TY - JOUR
AU1 - Hasenau, John, J
AB - Abstract As mentioned in other chapters, reproducibility of research data is very complicated and has numerous contributors for concerns. This chapter will discuss the animal housing systems and corresponding husbandry practices in regard to current practices and known and potential confounders in the research environment. This area has a very high impact for reproducibility and comparability of study data outcomes. Housing Systems and Husbandry: Terrestrial Animals the Basics General Animal Housing Concepts Types of Animal Rooms. Basically, animal rooms can be divided into 2 types: (1) rooms for housing animals using dry bedding cage systems, generally for housing small animals from rodents to rabbits; and (2) rooms for housing animals using hose-down caging systems, generally for housing nonhuman primates, canines, and small agriculture animals in cages or floor pens. One room design approach is a flexible system to accommodate either type of housing system, allowing maximum flexibility at a higher related cost; another approach is to design rooms for either one system or the other. This design aspect can have impacts on macroenvironmental conditions, with hose-down environments having greater variability and range in humidity and temperatures. The hose-down caging system requires floors sloping to floor drains, preferably in troughs, and the presence of a hose. This sloping of floors, critical to avoid pooled water, can be problematic in flexible rooms for rodent rack rolling or having the actual animals cage at an angle depending on the degree of slope. This can be offset by using wedges under the locked wheels to allow for level cages. The dry bedding housing system does not require floor drains or sloped floors. The disadvantage is that a room designed for both types of housing systems is not optimal for either. If the facility or parts of it only houses rodents (eg, a rodent barrier facility), then it is reasonable is to design it with no floor drains or sloping floors. These different configurations will have different inherent effects on the housed animals as well and are important considerations in study comparisons and evaluations. The size and shape of the animal room can vary depending on many factors, including the species to be housed, the type of housing system to be used, and the arrangement of the cages and racks in the room. Most rooms are sized based on the cage type used and support columns and beams infrastructure of the building. Caging layouts are usually done to maximize density. As an example, double-sided rodent racks are typically arranged library style, with multiple racks parked parallel and the end of each rack against a common wall, or with 2 rows on opposite walls with an aisle between them. Single-sided cage racks are typically parked with the back of the rack against a wall. A combination of both types combines the advantages of both. Conventional Animal Housing Conventional is a generic term, with no specific definition, referring to almost any type of laboratory animal housing facility, area within a facility, or animal room that is not specially designated otherwise (as a barrier or containment area). All animal rooms, whether conventional, barrier, or containment, are designed for ease of cleaning and have minimal built-ins. Animal rooms historically have been designed with operational efficiencies and staff conveniences as the primary driver; more recently animal behavioral needs focused on decreasing stress and distress have been emphasized. Barrier Animal Housing A barrier facility has come to be known as an animal housing system designed and managed to protect animals from potentially pathogenic or undesirable microbes. “Barrier” indicates “keeping these adventitious agents out” of animal models. The original primary use of barriers was to produce laboratory rodents. A similar level of barrier housing in the research environment has extended the need for barrier housing to the research facility. This increased need expanded further with the extensive use of immune-compromised animals and genetically modified rodents. Due to these concerns, barrier space may include space for wet laboratories, animal procedure laboratories, transgenic animal housing, specialized imaging equipment, irradiation equipment, etc. Studies that have been conducted within a barrier area should be identified as such in published articles. In all cases, the type of barrier should be specified as to set up and operation to better understand potential data impacts. Containment Animal Housing Containment refers to animal housing systems designed and managed to prevent the escape of experimental hazardous agents to which the animals have been exposed to protect workers, other animals, and the general environment. Containment equates to “keeping hazardous agents in” and minimizing staff exposures. The hazardous agents may be biological, chemical, or radiological. Containment, like barriers, can be achieved at the cage level, the room level, in an area within an animal facility, or the entire facility. There are many similarities between barrier and containment practices and procedures. When used together (containment within a barrier area) there are increased levels of containment. At all levels of containment, the primary objective is to contain the hazardous agent as close to the source as possible, ideally, at the cage level (eg, a microisolation cage or better, a hermetically sealed individual ventilated cages [IVC] cage). Microbiological agents are classified into 4 biosafety levels to BSL-4 according to the degree of risk