveness is recommended. This can be performed by evaluating processed materials by microbiologic culture or the use of organic material detection systems (e.g., adenosine triphosphate [ATP] bioluminescence) and/or by confirming the removal of artificial soil applied to equipment surfaces before washing.

Waste Disposal Conventional, biologic, and hazardous waste should be removed and disposed of regularly and safely (Hill 1999). There are several options for effective waste disposal. Contracts with licensed commercial waste disposal firms usually provide some assurance of regulatory compliance and safety. On-site incineration should comply with all federal, state, and local regulations (Nadelkov 1996).

Adequate numbers of properly labeled waste receptacles should be strategically placed throughout the facility. Waste containers should be leak-proof and equipped with tight-fitting lids. It is good practice to use disposable liners and to wash containers and implements regularly. There should be a dedicated waste storage area that can be kept free of insects and other vermin. If cold storage is used to hold material before disposal, a properly labeled, dedicated refrigerator, freezer, or cold room should be used that is readily sanitized.

Hazardous wastes must be rendered safe by sterilization, containment, or other appropriate means before their removal from the facility (DHHS 2009 or most recent edition; NRC 1989, 1995b). Radioactive wastes should be kept in properly labeled containers and their disposal closely coordinated with radiation safety specialists in accord with federal and state regulations; the federal government and most states and municipalities have regulations controlling disposal of hazardous wastes. Compliance with regulations concerning hazardous-agent use (see Chapter 2) and disposal is an institutional responsibility.

Infectious animal carcasses can be incinerated on site or collected by a licensed contractor. Use of chemical digesters (alkaline hydrolysis treatment) may be considered in some situations (Kaye et al. 1998; Murphy et al. 2009). Procedures for on-site packaging, labeling, transportation, and storage of these wastes should be integrated into occupational health and safety policies (Richmond et al. 2003).

Hazardous wastes that are toxic, carcinogenic, flammable, corrosive, reactive, or otherwise unstable should be placed in properly labeled containers and disposed of as recommended by occupational health and safety specialists. In some circumstances, these wastes can be consolidated or blended. Sharps and glass should be disposed of in a manner that will prevent injury to waste handlers.

Pest Control Programs designed to prevent, control, or eliminate the presence of or infestation by pests are essential in an animal environment. A regularly scheduled and documented program of control and monitoring should be implemented. The ideal program prevents the entry of vermin and eliminates their harborage in the facility (Anadon et al. 2009; Easterbrook et al. 2008). For animals in outdoor facilities, consideration should be given to eliminating or minimizing the potential risk associated with pests and predators.

Pesticides can induce toxic effects on research animals and interfere with experimental procedures (Gunasekara et al. 2008). They should be used in animal areas only when necessary and investigators whose animals may be exposed to them should be consulted beforehand. Use of pesticides should be recorded and coordinated with the animal care management staff and be in compliance with federal, state, or local regulations. Whenever possible, nontoxic means of pest control, such as insect growth regulators (Donahue et al. 1989; Garg and Donahue 1989; King and Bennett 1989; Verma 2002) and nontoxic substances (e.g., amorphous silica gel), should be used. If traps are used, methods should be humane; traps that catch pests alive require frequent observation and humane euthanasia after capture (Mason and Littin 2003; Meerburg et al. 2008).

Emergency, Weekend, and Holiday Care Animals should be cared for by qualified personnel every day, including weekends and holidays, both to safeguard their well-being and to satisfy research requirements. Emergency veterinary care must be available after work hours, on weekends, and on holidays.

In the event of an emergency, institutional security personnel and fire or police officials should be able to reach people responsible for the animals. Notification can be enhanced by prominently posting emergency procedures, names, or telephone numbers in animal facilities or by placing them in the security department or telephone center. Emergency procedures for handling special facilities or operations should be prominently posted and personnel trained in emergency procedures for these areas. A disaster plan that takes into account both personnel and animals should be prepared as part of the overall safety plan for the animal facility. The colony manager or veterinarian responsible for the animals should be a member of the appropriate safety committee at the institution, an “official responder” in the institution, and a participant in the response to a disaster (Vogelweid 1998).

