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Husbandry HOUSING Caging Caging is one of the primary components of a rodent's environment and can influence the well-being of the animals it houses. Many types of caging are available commercially. Those used to house rodents should have the following features: · They should accommodate the normal physiologic and behavioral needs of the animals, including maintenance of body temperature, normal movement and postural adjustments, urination and defecation, and, when indicated, reproduction. · They should facilitate the ability of the animal to remain clean and dry. · They should allow adequate ventilation. · They should allow the animals easy access to food and water and permit easy refilling and cleaning of the devices that contain food and water. · They should provide a secure environment that does not allow ani- mals to become entrapped between opposing surfaces or in ventilation openings. · They should be free of sharp edges or projections that could cause injury to the animals housed. · They should be constructed so that the animals can be seen easily without undue disturbance. 44
HUSBANDRY 45 · They should have smooth, nonporous surfaces that will withstand regular sanitizing with hot water, detergents, and disinfectants. · They should be constructed of materials that are not susceptible to corrosion. In selecting caging, one should pay close attention to the ease and thoroughness with which a cage can be serviced and sanitized. In addition to smooth, impervious surfaces that are free of sharp edges, cages should have minimal corners, ledges, and overlapping surfaces, because these fea- tures allow the accumulation of dirt, debris, and moisture. Cages should be constructed of durable materials that can withstand rough handling without ~ . . . . cupping or cracking. Sanitizing procedures, such as autoclaving and exposure to ionizing radiation, can alter the physical characteristics of caging materials over time and can greatly shorten useful life. Rodent cages are most commonly constructed of stainless steer 'or plastic (polyethylene, polypropylene, or polycarbonate), each of which has advantages and disadvantages. Galva ~ _ nized metal and aluminum have also been used but are generally less ac- ceptable because of their high potential for corrosion. Most rodent cages have at least one wire or metal grid surface to fur- nish ventilation and permit inspection of the animals in the'cage. Inspec- tion of animals can be further facilitated by the use of transparent plastic cages. Opaque plastic or metal cages might provide a more desirable envi- ronment for some studies or breeding programs; however, adequate inspec- tion of animals will usually require manipulation of each cage. The bottoms of rodent cages can be either solid or wire. The floors of solid-bottom cages usually are covered with bedding material that absorbs urine and moisture from feces, thereby improving the quality of the cage environment and allowing for easy removal of accumulated wastes. Solid- bottom cages provide excellent support for rodents' feet, minimizing the occurrence of pododermatitis and injuries. Wire-bottom cages are equipped with a wire-mesh grid, the spaces in which are large enough to allow the passage of feces. Generally, there are two to four wires per inch (2.5 cm) in the grid. These cages are normally mounted on racks that suspend them over waste-collection pans filled with absorbent material. This caging type minimizes contact with feces and urine and is thought to improve cage ventilation. However, careful consideration should be given to the size and species of rodents to be housed in wire-bottom cages because if their feet and legs can be entrapped in the wire ~rid, they can suffer severe trauma, including broken bones. In addition, older, heavier rodents can develop pododermatitis if the wires in the grid are too far apart or too small in diameter to provide adequate support for the feet. Specialized types of caging that serve specific functions are available for rodents, including caging designed to collect excrete, monitor physi _
46 RODENTS: LABORATORYANIMAL MANAGEMENT ologic characteristics, test behavioral responses, control aspects of the physical environment, and permit enhanced microbiologic control of the environ- ment. Such caging can pose special cleaning and sanitation problems. Various racking systems, both fixed and mobile, are available to hold either solid-bottom or wire-bottom cages. Racks should be constructed of durable, smooth-surfaced, nonporous materials that can be easily sanitized. Mobile racks are most commonly used because they allow greater flexibil- ity of room arrangement and are easier to clean than fixed racks. If fixed racks are used, adequate steps should be taken to protect floors or walls from damage caused by the weight of the racks and to provide for cleaning under and between the racks. Some racks incorporate devices that auto matically supply water directly to the cages they hold. Housing Systems Many types of housing systems with specialized caging and ventilation equipment are available for rodents. Generally, the purpose of these hous- ing systems is to minimize the spread of airborne microorganisms between cages; but they often do not prevent transmission of nonairborne fomites. The most frequently used of these systems is the filter-top cage, which has a spun-bound or woven synthetic filter that covers the wire-mesh top of a solid-bottom cage, thereby preventing the entry or escape of airborne par- ticles that can act as fomites for unwanted microorganisms. The use of filter tops restricts ventilation and can alter the microenvironment of the rodents housed in the cages; therefore, to maintain a healthful environment, it might be necessary to change the bedding and clean the cages more often (Keller et al., 19891. A cubicle (also called an Illinois cubicle or a cubical containment sys- tem) is an enclosed area of a room capable of housing one or more racks of cages. It is separated from the rest of the room by a door that usually opens and closes vertically. The cubicle is supplied by air that moves under the door from the room and is exhausted through the ceiling, or a separate air supply is provided to the cubicle through an opening in a wall, the base, or the ceiling. Cubicles have been used to reduce airborne cross contamina- tion between groups of animals housed in conventional plastic or wire- bottom cages (White et al., 19831. They provide better ventilation than many housing methods, but they do not protect against fomite transmission of microorganisms. Strict adherence to sanitation and other husbandry pro- cedures is required if cubicles are to be used effectively. In some housing systems, cages are individually ventilated with highly filtered air. In some, exhaust air is also filtered or controlled in a way that greatly minimizes the risk of contaminating animals in other cages and personnel in the animal rooms. Such systems can overcome the disadvan
HUSBANDRY 47 sages of using nonventilated filter-topped cages while minimizing airborne cross-contamination. A housing system that is particularly useful for maintaining the micro- biologic status of rodents has isolators made of rigid or flexible-film plastic that are designed to enclose a group of rodent cages. Built-in gloves allow the manipulation of animals and materials in the isolators. Isolators are supplied with filtered air and have a filtered exhaust; at least one transfer device is provided for moving sterilized or disinfected materials into the isolator. To maintain the microbiologic status of an isolated group of ani- mals, it is necessary to sterilize or otherwise disinfect all the interior sur- faces of the isolators, and all materials introduced into the isolators should be first sterilized or otherwise disinfected. Space Recommendations ~. ~ It is generally assumed that there are critical measures of cage floor area and cage height below which the physiology and behavior of labora- tory rodents will be adversely affected, thereby affecting the well-being of the animals and potentially influencing research outcomes. However, there are very few objective data for determining what those critical measures are or even whether such interactions exist. A number of studies designed to evaluate the effects of space on population dynamics have been conducted on wild and laboratory rodents housed in a laboratory environment (e.g., see Barnett, 1955; Christian and LeMunyan, 1958), but some of them used caging systems different from those generally used in laboratory animal facilities (e.g., see Davis, 1958; Joasoo and McKenzie, 1976; Thiessen, 1964~. Changes in behavior, reproductive performance, adrenal weights, A. ~ - glucocorticoid and catecholamine concentrations, immunologic function, numbers of some kinds of white blood cells (usually lymphocytes), and cage-use patterns have been assessed in those studies and suggested as indicators of stress and compromised well-being (e.g., see Barrett and Stockham, 1963; Bell et al., 1971; Christian, 1960; Poole and Morgan, 1976; White et al., 1989~. However, there has never been general agreement as to which physiologic and behavioral characteristics are indicative of well-being in rodents or what magnitude of change in them would be necessary to com- promise the well-being of the animals. With few objective data available, cage space recommendations have been based on the results of surveys of existing conditions and professional judgment and consensus. The Guide (NRC, 1996 et seq.) provides space recommendations for rodents. Space recommendations have also been de- veloped in other countries (CCAC, 1980; Council of Europe, 1990), but they are not totally compatible with those in the Guide. It is important to remember that space recommendations in the Guide serve only as a starting
48 RODENTS: LABORATORY ANIMAL MANAGEMENT point for determining space required by rodents and might need adjustment to fit the needs of the animals and the purposes for which they are housed. Although comprehensive studies involving all the characteristics asso- ciated with housing rodents are not available, sufficient information does exist to suggest that individually housed rodents and group-housed rodents have different space requirements. For the most part, laboratory rodents are social animals and probably benefit from living in compatible groups (Brain and Bention, 1979; NRC, 1978; White, 19901. Although more study is needed, rodents maintained for long periods, as in lifetime studies, appear to survive longer when housed in large, compatible social groups than when housed in small groups or individually (Hughes and Nowak, 1973; Rao, 19901. Individual housing is sometimes necessitated by the nature of the experimental protocol; in such instances, adequate space should be allotted to allow the animals to make normal postural adjustments, which will de- pend on the body size attained by the animals during the course of the experiment. Under those circumstances, current space guidelines might not be sufficient, especially if an animal's size exceeds the scope of the recom- mendations. Conversely, group-housed rodents would be expected to need less space per animal than individually housed rodents because each animal can also use the space of the other rodents with which it is housed. Studies have found that compatible social groups of rodents do not use all the available space recommended in current guidelines and probably do not require it for well-being (White, 1990; White et al., 1989~. Rodents housed in compat- ible groups share cage space by huddling together along walls and under overhanging portions of the cage, such as feeders, as well as piling up on top of each other during long rest periods. The center of the cage is used infrequently. Even if individually housed, rodents appear to prefer sheltered areas of the cage, especially if those areas have decreased light and height. Provid- ing such a confined space within a cage might be one way to enrich the environment of rodents. Sexually mature male rodents often fight when housed in groups for breeding or other purposes, but this behavior has never been shown to be a function of the amount of available floor space in the cage. Rather, the incidence of fighting appears to be related more to combining males into groups when they are sexually mature (especially if females are housed in the same room) or have participated in mating programs. Increasing the cage space is not effec- tive in preventing the development of such behavior or in eliminating it once it has occurred. Only separation of the animals into individual cages or into smaller, compatible groups is effective in eliminating fighting. In determining adequate cage space, it is important to consider the conditions of the experimental procedure and how long the animals will be