to humans (classified by the CDC-NIH in the publication “Biosafety in Microbiological and Biomedical Laboratories,” USDHHS 2009). Animal infections with infectious agents designate the corresponding facilities and management practices as animal biosafety levels (ABSL) 1 to 4. Studies with BSL-2 agents are commonly conducted in conventional animal rooms using appropriate equipment and ABSL-2 practices, as this is the highest level of biocontainment that can be achieved in conventional rooms. When available, an ABSL-3 facility is highly desirable for quarantine of rodents that are infected with adventitious agents or that are of unknown health status. These agents are not hazardous to humans but have the potential to be devastating for many if not most of the rodent studies in the facility. The increased work in microbiota studies with complex microbiomes has increased these types of studies in research facilities [1, 2]. As with barrier, containment areas and systems need to be well defined and explained so that experimental replications and transability in the study environment can be understood. Quarantine For the purpose of this chapter, it is important to indicate in study reports if animals used are either maintained in or had passed through a quarantine area. Most facilities today require a high-level rodent quarantine facility for holding animals until they can be documented to not be exposed and/or carriers of known adventitious agents, or the genetic line can be rederived by embryo transfer (less desirable is C-section and fostering). Ideally, rodents of unknown health status or animals known to be infected with agents hazardous to other rodents in the facility are best maintained in an ABSL-3 facility, utilize an animal facility in which no other rodent colonies are housed, or are in a strict quarantine area located away from regular rodent housing areas. In all cases, measures need to be in place to minimize the risk of pathogen spread by fomites (such as use of dedicated caging and cage sanitation equipment), including directional air flows and strictly enforced traffic patterns and hygiene practices for personnel who may work with Specific Pathogen Free and conventional mice [3]. In all, full reporting of the passage through or the maintenance of study animals in quarantine needs to be reported. Caging Small Rodents Small rodents (eg, rats, mice, hamsters, gerbils, and guinea pigs) are usually housed in solid-bottom, shoebox-type cages with various types of bedding materials covering the cage bottom. Open, stainless-steel, wire-bottom cages without bedding material do not allow natural behaviors to be exhibited, decrease the ability of thermoregulation of the animals, and are considered less desirable in terms of animal welfare but are occasionally used when required to achieve scientific objectives. Large guinea pigs are sometimes housed in rabbit cages, especially ones with perforated plastic floors. Rodent shoebox-type cages are made of various types of plastic materials. Biosecurity and sterility needs that require repeated cleaning with laboratory-grade detergents and steam sterilization without physical cage deterioration determine the choice and the quality of thermoplastics used for sustainable caging. Sustainable caging currently uses polypropylene, polycarbonate, polysulfone, and polyphenylsulfone compositions. Polypropylene (eg, Fortilene [Solvay]) was one of the first thermoplastics used. Polypropylene is a light, flexible material with high chemical inertness and thermal resistance up to 120°C (250°F). It may be translucent or opaque, depending on whether it is a copolymer or a reinforced copolymer. Historically opaque composites were used, but with the need for daily animal checks they were displaced using polycarbonate (Lexan [Sabic] or Makrolon [Covestro]). Polycarbonate is transparent, rigid, durable, and sanitizable withstanding autoclaving at 120°C (250°F) but eventually becomes brittle after multiple exposures to autoclave temperatures. The material most frequently used at this time for sustainable caging is polysulfone (Urdel PSU [Solvay]) due to the ability to handle thousands of repeated autoclavings (134°C [273°F]) while maintaining impact strength and transparency. Polysulfones are marketed with a lifespan of approximately 3 years under conditions of normal use. Polysulfone’s initial higher cost has been cost effective due to high durability and related longevity of the material. Polyphenylsulfone (Radel PPSU [Solvay]) withstands higher temperatures 143°C (289°F); thus, it is often referred to as “high-temperature” autoclave caging and has higher impact strength, allowing a lifespan of approximately 8 years. With PSU or PPSU there is some rodent visual protection provided from light intensities (~35% less light penetration than polycarbonate) due to the natural amber or yellow coloration, which is more apparent with PPSU. With appropriate nesting materials, observation of the animals can be more difficult, which has resulted in a clear caging option for PPSU as well. All polycarbonate caging releases Bisphenol A, an endocrine disruptor, when damaged. Historical research papers provided some of the first evidence of the harmful effects of exposure to this common chemical in both mice and nonhuman primates (NHPs). A recent publication has indicated that damaged polysulfone cages can release Bisphenol S, another endocrine disruptor, as well [4]. Other plastic polymers are used for disposable rodent caging. Polystyrene is rigid, with low impact and heat resistance and is translucent. It is usually used to form disposable cages suitable for toxic