Population Management

Identification Animal records are useful and variable, ranging from limited information on identification cards to detailed computerized records for individual animals (Field et al. 2007). Means of animal identification include room, rack, pen, stall, and cage cards with written, bar-coded, or radio frequency identification (RFID) information. Identification cards should include the source of the animal, the strain or stock, names and contact information for the responsible investigator(s), pertinent dates (e.g., arrival date, birth date, etc.), and protocol number when applicable. Genotype information, when applicable, should also be included, and consistent, unambiguous abbreviations should be used when the full genotype nomenclature (see below) is too lengthy.

In addition, the animals may wear collars, bands, plates, or tabs or be marked by colored stains, ear notches/punches and tags, tattoos, subcutaneous transponders, and freeze brands. As a method of identification of small rodents, toe-clipping should be used only when no other individual identification method is feasible. It may be the preferred method for neonatal mice up to 7 days of age as it appears to have few adverse effects on behavior and well-being at this age (Castelhano-Carlos et al. 2010; Schaefer et al. 2010), especially if toe clipping and genotyping can be combined. Under all circumstances aseptic practices should be followed. Use of anesthesia or analgesia should be commensurate with the age of the animals (Hankenson et al. 2008).

Recordkeeping Records containing basic descriptive information are essential for management of colonies of large long-lived animals and should be maintained for each animal (Dyke 1993; Field et al. 2007; NRC 1979a). These records often include species, animal identifier, sire and/or dam identifier, sex, birth or acquisition date, source, exit date, and final disposition. Such animal records are essential for genetic management and historical assessments of colonies. Records of rearing and housing histories, mating histories, and behavioral profiles are useful for the management of many species, especially nonhuman primates (NRC 1979a). Relevant recorded information should be provided when animals are transferred between institutions.

Medical records for individual animals can also be valuable, especially for dogs, cats, nonhuman primates, and agricultural animals (Suckow and Doerning 2007). They should include pertinent clinical and diagnostic information, date of inoculations, history of surgical procedures and postoperative care, information on experimental use, and necropsy findings where applicable.

Basic demographic information and clinical histories enhance the value of individual animals for both breeding and research and should be readily accessible to investigators, veterinary staff, and animal care staff.

Breeding, Genetics, and Nomenclature Genetic characteristics are important with regard to the selection and management of animals for use in breeding colonies and in biomedical research (see Appendix A). Pedigree information allows appropriate selection of breeding pairs and of experimental animals that are unrelated or of known relatedness.

Outbred animals are widely used in biomedical research. Founding populations should be large enough to ensure the long-term genetic heterogeneity of breeding colonies. To facilitate direct comparison of research data derived from outbred animals, genetic management techniques should be used to maintain genetic variability and equalize founder representations (Hartl 2000; Lacy 1989; Poiley 1960; Williams-Blangero 1991). Genetic variability can be monitored with computer simulations, biochemical markers, DNA markers and sequencing, immunologic markers, or quantitative genetic analyses of physiologic variables (MacCluer et al. 1986; Williams-Blangero 1993).

Inbred strains of various species, especially rodents, have been developed to address specific research needs (Festing 1979; Gill 1980). When inbred animals or their F1 progeny are used, it is important to periodically monitor genetic authenticity (Festing 1982; Hedrich 1990); several methods of monitoring have been developed that use immunologic, biochemical, and molecular techniques (Cramer 1983; Festing 2002; Groen 1977; Hoffman et al. 1980; Russell et al. 1993). Appropriate management systems (Green 1981; Kempthorne 1957) should be designed to minimize genetic contamination resulting from mutation and mismating.

Genetically modified animals (GMAs) represent an increasingly large proportion of animals used in research and require special consideration in their population management. Integrated or altered genes can interact with species or strain-specific genes, other genetic manipulations, and environmental factors, in part as a function of site of integration, so each GMA line can be considered a unique resource. Care should be taken to preserve such resources through standard genetic management procedures, including maintenance of detailed pedigree records and genetic monitoring to verify the presence and zygosity of transgenes and other genetic modifications (Conner 2005). Cryopreservation of fertilized embryos, ova, ovaries, or spermatozoa should also be considered as a safeguard against alterations in transgenes over time or accidental loss of GMA lines (Conner 2002; Liu et al. 2009).