HUSBANDRY 49 housed. Animals that become debilitated during the course of an experi- mental procedure might require increased cage space or an alteration in caging to accommodate limitations in motion, recumbent positions, and the need for alternative food and water sources. Older animals are less active than younger animals and use less of the cage space or available activity devices. The Guide (NRC, 1996 et seq.) and other guidelines also recommend cage heights. The recommendations do not appear to be related to the body size of rodents nor to their normal locomotion patterns. Laboratory rodents exhibit some vertical exploratory behavior when put into a new cage, and it has been suggested that relatively high cages be provided to accommodate this occasional behavior (Lawlor, 1990; Scharmann, 1991~. However, there is no good evidence to suggest that rodents require tall enclosures. On the contrary, as described previously, they tend to seek shelter under objects lower than recommended in existing guidelines. Depending on the caging type, existing height guidelines can be useful for ensuring that there is adequate space for side-wall or cage-top feeders and adequate clearance for sipper tubes or other watering devices. In summary, the space required to maintain rodents, either individually or in groups, depends on a number of factors, including age, weight, body size, sexual maturity, experimental intervention, behavioral characteristics, the duration of housing, group size, breeding activities, and availability of enrichment devices or sheltering areas within the cage. The relationships among those factors are complex, and there is not necessarily a direct corre- lation between body weight or surface area of the animals and the absolute floor area of the cage required or used by them. Guidelines should be used with professional judgment based on assessment of the animals' well-being. However, alterations that bring floor area or height of cages below recom- mended levels should be adequately justified and approved by the IACUC. ENVIRONMENT Microenvironment The microenvironment of a rodent is the physical environment that immediately surrounds it and is usually considered to be bounded by the primary enclosure or cage in which it resides. In contrast, the physical conditions in the secondary enclosure or animal room make up the macroenvironment. The components of the macroenvironment are often easier to measure and characterize than those of the microenvironment. The two environments are linked or coupled, but the character of each is often quite different and depends on a variety of factors, such as the numbers and species of rodents housed in the microenvironment, the design and con
50 RODENTS: LABORATORYANIMAL MANAGEMENT struction of the cages, and the types of bedding materials used (Beech, 1975; Woods, 1975; Woods et al., 1975~. The measurement of constituents of the microenvironment of rodents is often difficult because of the relatively small volume of the primary enclo- sure. Available data show that temperature, humidity, and concentrations of gases and particulate matter such as carbon dioxide, ammonia, meth- ane, sulfur dioxide, respirable particles, and bacteria are often higher in the microenvironment than in the macroenvironment (Beech, 1980; Clough, 1976; Flynn, 1968; Gamble and Clough, 1976; Murakami, 1971; Serrano, 19711. Although there is little information on the relation between the magnitude of exposure to some of those components and alterations in dis- ease susceptibility or changes in metabolic or physiologic processes, the available data clearly suggest that the characteristics of the microenviron- ment can have a substantial impact on research results (Broderson et al., 1976; Vessell et al., 1973, 1976~. Temperature Temperature and relative humidity are important components of the environment of all animals because they directly affect an animal's ability to regulate internal heat. They act synergistically to affect heat loss in rodents, which lose heat by insensible means, rather than by perspiring. Studies in the older literature, which were conducted without the benefit of modern systems for controlling conditions precisely or modern instrumen- tation, have established that extremes in temperature can cause harmful effects (Lee, 1942; Mills, 1945; Mills and Schmidt, 1942; Ogle, 1934; Sunstroem, 1927~. However, those studies were done on only a few labo- ratory species. Studies in the past generally focused on prolonged exposure of labora- tory animals to temperatures above 85°F (29.4°C) or below 40°F (4.4°C), which are required to achieve clinical effects (Baetjer, 1968; Mills, 1945; Weihe, 1965~. When exposed to those extreme temperatures, rodents use behavioral means (e.g., nest-building, curling up, huddling with others in the cage, and adjusting activity level) to attempt to adapt. If the tempera- ture change is brief or small, behavioral adaptation is sufficient; profound or prolonged temperature changes generally require physiologic or struc- tural adaptation as well. Physiologic adaptation includes alterations in metabolic rate, growth rate, and food or water consumption; hibernation or estivation; and the initiation of nonshivering thermogenesis. Structural ad- aptation includes alterations in fat stores, density of the hair coat, and struc- ture or perfusion of heat-radiating tissues and organs (e.g., tail, ears, scro- tum, and soles of the feet). Initiation of such changes usually requires exposure to an extreme temperature for at least 14 days.
HUSBANDRY 51 For routine housing of laboratory rodents, a consistent temperature range should be provided. However, there is little scientific evidence from which optimal temperature ranges for laboratory rodents can be determined. For each species, there is a narrow range of environmental temperatures at which oxygen consumption is minimal and virtually independent of change in ambient temperature. The range in which little energy is expended to main- tain body temperature is called the thermal neutral zone, and some have suggested that it is a range of comfortable temperatures for rodents (Beech, 1985; Weihe, 1965, 1976a). However, other evidence suggests that animals held within this temperature range do not necessarily achieve optimal growth and reproductive performance, and the optimal temperature range might be age-dependent (Blackmore, 1970; Weihe, 1965J. Moreover, measurements of thermal neutral zones are generally made on resting animals and do not take into account periods of increased activity or altered metabolic states, such as pregnancy. Thermal neutrality does not necessarily equate with comfort. In the absence of well-controlled studies that used objective mea- sures for determining optimal ranges, recommended temperature ranges for laboratory rodents have been independently developed by several groups on the basis of professional judgment and experience (e.g., CCAC, 1980; Council of Europe, 1990; NRC, 1996 et seq.~. Humidity Relative humidity varies considerably with husbandry and caging prac- tices. In addition, there is usually a difference between the relative humid- ity in the room and that in the animal cages. Many factors including cage material and construction, use of filter tops, number of animals per cage, frequency of bedding changes, and bedding type can affect the relative humidity in the rodents' immediate environment. Variations in relative humidity appear to be tolerated much better at some temperatures than at others. Studies in humans and limited in vitro work on survival of microorganisms have established a loose association between humidity and susceptibility to disease (Baetjer, 1968; Dunklin and Puck, 1948; Green, 1974; Webb et al., 1963), but there is no good evidence to establish this link in animals. Low relative humidity has been reported to be associated with the development of "ring tail" in rodents (Flynn, 1959; Njaa et al., 1957; Stuhlman and Wagner, 1971~; however, this condition has not been adequately studied and does not appear to be reproducible by lowering relative humidity in controlled laboratory experiments. Guidelines have been established for relative-humidity ranges based on experience and professional judgment (CCAC, 1980; Council of Europe, 1990; NRC, 1996 et seq.~. There is no evidence to support limiting the variation of relative humidity within these ranges; however, the combina
52 RODENTS: LABORATORYANIMAL MANAGEMENT lion of high relative humidity and high environmental temperature can af- fect the ability of rodents to dissipate heat by insensible means and should be avoided. Ventilation Ventilation Rate Ventilation refers to the process of using conditioned air to affect tem- perature, humidity, and concentrations of gaseous and particulate contami- nants in the environment. Ventilation is often characterized at the animal- room level as air exchanges per hour. However, as for other environmental conditions, there are no definitive data showing that the air-exchange range in existing guidelines (i.e., 10-15 air changes/hour) provides optimal venti- lation for laboratory rodents. Existing guidelines have been criticized as being based mainly on keep- ing odors below objectionable limits for humans (Beech, 1980; Runkle, 1964) and, in recent years, as being energy-intensive. An often-quoted study by Munkelt (1938) appears to support the first contention: his mea- sure of adequate ventilation was the ability to smell ammonia in the envi- ronment. Besch (1980) suggested that ventilation should be based on air- exchange rate per animal or animal cage because room air-exchange rates do not consider such factors as population density, room configuration, and cage placement within a room. Ultimately, however, the ventilation rate in animal rooms is governed by the heat loads produced in the rooms, which include not only heat produced by animals but also that produced by other heat-radiating devices, such as lighting (Curd, 1976~. Available evidence suggests that little additional control of the concen- trations of gaseous and particulate contaminants is gained by using air- exchange rates higher than those recommended in current guidelines (Barkley, 1978; Besch, 19801. The recommendation of providing a room air-ex- change rate of 10-15 changes/hour is still useful; however, this ventilation range might not be appropriate in some circumstances, especially if the diffusion of air within the room is inappropriate for the type and placement of cages. Other methods of providing equal or more effective ventilation, including the use of individually ventilated cages or enclosures and the adjustment of ventilation rate to accommodate unusual population densities, are also acceptable. Calculation of the amount of cooling required to control expected sen- sible and latent heat loads generated by the species to be housed and the largest expected population (ASHRAE, 1993) can be used to determine minimal ventilation requirements. However, that calculation does not take into account the generation of odors, particles, and gases, which might necessitate greater ventilation.
