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Guide for the Care and use of Laboratory Animals: Eighth Edition 3 Environment, 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
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Guide for the Care and use of Laboratory Animals: Eighth Edition 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 ANIMALS 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).
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Guide for the Care and use of Laboratory Animals: Eighth Edition 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,
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Guide for the Care and use of Laboratory Animals: Eighth Edition 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. 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. Micro- TABLE 3.1 Recommended Dry-Bulb Macroenvironmental Temperatures for Common Laboratory Animals Animal Dry-Bulb Temperature °C °F Mouse, rat, hamster, gerbil, guinea piga 20-26 68-79 Rabbit 16-22 61-72 Cat, dog, nonhuman primate 18-29 64-84 Farm animals, poultry 16-27 61-81 aDry-bulb room temperature settings for rodents are typically set below the animals’ LCT to avoid heat stress, and should reflect different species-specific LCT values. Animals should be provided with adequate resources for thermoregulation (nesting material, shelter) to avoid cold stress.
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Guide for the Care and use of Laboratory Animals: Eighth Edition environmental 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
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Guide for the Care and use of Laboratory Animals: Eighth Edition 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 highvelocity 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 prefer-
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Guide for the Care and use of Laboratory Animals: Eighth Edition able 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 nonhuman 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. Illumination 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
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Guide for the Care and use of Laboratory Animals: Eighth Edition 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-Rowland 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
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Guide for the Care and use of Laboratory Animals: Eighth Edition 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
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Guide for the Care and use of Laboratory Animals: Eighth Edition 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
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Guide for the Care and use of Laboratory Animals: Eighth Edition 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 condi-
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Guide for the Care and use of Laboratory Animals: Eighth Edition Hankenson FC, Garzel LM, Fischer DD, Nolan B, Hankenson KD. 2008. Evaluation of tail biopsy collection in laboratory mice (Mus musculus): Vertebral ossification, DNA quantity, and acute behavioral responses. JAALAS 47(6):10-18. Hanifin JP, Brainard GC. 2007. Photoreception for circadian, neuroendocrine, and neurobehavioral regulation. J Physiol Anthropol 26:87-94. Hartl DL. 2000. A Primer of Population Genetics, 3rd ed. Sunderland, MA: Sinauer Associates. Hasenau JJ, Baggs RB, Kraus AL. 1993. Microenvironments in microisolation cages using BALB/c and CD-1 Mice. Contemp Top Lab Anim Sci 32:11-16. Hedrich HJ. 1990. Genetic Monitoring of Inbred Strains of Rats. New York: Gustav Fischer Verlag. Heffner HE, Heffner RS. 2007. Hearing ranges of laboratory animals. JAALAS 46:20-22. Held SDE, Turner RJ, Wootton RJ. 1995. Choices of laboratory rabbits for individual or group-housing. Appl Anim Behav Sci 46:81-91 Hermann LM, White WJ, Lang CM. 1982. Prolonged exposure to acid, chlorine, or tetracycline in drinking water: Effects on delayed-type hypersensitivity, hemagglutination titers, and reticuloendothelial clearance rates in mice. Lab Anim Sci 32:603-608. Hess SE, Rohr S, Dufour BD, Gaskill BN, Pajor EA, Garner JP. 2008. Home improvement: C57BL/6J mice given more naturalistic nesting materials build better nests. JAALAS 47:25-31. Hilken G, Dimigen J, Iglauer F. 1995. Growth of Xenopus laevis under different laboratory rearing conditions. Lab Anim 29:152-162. Hill D. 1999. Safe handling and disposal of laboratory animal waste. Occup Med 14:449-468. Hoffman HA, Smith KT, Crowell JS, Nomura T, Tomita T. 1980. Genetic quality control of laboratory animals with emphasis on genetic monitoring. In: Spiegel A, Erichsen S, Solleveld HA, eds. Animal Quality and Models in Biomedical Research. Stuttgart: Gustav Fischer Verlag. p 307-317. Homberger FR, Pataki Z, Thomann PE. 1993. Control of Pseudomonas aeruginosa infection in mice by chlorine treatment of drinking water. Lab Anim Sci 43:635-637. Hotchkiss CE, Paule MG. 2003. Effect of pair-housing on operant behavior task performance by rhesus monkeys. Contemp Top Lab Anim Sci 42:38-41. Hubrecht RC. 1993. A comparison of social and environmental enrichment methods for laboratory housed dogs. Appl Anim Behav Sci 37:345-361. Hughes HC, Reynolds S. 1995. The use of computational fluid dynamics for modeling air flow design in a kennel facility. Contemp Top Lab Anim Sci 34:49-53 Ikemoto S, Panksepp J. 1992. The effect of early social isolation on the motivation for social play in juvenile rats. Dev Psychobiol 25:261-274. Ivy AS, Brunson KL, Sandman C, Baram TZ. 2008. Dysfunctional nurturing behavior in rat dams with limited access to nesting material: A clinically relevant model for early-life stress. Neuroscience 154:1132-1142. Jacobs BB, Dieter DK. 1978. Spontaneous hepatomas in mice inbred from Ha:ICR Swiss stock: Effects of sex, cedar shavings in bedding, and immunization with fetal liver or hepatoma cells. J Natl Cancer Inst 61:1531-1534. Jones DM. 1977. The occurrence of dieldrin in sawdust used as bedding material. Lab Anim 11:137. Karolewicz B, Paul IA. 2001. Group housing of mice increases immobility and antidepressant sensitivity in the forced swim and tail suspension tests. Eur J Pharmacol 415:97-201. Kaufman BM, Pouliot AL, Tiefenbacher S, Novak MA. 2004. Short- and long-term effects of a substantial change in cage size on individually housed, adult male rhesus monkeys (Macaca mulatta). Appl Anim Behav Sci 88:319-330.
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