Generation of animals with multiple genetic alterations often involves crossing different GMA lines and can lead to the production of offspring with genotypes that are not of interest to the researcher (either as experimental or control animals) as well as unexpected phenotypes. Carefully designed breeding strategies and accurate genotype assessment can help to minimize the generation of animals with unwanted genotypes (Linder 2003). Newly generated genotypes should be carefully monitored and new phenotypes that negatively affect well-being should be reported to the IACUC and managed in a manner to ensure the animals’ health and well-being.

Accurate recording, with standardized nomenclature when available, of both the strain and substrain or of the genetic background of animals used in a research project is important (NRC 1979b). Several publications provide rules developed by international committees for standardized nomenclature of outbred rodents and rabbits (Festing et al. 1972), inbred rats, inbred mice, and transgenic animals (FELASA 2007; Linder 2003). In addition, the International Committee on Standardized Genetic Nomenclature for Mice and the Rat Genome and Nomenclature Committee maintain online guidelines for these species (MGI 2009).


The variety of needs for fish and aquatic or semiaquatic reptiles and amphibians is as diverse as the number of species considered. This section is intended to provide facility managers, veterinarians, and IACUCs with basic information related to the management of aquatic animal systems (Alworth and Harvey 2007; Alworth and Vazquez 2009; Browne et al. 2007; Browne and Zippel 2007; Denardo 1995; DeTolla et al. 1995; Koerber and Kalishman 2009; Lawrence 2007; Matthews et al. 2002; Pough 2007). Specific recommendations are available in texts and journal reviews, and it will be necessary to review other literature and consult with experienced caregivers for further detail on caring for aquatic species (see Appendix A).

Aquatic Environment

Microenvironment and Macroenvironment

As with terrestrial systems, the microenvironment of an aquatic animal is the physical environment immediately surrounding it—the primary enclosure such as the tank, raceway, or pond. It contains all the resources with which the animals are in direct contact and also provides the limits of the animals’ immediate environment. The microenvironment is characterized by many factors, including water quality, illumination, noise, vibration, and temperature. The physical environment of the secondary enclosure, such as a room, constitutes the macroenvironment.

Water Quality

The composition of the water (water quality) is essential to aquatic animal well-being, although other factors that affect terrestrial microenvironments are also relevant. Water quality parameters and life support systems for aquatic animals will vary with the species, life stage, the total biomass supported, and the animals’ intended use (Blaustein et al. 1999; Fisher 2000; Gresens 2004; Overstreet et al. 2000; Schultz and Dawson 2003). The success and adequacy of the system depend on its ability to match the laboratory habitat to the natural history of the species (Godfrey and Sanders 2004; Green 2002; Lawrence 2007; Spence et al. 2008).

Characteristics of the water that may affect its appropriateness include temperature, pH, alkalinity, nitrogen waste products (ammonia, nitrite, and nitrate), phosphorus, chlorine/bromine, oxidation-reduction potential, conductivity/salinity, hardness (osmolality/dissolved minerals), dissolved oxygen, total gas pressure, ion and metal content, and the established microbial ecology of the tank. Water quality parameters can directly affect animal well-being; different classes, species, and ages in a species may have different water quality needs and sensitivities to changes in water quality parameters.

Routine measurement of various water characteristics (water quality testing) is essential for stable husbandry. Standards for acceptable water quality, appropriate parameters to test, and testing frequency should be identified at the institutional level and/or in individual animal use protocols depending on the size of the aquatic program. Staff managing aquatic systems need to be trained in biologically relevant aspects of water chemistry, how water quality parameters may affect animal health and well-being, how to monitor water quality results, and how water quality may affect life support system function (e.g., biologic filtration).

The specific parameters and frequency of testing vary widely (depending on the species, life stage, system, and other factors), from continuous monitoring to infrequent spot checks. Recently established systems and/or populations, or changes in husbandry procedures, may require more frequent assessment as the system ecology stabilizes; stable environments may require less frequent testing. Toxins from system components, particularly in newly constructed systems, may require special consideration such as leaching of chemicals from construction materials, concrete, joint compounds, and sealants (DeTolla et al. 1995; Nickum et al. 2004). Chlorine and chloramines used to disinfect water for human consumption or to disinfect equipment are toxic to fish and amphibians and must be removed or neutralized before use in aquatic systems (Tompkins and Tsai 1976; Wedemeyer 2000).