HUSBANDRY Air Quality 53 The quality of air used to ventilate animal rooms is another important consideration. Ventilation systems for rodent rooms incorporate various types of filtration of incoming air. Coarse filtration of the air supply is a minimal requirement for proper operation of ventilating equipment. Most facilities maintaining rodents of defined microbiologic status also use high- efficiency particulate air (commonly called HEPA) filters to decrease the risk of introducing rodent pathogens into the animal room through the fresh- air supply (Dyment, 1976; Harstad et al., 1967~. The required filter effi- ciency is a matter of professional judgment, and selection should be based on the perceived likelihood of introducing contaminated air into the room. Filtration of exhaust air from rodent rooms when air is not recycled is usually deemed unnecessary unless the exhaust air is likely to contain haz- ardous or infectious materials. Filters designed to remove chemicals from air are sometimes incorporated into exhaust systems to remove animal odors. Activated-chemical filters (e.g., those with activated charcoal) are often used for this purpose; however, their efficiency in removing odoriferous compounds, including ammonia, varies, and they require substantial mainte- nance to remain effective. The use of recycled air to ventilate animal rooms can save considerable amounts of energy. However, many animal pathogens can be airborne or travel on fomites, such as dust, so recycling of exhaust air into heating, ventilating, and air-conditioning systems that serve multiple rooms presents a risk of cross contamination. Exhaust air that is to be recycled should be HEPA-filtered to remove particles. HEPA filters are available in various efficiencies; the extent and efficiency of filtration should be proportional to ~ ~ 1 ~ ~ ~ ~ 1 the risk. toxic or odor-caus~ng gases produced oy aecompos~uon of animal wastes can be removed by the ventilating system with chemical absorption or scrubbing, but those methods might not be completely effective. Fre- quent bedding changes and cage-cleaning, a reduction in number of animals housed in a room, and a decrease in environmental temperature and humid- itv within limits recommended in the Guide (NRC, 1996 et seq.) can also assist in reducing the concentration of toxic or odor-caus~ng gases. Treatment of recycled air to remove either particulate or gaseous contami- nants is expensive and can be ineffective if filtration systems are improp- erly or insufficiently maintained. Therefore, recycling systems require regular monitoring for effective use. An energy-recovery system that reclaims heat and thereby makes it en- ergy-efficient to exhaust animal-room air totally to the outside is also accept- able, but these systems often require much maintenance to be effective. The recycling of air from nonanimal areas can be considered as an alternative to the recycling of animal-room air, but this air might require filtering and treat- ment to remove odors, toxic chemicals, and particles (White, 1982~.
54 Relative Air Pressures R ODENTS: LAB ORA TOR Y ANIMAL MANA CEMENT To minimize the potential for airborne cross-contamination between adjacent rodent rooms or between rodent rooms and other areas where con- taminants might be generated, it is important to consider controlling relative air pressures. By adjusting the rates of air flow to and from individual areas, one can produce a negative or positive pressure relative to adjoining areas. When the intent is to contain contaminants (e.g., in areas used to quarantine newly arrived animals, isolate animals infected or suspected of being infected with rodent pathogens, house animals or materials inoculated with biohazardous substances, or keep soiled equipment), air pressure in the containment area should be lower than that in surrounding areas. When the intent is to prevent the entry of contaminants, as in areas used to house specific-pathogen-free rodents or keep clean equipment, air pressure in the controlled area should be greater than that in surrounding areas. It is im- portant to remember, however, that many factors influence disease trans- mission between adjacent rooms; simply controlling air pressure is not suf- ficient to prevent transmission. Cage Ventilation Ventilation can easily be measured in rodent-holding rooms; however, conditions monitored in a room do not necessarily reflect conditions in the cages in the room. The large sample volumes required by the commonly used instruments that measure ventilation, as well as the size of the intruments themselves, preclude accurate measurement in cages (Johnstone and Scholes, 1976~. The degree to which cages are ventilated by the room air supply is affected by cage design; room air-diffuser type and location; number, size, and type of animals in the cages; presence of filter tops; and location of the cages. For example, cages without filter tops provide better air and heat exchange than those with filter tops, in which ventilation is substantially decreased (Keller et al., 19891. Rigidly maintaining room air quality and ventilation will not necessarily provide the same environment for similar groups of animals or even for similar cages in the same room. Individually ventilated cages provide better ventilation for the animals and, possibly, a more consistent environment (Lipman et al., 1992), but those systems are generally expensive. It has not been established whether rodents are uncomfortable when exposed to air movements (drafts) or that exposure to drafts has biologic consequences. However, movement of air in a room influences the ventila- tion of an animal's primary enclosure and so is an important determinant of microenvironment.
HUSBANDRY 55 Illumination Animal-room lighting can affect the eyes of laboratory rodents, espe- cially albino rodents. In examining the effects, there is a tendency to think only in terms of light intensity. However, it is the interaction of the three characteristics of light (spectral distribution, photoperiod, and intensity) that produces the effects (Brainard, 1988; Wurtman et al., 1985. Also contribut- ing to the effects of light exposure is the amount of time that rodents have their eyes open during the hours when the room is lit. Those factors should be kept in mind in reading the following discussion. Spectral Distribution Artificial lighting with white incandescent or fluorescent fixtures is preferred for rodent housing facilities because it provides consistent illumi- nation. The two types of lighting have similar spectra, although incandes- cent lighting generally has more energy in the red wavelengths and less energy in the blue and ultraviolet (UV) wavelengths than white fluorescent lighting. Although some fluorescent lighting more closely simulates the wavelength distribution of sunlight than incandescent lighting, no artificial lighting truly duplicates sunlight, and there is little reason to believe that the spectral distribution of one type of artificial lighting is superior to that of any other for rodent rooms. There is some evidence that UV light can increase the incidence of cataract formation in humans (Zigman et al., 1979) and in rodents exposed to very high levels (Zigman and Vaughan, 1974; Zigman et al., 19731. However, there is no evidence that UV-associated cataracts develop in rodents maintained under levels of illumination nor- mally found in animal rooms. UV radiation from fluorescent lights is elimi- nated when the lights are covered by plastic diffusing screens (Kaufman, 1987; Thorington, 1985~. Photoperiod Photoperiod (cycles of light and dark during the course of a single day) affects various physiologic and metabolic characteristics, including repro- ductive cycles, behavioral activity, and the release of hormones into the blood (Brainard, 1989; Reiter, 1991~. The rods and cones in the eye are influenced by photoperiod, requiring an interval of darkness for regenera- tion (LaVail, 1976; Williams, 1989; Williams and Baker, 1989~. There is evidence that exposure to even low-intensity light 64.6-193.7 lx (6-18 ft- candles) continuously for 4 days can cause degenerative retinal changes (Anderson et al., 1972; O'Steen, 1970; Williams, 1989~;
56 RODENTS: LABORATORYANIMAL MANAGEMENT Photoperiods in rodent rooms are usually controlled by automatic tim- ers. The cycles usually recommended are either 12 hours of light and 12 hours of dark or 14 hours of light and 10 hours of dark. For some mammals (e.g., hamsters), a longer period of light is important for normal reproduc- tion (Alleva et al., 1968~. In general, lighting in laboratory animal facilities does not reproduce that in nature, in that most light-timing devices do not provide any interval of reduced lighting intensity (simulating dawn and dusk). Changes or interruptions in light-dark cycles should be avoided because of the importance of photoperiod in normal rodent reproduction and other light-affected physiologic processes (Weihe, 1976b). Similarly, light from exterior windows and uncontrolled hallway lighting are usually undesirable. Light-timing devices in rodent facilities should be checked regularly for correct operation. Any system that can be overridden manually should be equipped with an indicator, such as a light, to remind personnel to turn off the override device or with a timer to turn it off automatically. Photoperiod can also be checked by photosensors linked to a computer-based monitor. Intensity The intensity of illumination is inversely proportional to the square of the distance from the source. Therefore, statements about intensity should indicate where it was measured. In animal facilities, such statements gener- ally specify distance above the floor; that implies that the illumination is uniformly diffused throughout the room. The actual intensity experienced by a rodent in an animal room is influenced not only by the relative loca- tions of its cage and the room lights, but also by cage material and design. The optimal light intensity required to maintain normal physiology and good health of laboratory rodents is not known. In the past, illumination of 807-1076 lx (75-100 ft-candles) or higher has been recommended to allow adequate observation of the animals and safe husbandry practices (NRC, 1978~. The point of measurement for that recommendation was never clearly stated, but it has been generally assumed that the recommendation referred to the illumination at maximal cage height in the center of the room. The recommended intensities, however, have been shown to cause retinal dam- age in albino mice (Greenman et al., 1982) and rats (Lad et al., 1978; Stotzer et al., 1970; Williams and Baker, 1980~. More recently, a light intensity of 323 lx (30 ft-candles) measured about 1.0 m (3.3 It) above the floor has been recommended as adequate for rou- tine animal care (Bellhorn, 1980; NRC, 1996 et seq.~. That intensity has been calculated to provide 32-40 lx (3.0-3.7 ft-candles) to rodents in the front of a cage that is in the upper portion of a cage rack. Exposure for up to 90 days to an intensity of around 300 lx during the light cycle has been
HUSBANDRY 57 reported not to cause retinal lesions in rats (Stotzer et al., 1970~; however, it is still questionable whether exposure to light of even this intensity can cause retinal lesions in albino animals if they are exposed for longer periods (Weisse et al., 1974~. Alternatives to providing a single light intensity in a room are to use variable-intensity controls and to divide rooms into zones, each lighted by a separate switching mechanism. Those alternatives conserve energy and provide sufficient illumination for personnel to perform their tasks adequately and safely. However, caution is necessary when instituting those alterna- tives. Boosting daytime room illumination for maintenance purposes has been shown to change photoreceptor physiology and can alter circadian regulation (Reme et al., 1991; Society for Research on Biological Rhythms, 1993; Terman et al., 19911. Noise Many sounds of varied frequencies and intensities are generated in ani- mal facilities during normal operation. Rodents emit ultrasonic vocaliza- tions that are an important part of their social and sexual behavior. Rats can hear frequencies as high as about 60-80 kHz but are relatively insensi- tive to frequencies less than 500 Hz (Kelly and Masterton, 1977; Peterson, 19801. Sounds are also produced by mechanical equipment (less than 500 Hz): by dog, cat, nonhuman primate, and pig vocalizations (up to 120 dB at 500 Hz); and by exterior conditions (e.g., highway noise). If acoustic energy is high enough (80-100 dB), both auditory and nonauditory changes can be detected in laboratory animals (Algers et al., 1978; Molter, 19781. The type of change produced depends on the pattern of sound presentation. Sound of uniform frequency and unchanging inten- sity can cause hearing loss in some rodents (Bock and Saunders, 1977; Burdick et al., 1978; Kelly and Masterton, 1977; Kraak and Hofmann, 1977~. Hamsters, guinea pigs, rats, and mice pass through developmental stages during which they are very susceptible to injury from sound of this type (Kelly and Masterton, 19771. Sound of irregular frequency and rapidly changing intensity that is presented to animals in an unpredictable fashion can cause stress-induced mechanical and metabolic changes (Anthony and Harclerode, 1959; Geber, 1973; Guha et al., 1976; Kimmel et al., 1976; Peterson et al., 1981~. Continuous exposure to acoustic energy above 85 dB can cause eosinopenia (Geber et al., 1966; Nayfield and Besch, 1981), in- creased adrenal weights (Geber et al., 1966; Nayfield and Besch, 1981), and reduced fertility (Zondek and Tamari, 1964~. Few studies are available on the long-term effects on rodents of sound comparable with that normally encountered in rodent rooms, and there are hardly any data on the sensitivity of rodents to intensity as a function of
58 RODENTS: LABORATORY ANIMAL MANAGEMENT frequency (Peterson, 1980~. In addition, no comparative damage-risk crite- ria have been established for rodents; therefore, recommendations for ac- ceptable noise in animal facilities are often based on extrapolations from humans (Peterson, 19801. As a general guideline, an effort should be made to separate rodent-housing areas from human-use areas, especially human- use areas where mechanical equipment is used or where noisy operations are conducted. Common soundproofing materials are not compatible with some of the construction requirements for animal facilities designed to house rodents, but attention can be given to separating rooms housing rodents from those housing noisy species, such as nonhuman primates, dogs, cats, and swine. The location of loud, unpredictable sources of noise such as intercoms, paging systems, telephones, radios, and alarms-should be care- fully considered because the noise from such sources can be stressful to some rodents. Extra care should be taken to control noise in rooms that house rodents that are subject to audiogenic seizures. Every reasonable effort should be made to house rodents in areas away from environmental sources of noise. FOOD Nutrition has a major influence on the growth, reproduction, health, and longevity of laboratory rodents, including their ability to resist pathogens and other environmental stresses and their susceptibility to neoplastic and nonneoplastic lesions. Providing nutritionally adequate diets is important not only for the rodents' welfare, but also to ensure that experimental re- sults are not biased by unintended or unknown nutritional factors. Provid- ing nutritionally adequate diets for laboratory rodents involves establishing requirements for about 50 essential dietary nutrients, formulating and manu- facturing diets with the required nutrient concentrations, and managing nu- merous factors related to diet quality. Factors that potentially affect diet quality include bioavailability of nutrients, palatability or acceptance by-the animals, preparation and storage procedures, and concentrations of chemi- cal contaminants. The estimated nutrient requirements of laboratory animal species are periodically reviewed and updated by a committee of the Na- tional Research Council (NRC, 1995), and information about the formula- tion, manufacture, and management of laboratory animal diets is available elsewhere (Coates, 1987; Knapka, 1983, 1985; McEllhiney, 1985; Navia, 1977; Rao and Knapka, 1987~. Types of Diets Adequate nutrition can be provided for laboratory rodents in different types of diets that are classified by the degree of refinement of the ingredi- ents used in their formulation (NRC, 1995~.
HUSBANDRY 59 Natural-ingredient diets are formulated with agricultural products and byproducts, such as whole grains (e.g., ground corn and ground wheat), mill byproducts (e.~., wheat bran, wheat middlings, and corn gluten meal), high - , . _ . ~ ~ 1 ~ 1 _ __ 11 1:~_ ~ 1~ ~ ~ ~ 1 ~ __~ ~ ~ ~ ~ ~ 1 An; ~ ~ 1 ~ ma.. ~ ~ ~ protein meals (e.g., soybean meal and Ilsn meal), processed IIllIlt;~-~1 5OU1~b (e.g., bone meal), and other livestock feed ingredients (e.g., dried molasses and alfalfa meal). Commercial diets are the most commonly used natural- ingredient diets, but special diets for specific research programs can also be of this type if appropriate attention is given to ingredient selection and diet formulation. Natural-ingredient diets are relatively inexpensive to manu- facture and are readily consumed by laboratory rodents. A natural-ingredient diet can be either an open-formula diet (informa- tion on the amount of each ingredient in the diet is readily available) or a closed-formula diet (information on the amount of each ingredient is privi- leged). The advantages of using natural-ingredient, open-formula diets in biomedical research have been discussed (Knapka et al., 1974~. There are two concerns about the use of natural-ingredient diets in biomedical research. First, such factors as varieties of plants, soil composi- tions, weather conditions, harvesting and storage procedures, and manufac- turing and milling methods influence the nutrient composition of ingredi- ents used in this type of diet to the extent that no two production batches of the same diet are identical (Knapka, 19831. This variation in dietary-nutri- ent concentrations introduces an uncontrolled variable that could affect ex- perimental results. Second, natural ingredients can be exposed to various naturally occurring or human-made contaminants, such as pesticide resi- dues, heavy metals, and estrogen. Diets manufactured from natural ingredi- ents can contain low concentrations of contaminants that might have no influence on animal health but could affect experimental results. For ex- ample, a lead concentration of 0.5-1 part per million is inherent in natural- ingredient rodent diets and is not generally detrimental to animal health; but it could substantially influence the results of toxicology studies designed to evaluate lead-containing test compounds. Purified diets are formulated with ingredients that have been refined so that in effect each ingredient contains a single nutrient or nutrient class. Examples of the ingredients are casein or soy protein isolate, which provide protein and amino acids; sugar and starch, which provide carbohydrates; vegetable oil and lard, which provide essential fatty acids and fat; a chemi- cally extracted form of cellulose, which provides fiber; and chemically pure inorganic salts and vitamins. The nutrient concentrations in this type of diet are less variable and more readily controlled than those in natural- ingredient diets. However, purified ingredients can contain low and vari- able concentrations of trace minerals, and batch-to-batch variation in their concentrations is inherent in the manufacture of purified diets. The poten- tial for chemical contamination of purified diets is low; however, they are
60 RODENTS: LABORATORY ANIMAL MANAGEMENT not always readily consumed by laboratory rodents, and they are more ex- pensive to produce than natural-ingredient diets. Chemically defined diets are formulated with the most elemental ingre- dients available, such as individual amino acids, specific sugars, chemically defined triglycerides, essential fatty acids, inorganic salts, and pure vita- mins. Use of this type of diet provides the highest degree of control over dietary nutrient concentrations. However, chemically defined diets are not readily consumed by laboratory rodents, and they are usually too expensive for general use. The dietary nutrient concentrations in chemically defined diets are theo- retically fixed at the time of manufacture; however, the bioavailability of nutrients can be altered by oxidation or nutrient interactions during diet storage. Liquid chemically defined diets that can be sterilized by filtration have been developed (Pleasants, 1984; Pleasants et al., 1986~. Criteria for Selecting Optimal Rations Selection of the most appropriate type of diet for a particular animal colony depends on the reproductive or experimental objectives. One of the most important considerations is the amount of control over dietary-nutrient composition that is necessary to attain the objectives. For example, the use of a purified diet is essential for studies designed to establish quantitative requirements for micronutrients because the batch-to-batch variation in nu- trient concentrations inherent in natural-ingredient diets would compromise experimental results. Conversely, the variation in nutrient concentrations in natural-ingredient diets would have no detectable influence on rodent pro- duction colonies because the nutrient concentrations are generally greater than those required in a nutritionally adequate diet. The use of chemically defined diets might be required for studies whose objectives involve dietary concentrations of single amino or fatty acids. The potential for chemical contamination is an important consideration in selecting a diet for rodents that will be used in toxicology studies. Even though the concentrations of chemical contaminants in natural-ingredient diets are so low that they generally do not jeopardize animal health, they might be high enough to compromise results of toxicology studies. The results of some immunology studies might also be influenced by the use of natural-ingredient diets because some ingredients, particularly those of ani- mal origin, contain antigens. Purified diets should be considered for ani- mals used in both kinds of studies, although their cost can increase the cost of conducting the research, especially in life-span studies that use large numbers of rodents. Any diet selected should be accepted by the animals, otherwise consid- erable amounts will be wasted. This is expensive and constitutes a major
HUSBANDRY 61 disadvantage in studies that require quantification of dietary consumption. Diets should be nutritionally balanced and free of toxic or infectious agents. If diet is a factor in a study, the diet selected should be readily reproducible to ensure that research results can be verified by replication. Quality Assurance Although reputable laboratory animal feed manufacturers develop elaborate programs to ensure the production of high-quality products, additional pro- cedures are often required to ensure that the diets are nutritionally adequate. The shelf life of any particular feed lot depends on the environmental con- ditions during storage. Nutrient stability of animal feeds generally increases as temperature and humidity in the storage environment decrease. Natural- ingredient rodent diets stored in air-conditioned areas in which the tempera- ture is maintained below 21°C (70°F) and the humidity below 60 percent should be used within 180 days of manufacture. Vitamin C in diets stored under these conditions has a shelflike of only 90 days. If a vitamin C- containing diet stored for more than 90 days is to be fed to guinea pigs, an appropriate vitamin supplement should be added. To monitor compliance with these guidelines, storage containers should be marked with the date of manufacture of the food stored therein. Diets stored for longer periods or under conditions other than those recommended above should be assayed for the most labile nutrients (i.e., vitamin A, thiamine, and vitamin C) before use. Diets formulated without antioxidants or with large amounts of highly perishable ingredients, such as fat, might require special handling or storage procedures. Given the potential importance of diet quality and consistency to experi- mental results, a routine program of nutrient testing should be implemented to verify the composition of diets fed to research animals. Accidental omission or inclusion of ingredients in the manufacturing process, although uncommon, can have disastrous consequences on research projects. Discrepancies be- tween expected and actual nutrient concentrations in laboratory animal diets can arise from errors in formulation, which can result in hazardous concentra- tions of nutrients that are toxic when present in excess of requirements (e.g., vitamins A and D, copper, and selenium); losses of labile nutrients during manufacture or storage; variation in nutrient content of ingredients used in diet formulation; and errors associated with diet sampling or analysis. Although most laboratory animal feed manufacturers will provide data on the complete nutrient composition of rodent diets, it is often difficult to ascertain the source of these data (i.e., whether they are calculated, representative of several diet production batches, or representative of a single production batch). Therefore, it is suggested that feed manufacturers routinely be asked to provide the results of nutrient assays of representative samples of their diets.