Life Support System

The phrase life support system refers to the physical structure used to contain the water and the animals as well as the ancillary equipment used to move and/or treat the water. Life support systems may be simple (e.g., a container to hold the animal and water) or extremely complex (e.g., a fully automated recirculating system). The type of life support system used depends on several factors including the natural habitat of the species, age/size of the species, number of animals maintained, availability and characteristics of the water required, and the type of research.

Life support systems typically fall into three general categories: recirculating systems where water (all or part) is moved around a system, flow-through systems where water is constantly replaced, or static systems where water is stationary and periodically replenished or replaced. The water may be fresh, brackish, or salt and is maintained at specific temperatures depending on the species’ needs.

The source of water for these systems typically falls into four general categories: treated wastewater (e.g., municipal tap water), surface water (e.g., rivers, lakes, or oceans), protected water (e.g., well or aquifer water), or artificial water (e.g., reverse osmosis or distilled water). Artificial saltwater may be created by adding appropriate salt to freshwater sources. Source water selection should be based on the provision of a consistent or constant supply, incoming biosecurity level requirements, water volumes needed, species selection, and research considerations.

Recirculating systems are common in indoor research settings where high-density housing systems are often needed. Most recirculating systems are designed to exchange a specific volume of water per unit time and periodically introduce fresh water into the system. These systems are the most mechanically advanced, containing biologic filters (biofilters) that promote conversion of ammonia to nitrite and nitrate via nitrifying bacteria, protein skimmers (foam fractionators) and particulate filters to remove undissolved and dissolved proteins and particulate matter, carbon filters to remove dissolved chemicals, and ultraviolet or ozone units to disinfect the water. The systems generally contain components to aerate and degas the water (to prevent gas oversaturation) and to heat or cool it, as well as automated dosing systems to maintain appropriate pH and conductivity. Not all elements are present in all systems and some components may accomplish multiple functions. Recirculating systems may be designed so that multiple individual tanks are supplied with treated water from a single source, as is the case with “rack” systems used for zebrafish (Danio rerio) and Xenopus laevis and X. tropicalis, as examples (Fisher 2000; Koerber and Kalishman 2009; Schultz and Dawson 2003).

The development and maintenance of the biofilter is critical for limiting ammonia and nitrite accumulation in recirculating systems. The biofilter must be of sufficient size (i.e., contain a sufficient quantity of bacteria) to be capable of processing the bioload (level of nitrogenous waste) entering the system. The microorganisms supported by the biofilter require certain water quality parameters. Alterations in the aquatic environment (e.g., rapid changes in salinity, temperature, and pH) as well as the addition of chemicals or antimicrobials may significantly affect the microbial ecology of the system and therefore water quality and animal well-being. If damaged, biofilter recovery may take weeks (Fisher 2000). Changes in water quality parameters (e.g., pH, ammonia, and nitrite) may negatively affect animal health and the efficiency of the biofilter, so species sensitive to change in water quality outside of a narrow range require more frequent monitoring.

Continuous or timed flow-through systems can be used where suitable water is available to support the species to be housed (e.g., in aquaculture facilities). These systems may use extremely large volumes of water as it is not reused. The water may be used “as is” or processed before use, for example by removing sediments, excessive dissolved gases, chlorine, or chloramines, and by disinfecting with UV or ozone (Fisher 2000; Overstreet et al. 2000). Static systems vary in size from small tanks to large inground ponds, and may use mechanical devices to move and aerate water.

Temperature, Humidity, and Ventilation

The general concepts discussed in the Terrestrial Animals section also apply to the aquatic setting. Most aquatic or semiaquatic species (fish, amphibians, and reptiles) used in research are poikilotherms, which depend, for the most part, on the temperature of their environment to sustain physiologic processes, such as metabolism, reproduction, and feeding behavior (Browne and Edwards 2003; Fraile et al. 1989; Maniero and Carey 1997; Pough 1991). Temperature requirements are based on the natural history of the species and can vary depending on life stage (Green 2002; Pough 1991; Schultz and Dawson 2003). Water temperature may be controlled at its source, within the life support system, or by controlling the macroenvironment. Some semiopen systems (e.g., raceways by a river) depend on source water temperature and thus enclosure water temperature will vary with that of the source water.

The volume of water contained in a room can affect room temperature, temperature stability, and relative humidity. Likewise the thermal load produced by chiller/heater systems can affect the stability of the macroenvironmental temperature. Air handling systems need to be designed to compensate for these thermal and moisture loads. Macroenvironmental relative humidity levels are generally defined by safety issues and staff comfort, since room humidity is not critical for aquatic species; however, excessive moisture may result in condensation on walls, ceilings, and tank lids, which may support microbial growth and serve as a source of contamination or create a conducive environment for metal corrosion. In a dry environment (e.g., indoor heating during cold weather or outdoor housing in some climates/seasons), evaporation rates may be higher, potentially requiring the addition of large quantities of water to the system and monitoring for increases in salinity/conductivity, contaminants, or other water quality aberrations. Some amphibians and reptiles may need elevated microenvironmental humidity (in excess of 50–70% relative humidity), which may require maintaining elevated macroenvironmental humidity levels (Pough 1991; St. Claire et al. 2005).

Room air exchange rates are typically governed by thermal and moisture loads. For fish and some aquatic amphibians, the microenvironmental air quality may affect water quality (i.e., gas exchange), but appropriate life support system design may reduce its importance. Airborne particulates and compounds (e.g., volatile organic compounds and ammonia) may dissolve in tank water and affect animal health (Koerber and Kalishman 2009). As the aerosolization of water can lead to the spread of aquatic animal pathogens (e.g., protozoa, bacteria) within or throughout an aquatic animal facility, this process should be minimized as much as possible (Roberts-Thomson et al. 2006; Wooster and Bowser 2007; Yanong 2003).


Aquatic and semiaquatic species are often sensitive to changes in photoperiod, light intensity, and wavelength (Brenner and Brenner 1969). Lighting characteristics will vary by species, their natural history, and the research being conducted. Gradual changes in room light intensity are recommended, as rapid changes in light intensity can elicit a startle response in fish and may result in trauma. Some aquatic and semiaquatic species may need full-spectrum lighting and/or heat lamps to provide supplemental heating to facilitate adequate physiological function (e.g., aquatic turtles provided with a basking area; Pough 1991).

Noise and Vibration

General concepts discussed in the Terrestrial Animals section apply to aquatic animals. These animals may be sensitive to noise and vibration, which are readily transmitted through water. Species vary in their response and many fish species acclimate to noise and vibration, although these may cause subclinical effects (Smith et al. 2007). Vibration through floors can be reduced by using isolation pads under aquaria racks. Some facilities elect to place major components of the life support system (e.g., filters, pumps, and biofilters) outside the animal rooms to reduce vibration and noise.

Aquatic Housing

Microenvironment (Primary Enclosure)

The primary enclosure (a tank, raceway, pond, or pen holding water and the animal) defines the limits of an animal’s immediate environment. In research settings, acceptable primary enclosures

  • allow for the normal physiological and behavioral needs of the animals, including excretory function, control and maintenance of body temperature, normal movement and postural adjustments, and, where indicated, reproduction. In some poikilothermic reptiles and amphibians, microenvironmental temperature gradients may be needed for certain physiologic functions such as feeding and digestion.
  • allow conspecific social interactions (e.g., schooling in fish species).
  • provide a balanced, stable environment that supports the animal’s physiologic needs.
  • provide the appropriate water quality and characteristics, and permit monitoring, filling, refilling, and changing of water.
  • allow access to adequate food and removal of food waste.
  • restrict escape or accidental entrapment of animals or their appendages.
  • are free of sharp edges and/or projections that could cause injury.
  • allow for observation of the animals with minimal disturbance.
  • are constructed of nontoxic materials that do not leach toxicants or chemicals into the aquatic environment.
  • do not present electrical hazards directly or indirectly.