62 RODENTS: LABORATORY ANIMAL MANAGEMENT Testing samples of natural-ingredient diets used in research colonies is particularly important because the nutrient concentrations measured by analy- sis can differ from the expected concentrations. Samples for assay should be collected from multiple bags or containers within a single production batch of feed (i.e., in which all containers bear the same manufacture date). The con- tainers sampled should be selected at random; traditionally, the number sampled equals the square root of the total number of containers in a single shipment or production batch. The objective is to obtain a sample of diet that is represen- tative of the entire lot being assayed. Nutrient analyses should be conducted by a laboratory with an established reputation in assaying feed samples, and all assays should be conducted in accordance with the most recent methods published by the Association of Official Analytical Chemists (Helrich, 1990~. Analyses should include at least proximate constituents (i.e., moisture, crude protein, ether extract, ash, and crude fiber) and any nutrients that are under study or that could influence the study. Some vitamins and other nutrients required at trace concentrations might be difficult to assay because of low concentrations, interfering compounds, or both. The presence of biologic contaminants in diets is a cause for concern in most research and production rodent colonies. Unwanted agents in the diet include pathogenic bacteria and viruses, insects, and mites. Diets for axenic and microbiologically associated rodents should be sterilized before use, as should those for severely immunodeficient rodents (i.e., athymic rodents and mice homozygous for the mutation scid) (NRC, 1989J. Diets for spe- cific-pathogen-free (SPF) rodents should be subjected to some degree of decontamination, such as pasteurization. It is also prudent to decontaminate diets, at least partially, for conventionally maintained rodents, particularly when they are used in long-term studies. Steam autoclaving is the most widely used method for eliminating biologic contaminants from diets (Coates, 1987; Foster et al., 1964; Williams et al., 1968~. However, this process can decrease the concentrations of heat-labile nutrients (Zimmerman and Wostmann, 19631. To ensure that adequate amounts of the most heat-labile vitamins (e.g., vitamins A and C and some of the B complex) will remain after autoclaving, consideration should be given to purchasing autoclavable diets that have been fortified with those vitamins. The magnitude of fortifi- cation in autoclavable diets is not generally high enough to be toxic to rodents; however, the routine use of autoclavable diets without autoclaving is not recommended, because the increased vitamin concentrations could influence experimental results. The level of sterility required for axenic or microbiologically associ- ated rodents requires that the temperature of the diet be raised above 100°C (212°F). To ensure that all the diet in the autoclave attains this tempera- ture, it is recommended that the diet be exposed to a temperature of 121°C (250°F) for 15-20 minutes. Diets should not be subjected to the maximal
HUSBANDRY 63 autoclaving temperature longer than necessary to achieve sterilization (Coates, 1987). To ensure proper operation of the autoclave, sterility of the diet, and adequate concentrations of labile nutrients, validation procedures are required, including periodic evaluation of autoclave operation by qualified personnel, use of commercially available heat indicators, culture of autoclaved feed samples for biologic contaminants, and assay of autoclaved feed samples to verify nutritional adequacy. Clarke et al. (1977) have described procedures for sam- pling and assaying feeds for various pathogenic organisms and provided stan- dards for the number and kinds of organisms that are acceptable in diets. Autoclaving at 80°C (176°F) for 5-10 min is required for pasteurization of diets. At that temperature, vegetative forms, but not spores, of microor- ganisms are destroyed (Coates, 1987~. Pasteurized diets are generally ac- ceptable for use in both specific-pathogen-free and conventional rodent colonies. Pasteurization, rather than sterilization, is used because there is less nutrient loss, and the diets are more readily consumed than are sterilized diets. Laboratory rodent diets also can be decontaminated by ionizing radia- tion (Coates, 1987; Coates et al., 1969; Ley et al., 1969), and diets sterilized in this way are now commercially available. Ethylene oxide fumigation has also been used to decontaminate diets (Meter and Hoag, 19661. All animal diets, particularly those produced from natural ingredients, can contain or become contaminated with various manufactured or naturally occurring chemicals, including pesticide residues, bacterial or plant toxins, mycotoxins, nitrates, nitrites, nitrosamines, and heavy metals (Fox et al., 1976; Newberne, 1975; Yang et al., 1976~. Procedures, if any, for detecting these chemicals are often difficult and expensive. Testing for contaminant concentrations in natural-ingredient diets should be routine in toxicologic research and might be valuable in some other studies. On the basis of observed contaminant concentrations and potential toxic effects, Rao and Knapka (1987) developed a list of recommended limits for about 40 chemical contaminants. The authors also proposed a scoring sys- tem for diets used in chemical toxicology studies that permits separation of tested diets into those acceptable for long-term use, those acceptable only for short-term or transitory use, and those which should be rejected. Laboratory animal diets designated as "certified" are commercially avail- able. Although the term is subject to different interpretations, in most cases the certification guarantees that the concentration of each contaminant on a specific list will not exceed the indicated maximum. Because the maximal concentrations usually are established by the diet manufacturer, the use of certified diets might not be appropriate for studies in which the acceptable concentrations of contaminants could influence experimental data indepen- dently or through an additive effect. In addition, a diet might have contami- nants that are not included in the certification but are of concern in specific research projects.
64 RODENTS: LABORATORYANIMAL MANAGEMENT Caloric Restriction Traditionally, the criterion used to evaluate laboratory rodent diets for nutritional adequacy has been maximal growth or reproduction of the ani- mals consuming the diet. Laboratory rodents generally are given ad libitum access to such diets throughout their lives. However, during the past 60 years, many studies have shown beneficial effects of caloric restriction in various species, including laboratory rodents (Bucci, 1992; Snyder, 1989; Weindruch and Walford, 1988; Yu, 19909. It has been reported that caloric restriction increases life expectancy and life span, decreases the incidence and severity of degenerative diseases, and delays the onset of various neoplasias. The objective of caloric restriction is to reduce calories without malnour- ishing the animals. That objective is generally accomplished by supplement- ing a diet with micronutrients and then limiting dietary consumption to 60-80 percent of the dietary consumption of animals that are fed ad libitum; this procedure results in decreased total caloric consumption. Although studies have been conducted in which the total fat (Iwasaki et al., 1988), protein (Davis et al., 1983; Goodrick, 1978), or carbohydrate (Kubo et al., 1984; Yu et al., 1985) consumption has been limited individually, only reduction in caloric intake results in the full range of dietary-restriction-related beneficial effects. Hypotheses explaining the results of dietary restriction studies have been re- viewed and discussed (Keenan et al., 19943. Numerous questions still need to be addressed to determine by what mechan- isms dietary or caloric restriction influences various life processes, and the quantitative nutrient or energy requirements necessary to achieve the effects associated with dietary restriction have not been established. However, the reported data show that ad libitum feeding might not be universally desirable for rodents used in long-term toxicologic or aging studies, and this factor should be a prime consideration when designing such studies. WATER Laboratory rodents should have ad libitum access to fresh, potable, uncontaminated drinking water, which can be provided by using water bottles and drinking tubes or an automatic watering system. Occasionally, it is necessary to train animals to use automatic watering devices. If water bottles are used, it is better to replace than to refill them; however, if they are refilled, each bottle should be returned to the cage of origin to minimize potential cross-contamination with microbial agents. If automatic watering devices are used, they should be examined routinely to ensure proper opera- tion. The drinking nozzles on these devices should be sanitized regularly, and the pipe distribution system should be flushed or disinfected routinely. Water is a potential source of microbial or chemical contaminants. A1- though a water source might be in compliance with standards that ensure
HUSBANDRY 65 purity of water supplied for human consumption, additional treatment might be required to ensure that water constituents do not compromise animal- colony objectives. Treatments used to limit or eliminate bacteria in water intended for laboratory rodents maintained in axenic or SPF environments include distillation, sterilization by autoclaving, hyperacidification, reverse osmosis, ultraviolet treatment, ultrafiltration, ozonation, halogenation, and irradiation (Bank et al., 1990; Engelbrecht et al., 1980; Fidler, 1977; Green and Stumpf, 1946; Hall et al., 1980; Hann, 1965; Hermann et al., 1982; Kool and Hrubec, 1986; Newell, 1980; Tobin, 1987; Tobin et al., 1981; Wegan, 19821. The advantages, disadvantages, and potential effects of water treatment on an animal's physiologic response to experimental treat- ments should be evaluated before a method of water decontamination is initiated. In general, any treatment that decreases water consumption is potentially detrimental to the animals' health and welfare. Drinking water of animals used in toxicology experiments, particularly those of long duration, should be periodically assayed for compounds that might influence experimental results, even when exposures are small. Mineral concentrations in water can have a profound influence on experimental results in studies designed to establish dietary mineral requirements for laboratory rodents. Distilled or deionized drinking water should be provided to rodents used in studies in which the amounts of minerals consumed are critical. BEDDING Bedding materials are used to absorb spilled water, minimize urinary and fecal soiling of the animals, and assist in decreasing the generation of odors and gaseous contaminants caused by bacterial decomposition of urine and feces. Bedding material can be used either as contact bedding in solid- bottom cages or as noncontact bedding in waste-collection pans placed be- neath wire-bottom cages. Contact bedding provides thermal insulation for the animals and is often used as nesting material in breeding colonies. Abrasive or toxic materials should not be used as contact bedding. Most products used for bedding in rodent colonies are byproducts of vari- ous industries. During the manufacturing process, these byproducts are occa- sionally subjected to conditions that are conducive to microbial contamination. Many of the commercially available rodent bedding materials are subjected to heat treatment before packaging; however, microbiologic recontamination can occur during shipment from the manufacturing plant to the animal facility. For maximal protection from potential microbiologic contamination, contact and noncontact bedding products should be sterilized before use. Hardwood and softwood are the most commonly used rodent bedding materials. Wood products should be screened to eliminate splinters or sliv- ers and should be free of foreign materials, such as paint, wood preserva
66 RODENTS: LABORATORY ANIMAL MANAGEMENT fives, chemicals, heavy metals, and pesticides. Some manufacturers will provide an assurance that the bedding is free of specified contaminants. The moisture content of wood products should be high enough to prevent excessive dust but low enough to provide adequate absorbency. Cedar- wood products are often mixed with other bedding material to mask animal- room odors; however, their use is not recommended because the aromatic hydrocarbons inherent in these products can alter hepatic microsomal en- zyme activity and potentially influence experimental results (Cunliffe-Beamer etal., 1981;Ferguson, 1966;Porter and Lane-Petter, 1965;Vesell, 1967; Vesell et al., 1976~. Furthermore, masking animal-room odors with cedar products is not a substitute for good sanitation practices. Plant byproducts and other cellulose-containing materials (including ground corncobs) are readily available as bedding for laboratory rodents. Laminated-paper products are available for use in waste-collection pans, and shredded-paper products are marketed for use as contact bedding for rodents. Corncob and paper products treated with germicides or antibiotics to control bacterial growth are also available. However, the routine use of antibiotic-treated bedding materials might cause antibiotic-resistant strains of bacteria to develop or influence experimental results. Bedding products manufactured specifically for use as rodent nesting materials are available. The use of such products, which might enhance neonatal survival in inbred rodent strains with inherently low reproduction rates, should be considered. All rodent bedding products should be packaged in sealed, nonporous bags. Bags of bedding material should be stored in vermin-proof areas on pallets that do not touch the walls. When the bedding material is removed from the bags, it should be stored in metal or plastic containers that can be closed securely. The storage containers should be sanitized routinely. SANITATION Cleaning Adequate sanitation is an integral part of maintaining laboratory ro- dents. Clean, sanitary conditions limit the presence of adventitious and opportunistic microorganisms, thereby decreasing their potential for com- promising rodent health or causing adverse interactions with experimental procedures. Complete sterilization of the rodents' environment is seldom practical or necessary unless animals of highly defined microbiologic status or compromised immune status are used. All components of the animal facility should undergo regular and thor- ough cleaning, including animal rooms, support areas (e.g., storage areas), cage-washing facilities, corridors, and procedure rooms. They should be cleaned with detergents and, when appropriate, disinfectant solutions to rid
HUSBANDRY 67 them of accumulated dirt and debris. Many such products are available. Selection of a cleaning agent should be based on how much and what kind of material is adhering to surfaces, as well as on the type of microbiologic contamination present (Block, 1991~. Monitoring of sanitation procedures should be appropriate to the pro- cess and materials used and might include visual inspection, monitoring of water temperatures, and microbiologic monitoring. It has been suggested that the effectiveness of sanitation procedures can be assessed by the inten- sity of animal odors, particularly ammonia; however, this should not be the sole means of assessing cleanliness, because too many variables are in- volved. Agents used to mask animal odors should not be used in rodent housing facilities; these agents cannot substitute for good sanitation prac- tices, and their use exposes animals to volatile substances that can alter basic physiologic and metabolic processes. The frequency with which surfaces are cleaned should be determined by how much use an area receives and the nature of potential contamina- tion. Sweeping, mopping, and scrubbing with disinfectant agents should take place in a logical sequence. Cleaning utensils should be constructed of materials that resist corrosion and do not absorb dirt or debris. They should be stored in a neat, organized fashion. Wall-mounted hangers are useful for storing cleaning utensils because they reduce clutter, facilitate drying, and minimize contamination by keeping utensils off the floor. Cleaning utensils should be assigned to specific areas and should not be transported between areas. They should be regularly cleaned and dried, and there should be a regular schedule for replacing worn-out utensils. Soiled bedding material should be removed and replaced with clean, drY bedding as often as is necessary to keep the animals clean and dry. The ~ _ ~ . . . ~- ~ ' 1 · ~ ~ 1_ _ _ _ 1 1 1~ _ 1~ _ ~ ~ 1 _ _ frequency Is a matter of professional Judgment and snout oe uas';u use various factors, including the number and size of the animals housed in each cage, the anticipated urinary and fecal output, and the presence of debilitating conditions that might limit an animal's ability to access clean areas of the cage. Bedding should be changed in a manner that reduces exposure of the animals and personnel to aerosolized waste materials. Laminar-flow bed- ding dump stations or similar devices can be used to control aerosol materi- als. If animals have been exposed to hazardous materials that are excreted in the urine or feces, additional precautions might be needed to prevent exposure of personnel while they are changing the bedding. Frequent bedding changes can sometimes be counterproductive, for ex- ample, during portions of the postpartum period, changing the bedding re- moves pheromones, which are essential for successful reproduction (e.g., pheromones are necessary for synchronization of ovulation). Research ob- jectives might also preclude frequent bedding changes. Under such circum 1
68 RODENTS: LABORATORYANIMAL MANAGEMENT stances, an exception to the regular bedding-change and cage-cleaning schedule can be justified. Cages, cage racks, and accessory equipment, such as feeders and water- ing devices, should be cleaned and sanitized regularly to minimize the buildup of debris and to keep them free from contamination. Extra caging makes it easier to maintain a systematic schedule. Cleaning frequency will depend on the amount of bedding used, the frequency of bedding changes, the number of animals per cage, and other factors. In general, rodent cages and cage accessories will need to be washed at least once every 2 weeks. Solid- bottom rodent cages, water bottles, and sipper tubes usually require weekly cleaning. Some types of cage racking, large cages with very low animal density and frequent bedding changes, cages housing animals in gnotobiotic conditions, and cages used under other special circumstances might require less frequent cage-cleaning. Filter-top cages without forced-air ventilation and cages containing rodents with increased rates of production of feces or urine might require more frequent cleaning. Cage-cleaning, debris removal, and disinfection can be accomplished in a single step or in multiple steps. Cage-cleaning and debris removal usually require the application of a detergent or surfactant solution coupled with me- chanical action to remove adherent material from cage surfaces. Some labora- tory rodents, such as guinea pigs and hamsters, produce urine with high con- centrations of proteins and minerals. Their urine often binds aggressively to cage surfaces, which therefore require treatment with acid solutions before washing. Some detergents are rendered inactive at high temperatures, so, it is important to follow the manufacturer's instructions carefully. Disinfection of cages is the process of killing vegetative forms of pathogenic bacteria. It can be accomplished by the action of either chemicals or hot water. If chemicals are used as the sole means of disinfection, careful attention should be paid to the concentration of the disinfectant solution's active ingredients, and the solution should be regularly changed in accor- dance with the manufacturer's instructions. When hot water is used either alone or in combination with disinfectant chemicals, temperatures and ex- posure times should be appropriate for adequate disinfection. Generally, the water temperature required for adequate disinfection precludes its use in anything but mechanical cage-washing equipment. Cleaning and disinfection of cages can be done efficiently in mechani- cal cage washers. Washing times and conditions should be sufficient to kill vegetative forms of common bacteria and other microorganisms that are presumed to be controllable by sanitization. Microorganisms are killed by a combination of heat and the length of exposure to that heat (called the cumulative heat factory. Using high temperatures for short periods will produce the same cumulative heat factor and have the same effect on micro- organisms as using lower temperatures for longer periods (Wardrip et al.,
HUSBANDRY 69 1994~. To achieve effective disinfection, water temperatures for washing and rinsing can vary from 58°C (143°F) to 82°C (180°F) or more. Recom- mendations for some types of mechanical cage washers using hot water alone for disinfection have been developed by the National Sanitation Foun- dation International (1990~. Detergents and chemical disinfectants are known to enhance the effectiveness of hot water but must be thoroughly rinsed from surfaces to avoid harm to personnel and animals. Cages and equipment can be effectively washed and disinfected by hand if appropriate attention is given to detail. Chemicals should be com- pletely rinsed from surfaces, and personnel should have appropriate equip- ment to protect them from prolonged exposure. Large pieces of caging equipment, such as racks, can be washed by hand; if large numbers are to be cleaned, portable cleaning equipment that dispenses detergent and hot water or steam under pressure might be more efficient. Large mechanical washing machines designed to accommodate racks and other pieces of large equipment are also commercially available. Water bottles, sipper tubes, stoppers, and other small pieces of equip- ment should be washed with detergents, hot water, and, if appropriate, chemical agents to destroy vegetative forms of microorganisms. This process can be manual, if high-temperature rinse water is not used, or performed with me- chanical washing equipment built especially for this purpose or a multiple -~ r ~1 · , . , ~1 purpose cage-washing machine. Water bottles and sipper tubes can also be autoclaved after routine washing to ensure adequate sanitation. If large numbers of water bottles or other small pieces of equipment are to be washed by hand, powered rotating brushes can be used to ensure adequate cleaning. Small items should be dipped or soaked in detergent and disinfec- tant solutions to maximize contact time. Therefore, large, two-compartment sinks are generally required if small items are to be hand washed. If automatic watering systems are used, they should incorporate some mechanism to ensure that bacteria and debris do not build up in the water- ing devices. These systems are usually periodically flushed with large volumes of water or appropriate chemical agents and then rinsed to remove chemicals and associated debris. Constant-recirculation loops that use fil- ters, ultraviolet light, or other treatment procedures to sterilize recirculated water can also be used. Common methods of disinfection and sanitization are adequate for most rodent holding facilities. However, if pathogenic microorganisms are present or if rodents with highly defined microbiologic flora or compromised immune systems are maintained, it might be necessary to sterilize caging and other associated equipment after cleaning and disinfection. In such instances, access to an autoclave, gas sterilizer, or device capable of sterilizing with ionizing radiation is required. Whenever such sterilization processes are used, some form of regular monitoring is required to ensure their effectiveness.