Environmental Enrichment and Social Housing

Environmental enrichment strategies for many aquatic species are not well established. The implications of a barren versus an enriched environment on well-being, general research, growth, and development are unknown or poorly defined, as is true of individual versus group (social) housing for many species. When used, enrichment should elicit species-appropriate behaviors and be evaluated for safety and utility.

Generally, schooling fish species are housed with conspecifics, and many amphibians, especially anuran species, may be group housed. Aggression in aquatic animals does occur (van de Nieuwegiessen et al. 2008; Speedie and Gerlai 2008) and, as for terrestrial animals, appropriate monitoring and intervention may be necessary (Matthews et al. 2002; Torreilles and Green 2007). Some species need appropriate substrate (e.g., gravel) to reproduce or need substrate variety to express basic behaviors and maintain health (Overstreet et al. 2000). Improved breeding success in enriched environments has been reported but further research in this area is needed (Carfagnini et al. 2009). For many species (including, e.g., X. laevis), visual barriers, hides, and shading are appropriate (Alworth and Vasquez 2009; Torreilles and Green 2007). Most semiaquatic reptiles spend some time on land (basking, feeding, digesting, and ovipositing) and terrestrial areas should be provided as appropriate.

Sheltered, Outdoor, and Naturalistic Housing

Animals used in aquaculture are often housed in situations that mimic agricultural rearing and may be in outdoor and/or sheltered raceways, ponds, or pens with high population densities. In these settings, where natural predation and mortalities occur, it may be appropriate to measure animal “numbers” by using standard aquaculture techniques such as final production biomass (Borski and Hodson 2003).


Space recommendations and housing density vary extensively with the species, age/size of the animals, life support system, and type of research (Browne et al. 2003; Green 2009; Gresens 2004; Hilken et al. 1995; Matthews et al. 2002). In the United States, for example, adult zebrafish (Danio rerio) in typical biomedical research settings are generally housed 5 adult fish per liter of water (Matthews et al. 2002), but this housing density varies when breeding and for housing younger animals (Matthews et al. 2002). This guidance is not necessarily relevant for other species of fish, and may change as research advances (Lawrence 2007). X. laevis adults may be housed at 2 liters of water per frog (NRC 1974), but a wide variety of housing systems are currently used in research settings (Green 2009). Institutions, investigators, and IACUC members should evaluate the appropriate needs of each species during program evaluations and facility inspections and continue to review ongoing research in these areas.

Aquatic Management

Behavior and Social Management

Visual evaluations of aquatic and semiaquatic animals are typically used for monitoring. To avoid damage to the protective mucus layers of the skin and negative effects on immune function (De Veer et al. 2007; Subramanian et al. 2007; Tsutsui et al. 2005), handling of these species should be kept to the minimum required (Bly et al. 1997). Appropriate handling techniques vary widely depending on the species, age/size, holding system, and specific research need (Fisher 2000; Matthews et al. 2002; Overstreet et al. 2000); they should be identified at the facility or individual protocol level.

Latex gloves have been associated with toxicity in some amphibians (Gutleb et al. 2001). The use of appropriate nets by well-trained personnel can reduce skin damage and thus stress. Nets should be cleaned and disinfected appropriately when used in different systems and should be dedicated to animals of similar health status whenever possible.

Exercise and activity levels for aquatic species are minimally described but informed decisions may be extrapolated from studies of behavior of the same or similar species in the wild (Spence et al. 2008). Some aquatic species do not rest and constantly swim; others may rest all or a significant portion of the day. Water flow rates and the provision of hides or terrestrial resting platforms (e.g., for some reptiles and amphibians) need to be appropriate for species and life stage.


Food The general principles relating to feeding of terrestrial animals are applicable to aquatic animals. Food should be stored in a type-appropriate manner to preserve nutritional content, minimize contamination, and prevent entry of pests. Food delivery methods should ensure that all animals are able to access food for a sufficient period of time while minimizing feeding aggression and nutrient loss. Feeding methods and frequency vary widely depending on the species, age/size of species, and type of life support system. Many aquatic or semiaquatic species are not provided with food ad libitum in the tank, and in some cases may not be fed daily.

Commercial diets (e.g., pellets, flakes) are available for certain species and storage time should be based on manufacturer recommendations or follow commonly accepted practices. In aquatic systems, particularly in fish rearing or when maintaining some amphibian and reptile species, the use of live foods (e.g., Artemia sp. larva, crickets, or mealworm beetle larvae) is common. Live food sources need to be maintained and managed to ensure a steady supply and the health and suitability of the organism as a food. Care should be taken to feed a complete diet to avoid nutritional deficiencies.

Water (see also section on Water Quality) Aquatic animals need access to appropriately conditioned water. Fully aquatic animals obtain water in their habitat or absorb it across their gills or skin. Some semiaquatic amphibians and reptiles may need “bowls” of water for soaking and drinking, and water quality should be appropriate (see Terrestrial Animals section). Chlorine or chloramines may be present in tap water at levels that could be toxic to some species.

Substrate Substrates can provide enrichment for aquatic animals by promoting species-appropriate behavior such as burrowing, foraging, or enhanced spawning (Fisher 2000; Matthews et al. 2002; Overstreet et al. 2000). They may be an integral and essential component of the life support system by providing increased surface area for denitrifying bacteria (e.g., systems with undergravel filtration), and need routine siphoning (i.e., hydrocleaning) to remove organic debris. System design and species needs should be evaluated to determine the amount, type, and presentation of substrate.

Sanitation Sanitation of the aquatic environment in recirculating systems is provided through an appropriately designed and maintained life support system, regular removal of solid waste materials from the enclosure bottom, and periodic water changes. The basic concept of sanitation (i.e., to provide conditions conducive to animal health and welfare) is the same for terrestrial and aquatic systems. However, sanitation measures in aquatic systems differ from those for terrestrial systems because much of the nitrogenous waste (feces and urine) and respiratory output (carbon dioxide) is dissolved in the water.

A properly functioning life support system, designed to process the bioload, will maintain nitrogenous wastes within an acceptable range. Solids may be removed in a variety of ways, depending on the design of the system; generally they are removed by siphoning (hydrocleaning) and/or filtration. Depending on the type, filters need routine cleaning or replacement or, if self-cleaning, proper maintenance; in saltwater systems dissolved proteins may be removed by protein skimmers. Reducing organic solids limits the quantities of nitrogen and phosphorus that need to be removed from the system, both of which can accumulate to levels that are toxic to fish and amphibians. The biologic filter (denitrifying bacteria) typically removes ammonia and nitrite, potential toxins, from aquatic systems. Nitrate, the end product of this process, is less toxic to aquatic animals but at high levels can be problematic; it is generally removed through water changes, although large systems may have a specialized denitrification unit to reduce levels.

Disinfection is usually accomplished through water treatment (e.g., filtration and application of UV light or ozone) and/or water changes. Chlorine and most chemical disinfectants are inappropriate for aquatic systems containing animals as they are toxic at low concentrations; when used to disinfect an entire system or system components, extreme care must be taken to ensure that residual chlorine, chemical, and reactive byproducts are neutralized or removed. The type of monitoring and frequency varies depending on the disinfection method, the system, and the animals.

Algal growth is common in aquatic systems and increases with the presence of nitrogen and phosphorus, particularly in the presence of light. Excessive growth may be an indication of elevated nitrogen or phosphorus levels. Algal species seen with recirculating systems are generally nontoxic, although species capable of producing toxins exist. Algae are typically removed using mechanical methods (i.e., scrubbing or scraping). Limiting algal growth is important to allow viewing of the animals in the enclosure. Cyanobacteria (commonly called blue-green algae) growth is also possible and may be common in freshwater aquaculture. The same factors that promote algae growth also promote cyanobacteria growth. As with algae, while most species are harmless, some species can produce clinically relevant toxic compounds (Smith et al. 2008).

Tank (cage) changing and disinfection are conducted at frequencies using methods that often differ from terrestrial systems. Because waste is dissolved in the water and/or removed as solids by siphoning or filtration, regular changing of tanks is not integral to maintaining adequate hygiene in typical aquatic systems. The frequency of cleaning and disinfection should be determined by water quality, which should permit adequate viewing of the animals, and animal health monitoring. System components such as lids on fish tanks, which may accumulate feed, may require sanitation as often as weekly depending on the frequency and type of feed and the system’s design.