70 RODENTS: LABORATORYANIMAL MANAGEMENT Waste Containment and Disposal Proper sanitation of an animal facility requires adequate containment, as well as regular and frequent removal of waste. Waste containers should be constructed of either metal or plastic materials and should be leakproof. They should be equipped with tight-fitting lids and, where appropriate, pro- vided with disposable plastic liners for ease of waste removal. They should also be adequately labeled to distinguish between containers for hazardous and nonhazardous wastes; a color-coding system often proves useful. If hazardous biologic waste is generated, an inventory sheet might be necessary for each waste container, so that the type of waste and the approxi- mate quantity of hazardous material can be recorded. Waste containers for animal tissues or carcasses should be lined with leakproof, disposable liners that will withstand being refrigerated or frozen to reduce tissue decomposition. If wastes are collected and stored before removal from the site, the storage area should be physically separated from other facilities used to house animals or store animal-related materials; Waste-storage areas should be cleaned regu- larly and kept free of insects and other vermin. All waste containers and associated implements should be cleaned and disinfected frequently. Waste materials from rodent housing facilities can be disposed of in various ways (depending on the type of waste), including incineration, agri- cultural comporting, and landfill disposal. Hazardous waste must be sepa- rated from other waste, and its classification and handling are controlled by a variety of local, state, and federal agencies. Some form of pretreatment- such as autoclaving, chemical neutralization, or compaction with absorbents- might be required. The National Safety Council (1979) has recommended procedures for disposal of hazardous waste. It is the institution's responsi- bility to comply with all federal, state, and municipal statutes and ordi- nances regarding the control, movement, and disposal of hazardous waste. Pest Control All rodent housing facilities should have a program to prevent, control, or eliminate infestation by pests (including insects and wild and escaped rodents). The program should include regular inspection of the premises for signs of pests, a monitoring system that uses rodent traps and insect- collection devices to capture pests, and regular evaluation of the integrity and condition of the animal facilities. The pest-control program should focus on preventing the entry of vermin into the facility (by sealing poten- tial points of entry and eliminating sites outside the facility where vermin can breed or be harbored) and maintaining an environment in which pests cannot sustain themselves and reproduce. Only if those methods are inef- fective should the use of pesticides be considered.
HUSBANDRY 71 If pesticides are required, relatively nontoxic substances (e.g., boric acid, amorphous silica gel, and insect-growth regulating hormones) and me- chanical devices (e.g. ' ~ ' ' '~ devices) should be used in preference to toxic materials, especially for controlling insect pests. If a toxic compound is to be used in animal areas, it should be used only after consultation with the investigators whose ani- mals are housed in the facility because of potential effects on the animals' health and possible interference with research results. The application of toxic pesticides should be coordinated with those responsible for the man- agement of the animal-care program and carried out by licensed applicators in compliance with local, state, and federal regulations. The pest-control program should be adequately documented, including records of dates and methods of application of pesticides and possibly records of'inspection, results of monitoring and trapping programs, records of sightings and identification of pests, and maintenance schedules. , adhesive traps, air curtains, and ~nsec~-e~ec~rocuuon IDENTIFICATION AND RECORDS Adequate individual or group identification of rodents and appropriate records of their care and use are essential to the conduct of biomedical research programs. Individual identification of Rodents is not always re- quired; when necessary, it can be accomplished in various ways, including ear-punching, use of ear tags, tattooing (usually on the tail), or implanting electromagnetic transponders. If ear tags are used, they should be light enough so that they do not visibly change the animal's head posture, and surrounding tissues should be monitored for inflammation. Dyes are occa- sionally used on the fur, skin, or tail for temporary identification. In gen- eral, amputation of digits (toe-clipping) is no longer an acceptable method of identification, because more humane methods can usually be substituted. Individual animals or croups of animals can also be identified with cage identification cards. ~, If cards are used sufficient information is re quired to identify and characterize the animals in the cage adequately. This information can include such details as the name and location (e.g., office location, telephone number, and division or department name) of the re- sponsible investigator; the species, strain, or stock of the animals; the sex of the animals; the number of animals in the cage; the source of the animals; institutional identification numbers (e.g., IACUC-approved protocol num- ber and purchase-order number); and, when appropriate, other identifying information pertaining to the project (e.g., group designation and age or weight specifications). Bar-code identifiers can also be included on the cage card to aid in identifying the animals and linking their identification with other, more detailed records. Color-coding the cage cards and labeling _ . .
72 RODENTS: LABORATORYANIMAL MANAGEMENT cage racks and animal holding rooms are effective management tools for locating and identifying animals. Some research protocols require that records be kept on individual ani- mals, for example, when animals are used in breeding programs or are exposed to hazardous agents. Detailed surgical records are not commonly maintained on individual rodents but might be helpful in some situations such as when complex surgical procedures are being used or when new procedures are being developed. RODENTS OTHER THAN RATS AND MICE Guinea Pigs One of the most striking ways in which guinea pigs (Cavia porcellus) differ from rats and mice is the guinea pigs' absolute requirement for exog- enous vitamin C, a requirement that is shared with humans and only a few other species. Because of that requirement, guinea pig diets must be forti- fied with vitamin C. As an alternative, vitamin C can be added to the drinking water or provided in the form of food supplements, including such vegetables as kale, that are high in vitamin C. The use of food supplements should be approached with some caution because of the possibility of con- tamination with chemicals or microorganisms that could influence the course of experimentation. Vitamin C is a very labile compound, so storage condi- tions of foods containing it and heat treatment of such foods, including autoclaving, are of particular concern. The guinea pigs' body conformation makes design and placement of feeders important. Feeders should be designed to avoid trauma to the chin and neck area of guinea pigs. Guinea pigs will occasionally rear up on their hind legs, but they will not accept food from feeders suspended overhead. Bowls for food and water can be used instead of more conventional feeding and watering devices; but guinea pigs like to nest in such receptacles, and that causes waste and contamination of food. Feeders that have a J shape are best suited to address these concerns and are used most commonly. Guinea pigs, like other rodents, tend to eat and drink throughout the day and night. They become habituated to a particular diet and have de- fined taste preferences. Any changes in the composition of the food- especially changes in size, shape, consistency, or taste-can cause a sharp decline in food consumption. If the animals fail to adapt to the new food, severe weight loss or even starvation and death can occur; therefore, new food should be introduced gradually. Guinea pigs often grow to weigh more than 1 kg and have relatively small feet. They have a well-developed startle response that causes them to make sudden movements in response to unfamiliar sounds; when they are
HUSBANDRY housed in groups 73 , this might be manifested as a stampede. Those two traits make cage-floor design particularly important. Wire-bottom cages should be designed to provide sufficient support for the animals' feet to prevent pressure sores, and the space between the wires in the floor grid should be small enough to preclude entrapment of animals' feet. Guinea pigs also differ substantially from rats and mice in having a vaginal closure membrane and a long gestation period. Gestation in guinea pigs can range from 59 to 72 days; 63 to 68 days is the average. Gestation length can be affected by several characteristics, including litter size, which is usually one to three pups (McKeown and Macmahon, 1956~. Female and male guinea pigs reach puberty as early as 4-5 weeks old and 8-10 weeks old, respectively, but are best mated when 2.5-3 months old or when they weigh 450-600 g (Ediger, 1976~. Because a relatively large fetal mass is expelled at parturition, a female should be bred before she is 6 months old to minimize the likelihood of being excessively fat or having firm fusion of the symphysis pubis. If the symphysis pubis is fused, it cannot separate the approximate 0.5 in. needed for passage of fetuses through the birth canal; the result can be severe reproductive problems and death of both fetus and mother. Strain 13 guinea pigs, which are highly inbred, should be housed to protect them from or immunized against the common bacterium Bordetella bronchiseptica (Ganaway et al., 1965~. Treating guinea pigs for bacterial infections should be approached with caution because antibiotics can cause acute effects. Some can be administered safely; others, such as penicillin, can cause toxemia and death (fakes et al., 1984; Wagner, 1976~. The problem appears to be associated with the excretion of the antibiotics into the gastrointestinal tract and the resulting disturbance of the microbiologic flora on which the guinea pig depends for much of its digestive processes. Guinea pigs produce large volumes of urine that contain substantial quantities of dissolved minerals and protein. Their urine adheres tena- ciously to surfaces, and soaking in dilute solutions of organic acids is often required before cages are cleaned. Urination and dragging the perineum across the floor of the cage are common methods by which guinea pigs mark freshly cleaned cages. Hamsters Laboratory hamsters belong to the subfamily Cricetidae. The most common and most readily available commercially is the Syrian hamster, Mesocricetus auratus (sometimes called the golden hamster). Syrian ham- sters are native to arid regions of the Middle East and have become well adapted to conserving water, which they obtain principally through food. In a laboratory environment, hamsters will drink water from water bottles,
74 RODENTS: LABORATORYANIMAL MANAGEMENT bowls, or automatic watering systems. Hamsters secrete highly concen- trated urine that contains large quantities of mineral salts; their urine tends to leave deposits on cage surfaces that are often difficult to remove and might require the application of dilute acids. Hamsters are often aggressive toward each other, and care should be taken when they are housed in groups. Hamsters that fight must be sepa- rated to prevent injury. Cannibalization can occur in group-housed animals when an animal becomes sick or debilitated. It is important to separate animals that are observed to be clinically abnormal. Vitamin E is an important nutritional requirement of hamsters; vitamin E deficiency has been associated with muscular dystrophy (West and Ma- son, 1958J and fetal central nervous system hemorrhagic necrosis (Keeler and Young, 19793. Most commercial rodent diets are supplemented with vitamin E, but care is required to ensure the adequacy of vitamin E if special-formula, purified, or semipurified diets are used (Balk and Slater, 1987~. The method of food presentation is important. If food is placed in suspended feeders, hamsters will remove it from the feeder and pile it on the floor. Location of the food pile is peculiar to individual hamsters and will vary from one cage environment to the next. Moving food away from a pile will cause the hamsters to retrieve it and move it back. Given that behavioral pattern, feeding hamsters on the floor of the cage is considered acceptable (9 CFR 3.291. Hamsters have cheek pouches in which they hold and transport food; a full cheek pouch should not be mistaken for a pathologic condition. Hamsters have very loose skin, particularly over the shoulders. Care should be taken when picking them up so that they do not turn around and bite the handler. Hamsters can be tamed by regular, gentle handling. With- out such taming, they can be aggressive toward the handler. Many species of hamsters hibernate if conditions are right. Various environmental influences seem important, including seasonality, photope- riod, ambient temperature, availability of food, and isolation. To avoid hibernation, temperatures should be maintained within ranges specified in the Guide (NRC, 1996 et seq.~. Hamsters, like guinea pigs, are susceptible to antibiotic associated tox- icity and enterocolitis. Although successful use of antibiotics in hamsters has been reported, the reports usually involve smaller than therapeutic dos- ages of antibiotics or the use of particular antibiotic preparations that are not excreted into the gastrointestinal tract (fakes et al., 1984; Small, 19871. As a general rule, antibiotics should be avoided in hamsters. Estrus in hamsters is similar to that in mice, lasting 4-5 days; however, the gestation period is considerably shorter than that in mice an average of 16 days. Hamsters are commonly pair-mated; the female is taken to the male's
HUSBANDRY 75 cage for breeding on detection of a stringy vaginal discharge that occurs when the female is in estrus. The female can be removed from the male's cage after mating is observed; however, conception is sometimes improved by leaving her with the male for 24 hours. Removing the female after that time mini- mizes fighting and allows the male to breed with other females. For optimal reproduction, the light cycle should be maintained at 14 hours of light and 10 hours of dark, which is slightly different from that for other rodents. Litter size ranges from 4 to 16 pups; first litters tend to be smaller than subsequent litters. Cannibalism of pups is common, especially in first litters. It is impor- tant to furnish enough bedding or nesting material for the neonates to stay well hidden and to provide the dam with enough food to allow her to be undis- turbed from about 2-3 days before birth until about 7-10 days after birth (Balk and Slater, 1987; Harkness and Wagner, 19891. Gerbils Gerbils (Meriones unguiculatus) do well in solid-bottom cages. Gerbils tend to stand and sit upright and often exhibit a digging or scratching be- havior in the corners of cages while in an upright posture. Therefore, cages that are tall enough for this behavior are generally preferred. Gerbils tend to form social relationships early in life, and groups estab- lished at puberty tend to exhibit minimal fighting or other aggressive be- havior; aggressive behavior is more common when individual animals are put together later in life. New mates are not accepted easily. For those reasons, it is prudent to select a paired-mating scheme for establishment of colonies and not to regroup gerbils often. Estrus in gerbils lasts 4-6 days; gestation in nonlactating females is about 24-26 daYs. If females are bred in the postpartum period, implanta- tion is delayed, and gestation can be as long as 48 days A, To avoid postpar tum mating, the male can be removed from the cage, but he should be returned to his mate within 2 weeks to decrease the possibility of fighting (Harkness and Wagner, 19891. Average litter size is 3-7. Gerbils are generally very tame and rarely bite unless mishandled. When they are excited, they will jump and dart about to resist being caught. Gerbils should not be suspended by holding their tails, because the skin over the tail is relatively loose and can be pulled off easily. Commercial rodent diets are usually acceptable for gerbils, provided that they have a low fat content. Because of the gerbils' unique fat metabo- lism, it is not uncommon for them to develop high blood cholesterol con- centrations on diets containing fat at 4 percent or more. When fed a diet high in fat, gerbils tend to store the fat and become obese. In females, the fat accumulation can be associated with reproductive difficulty.
76 RODENTS: LABORATORYANIMAL MANAGEMENT Chinchillas Chinchillas (Chinchilla laniger) have been farmed for pelts since 13 animals were imported from South America to California in 1927. Most domestic stock is believed to be descended from those animals (Anderson and Jones, 1984~. Chinchillas can be housed in wire-mesh or solid-bottom cages; the latter are preferred for breeding (Clark, 1984; Weir, 19761. They are fastidious groomers and should be provided with a box containing a mixture of silver sand and Fuller's earth for a short period daily to allow dust bathing (Clark, 1984~. Chinchillas tolerate cold but are very sensitive to heat; the suggested temperature is 20°C (68°F) (Weir, 19761. Commercial chinchilla feed is available, but standard guinea pig rations can also be used (Clark, 1984; Weir, 1976~. They might require a source of roughage, such as hay (Weir, 1967~. Water and food should be made available ad libitum. The system used most commonly for breeding chinchillas is to put one male with several females in a large cage. However, females are larger than males and are very aggressive toward both males and other females, and it is necessary to provide refuges, such as nesting boxes, for animals that are being attacked. An "Elizabethan collar" can be used to keep an aggressive female from following an animal that she is attacking into its refuge. A light:dark ratio of 14:10 hours is adequate (Weir, 1967~. The mean gesta- tion period is 111 days, with a range of 105-118 days (Clark, 19841. Chin- chilla litter size ranges from one to six, with a mean of two. The young are born fully furred and with open eyes, and they begin eating solid food within 1 week but are not completely weaned until they are 6-8 weeks old. Females do not build nests. REFERENCES Algers, B., I. Ekesbo, and S. Stromberg. 1978. The impact of continuous noise on animal health. Acta Vet. Scand. 67(Suppl.): 1 -26. Alleva, J. J., M. V. Waleski, F. R. Alleva, and E. J. Umberger. 1968. Synchronizing, effect of photoperiodicity on ovulation in hamsters. Endocrinology 82:1227-1235. Anderson, K. V., F. P. Coyle, and W. K. O'Steen. 1972. Retinal degeneration produced by low-intensity colored light. Exp. Neurol. 35:233-238. Anderson, S., and J. K. Jones, Jr., eds. 1984. Orders and Families of Recent Mammals of the World. New York: John Wiley and Sons. 686 pp. Anthony, A., and J. E. Harclerode. 1959. Noise stress in laboratory rodents. II: Effects of chronic noise exposures on sexual performance and reproductive function of guinea pigs. J. Acoust. Soc. Am. 31:1437-1440. ASHRAE (American Society Heating, Refrigeration, and A Engineers, Inc.). 1993. Chapter 9: Environmental Control for Animals and Plants. In 1993 ASHRAE Handbook: Funda mentals, I-P edition. Atlanta: ASHRAE Baetjer, A. M. 1968. Role of environmental temperature and humidity in susceptibility to disease. Arch. Environ. Health 16:565-570. Balk, M. W., and G. M. Slater. 1987. Care and management. Pp. 61-67 in Laboratory Ham- sters, G. L. Van Hoosier, Jr., and C. W. McPherson, eds. Orlando, Fla.: Academic Press.
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