Cleaning and Disinfection of the Macroenvironment As with terrestrial systems, all components of the animal facility, including animal rooms and support spaces (e.g., storage areas, cage-washing facilities, corridors, and procedure rooms), should be regularly cleaned and disinfected as appropriate to the circumstances and at a frequency determined by the use of the area and the nature of likely contamination. Cleaning agents should be chosen and used with care to ensure there is no secondary contamination of the aquatic systems.

Cleaning implements should be made of materials that resist corrosion and withstand regular sanitation. They should be assigned to specific areas and should not be transported between areas with different risks of contamination without prior disinfection. Worn items should be replaced regularly. The implements should be stored in a neat, organized fashion that facilitates drying and minimizes contamination or harborage of vermin.

Waste Disposal Wastewater treatment and disposal may be necessary in some facilities depending on water volume, quality, and chemical constituents. Local regulations may limit or control the release of wastewater.

Pest Control Terrestrial animal pest control principles apply to aquatic systems but, due to transcutaneous absorption, aquatic and semiaquatic species may be more sensitive to commonly used pest control agents than terrestrial animals. Before use, an appropriate review of chemicals and methods of application is necessary.

Emergency, Weekend, and Holiday Care As with terrestrial species, aquatic animals should receive daily care from qualified personnel who have a sufficient understanding of the housing system to identify malfunctions and, if they are unable to address a system failure of such magnitude that it requires resolution before the next workday, access to staff who can respond to the problem. Automated monitoring systems are available and may be appropriate depending on system size and complexity. Appropriate emergency response plans should be developed to address major system failures.

Population Management

Identification Identification principles are similar to those for terrestrial animals. Identification criteria are based on the species and housing system. Identification methods available for use in aquatic species include fin clipping, genetic testing (Matthews et al. 2002; Nickum et al. 2004), identification tags, subcutaneous injections of elastomeric or other materials (Nickum et al. 2004), individual transponder tags (in animals of sufficient size), and, as applicable, external features such as individual color patterns. Because it can be difficult to individually identify some small aquatic animals throughout their life, group identification may be more appropriate in some situations (Koerber and Kalishman 2009; Matthews et al. 2002).

Aquatic Animal Recordkeeping Adequate recordkeeping is necessary in aquatic system management. In general, the same standards used for terrestrial animals apply to aquatic and semiaquatic species, although modifications may be necessary to account for species or system variations (Koerber and Kalishman 2009).

Although many aquatic animals are maintained using group (vs. individual) identification, detailed animal records are still necessary. Animal information that may routinely be captured, particularly in biomedical research with fish, includes species; genetic information (parental stock identification, genetic composition); stock source; stock numbers in system; tank identification; system life support information; breeding; deaths; illnesses; animal transfers within and out of the facility; and fertilization/hatching information (Koerber and Kalishman 2009; Matthews et al. 2002). Records should be kept concerning feeding information (e.g., food offered, acceptance), nonexpired food supplies to ensure sustenance of nutritional profile, and any live cultures (e.g., hatch rates and information to ensure suppliers’ recommendations are being met; Matthews et al. 2002).

Records of water quality testing for system and source water and maintenance activities of the life support system components are important for tracking and ensuring water quality. The exact water quality parameters tested and testing frequency should be clearly established and will vary with such factors as the type of life support system, animals, and research, as discussed under Water Quality. Detailed tracking of animal numbers in aquatic systems is often possible with accurate records of transfers, breeding, and mortalities (Matthews et al. 2002). In some cases where animals are housed in large groups (e.g., some Xenopus colonies) periodic censuses may be undertaken to obtain an exact count. In large-scale aquaculture research it may be more appropriate to measure biomass of the system versus actual numbers of animals (Borski and Hodson 2003).


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Gnotobiotic: germ-free animals or formerly germ-free animals in which the composition of any associated microbial flora, if present, is fully defined (Stedman’s Electronic Medical Dictionary 2006. Lippincott Williams & Wilkins).


Rabbits and some rodents, such as guinea pigs and hamsters, produce urine with high concentrations of proteins and minerals. These compounds often adhere to cage surfaces and necessitate treatment with acid solutions before and/or during washing.

Copyright © 2011, National Academy of Sciences.
Bookshelf ID: NBK54046


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