or radioactive applications when decontamination of the sustainable cage is impractical or too dangerous for personnel. Cage type, age, and composition material (if known) should be mentioned for comparison purposes when writing a study report. Micro-Isolation Caging System(s) The micro-isolator caging system's major component is the micro-isolation cage. Two main types of micro-isolation caging designs exist: ventilated (commonly used in contemporary animal facilities) as the IVCs in their custom cage racks, or static (less common) as the static micro isolation units not requiring a specific rack style. In addition, 2 other primary components are required for proper micro-isolation function and use: (1) a HEPA-filtered air cabinet, often referred to as animal transfer stations or cabinets (ATSs), to be used anytime the cages are opened, either to transfer the animals to a clean cage or to perform experimental procedures with them; and (2) rigorous operational sanitation procedures to be used immediately prior to and during the time when the cage is opened. Micro-isolation cages for rodents have already been described in this chapter as being highly effective in both barrier and containment environments. Static micro-isolation caging may be more suited for containment of high-level hazardous agents than IVCs unless the ventilated cages are tightly sealed. Hermetically sealed IVCs with individual cage-level HEPA filtration (both in and out) are available for high-level hazardous agents and for use in microbiome work as an alternative to isolator housing. Regarding the ATS, class II type A biocontainment cabinets are well suited for use in barrier and containment situations since they are designed to protect the operator’s environment outside the cabinet from agents in the cabinet as well as the product inside the cabinet. The product is the mice in the opened cage. Protection is from contaminants outside the cabinet. When there is a disease threat (eg, mouse hepatitis virus), the management objective is to contain the offending infectious agent until it can be detected and eliminated. There are ATSs available that do both, keeping agents from getting into or escaping from the cabinet, that are more user friendly but that do not qualify to be classified as biocontainment cabinets. Which one is chosen will depend on how much emphasis one places on containment when working in a barrier setting. Considerations should also be given with regards to noise and vibration levels of the ATS and racks with blowers that could negatively impact the research [5, 6]. The rigorous operational sanitation procedures include wearing appropriate personal protective equipment or, at a minimum, protective sleeves to cover the arms and the hands with gloves and opening 1 cage at a time in the cabinet, except when both a soiled and a clean cage are required when transferring animals to a clean cage; between working with cages in the cabinet, using a fast-acting, high-level disinfectant on the cabinet work surface, the outside of the cage, the gloved hands, and any instrumentation that touches the animals in reality has the greatest impact on barrier and containment aspects of the operation. More emphasis has been placed on the types of disinfectants used and potential effects on animal behaviors [7] and animal generated study data, especially with research equipment [8]. This area is still in debate. Behavioral testing of Swiss Webster, C57Bl/6, and BALB/c mice showed that hypochlorite- and peroxide-based products were clearly aversive, and yet in elevated maze tests (EMT), cleaning the equipment compared with leaving it soiled did not affect performance in male or female C57 mice, nor did cleaning agent choice (isopropyl alcohol, chlorine dioxide, and bleach), but there was aversion of all cleaning agents noted with a separate 2-choice light/dark test. Static micro-isolation cages are very effective at protecting rodents from microbial contamination when used in combination with a HEPA-filtered mass air displacement cabinet when cages are opened [9]. These cages can have rapid elevations of humidity, ammonia, and carbon dioxide [10–13] necessitating more frequent cage changes (usually 2 times weekly) than rodents maintained in IVCs. There are concerns of increased stress on animals with frequent cage changes. Plasma corticosterone levels tended to be lower when cage-changing intervals were longer [14], and fecal corticosterone levels stayed even with longer cage change intervals [15]. Air Qualities Ventilated micro-isolation caging was designed to better control the microenvironment [11, 16]. Directly ventilating each micro-isolation cage with HEPA-filtered air significantly slows the buildup of ammonia, etc. in the cage, thus decreasing the cage change frequency to once a week or even once every 2 or 3 weeks. Direct exposure of animals to air moving at high velocity (drafts) should be avoided as the speed of air to which animals are exposed affects the rate at which heat and moisture are removed from an animal [17, 18]. If ventilated primary enclosures have adequate filtration to address contamination risks, air exhausted from the microenvironment may be returned to the animal housing room, although it is generally preferable to exhaust these systems directly into the building’s exhaust system to reduce heat load and macroenvironmental contamination. Some ventilated micro-isolation cage racks that control the cage supply and exhaust air can selectively maintain the air pressure in the cage positively or negatively relative to the room air pressure. Some of the blower units are now designed to provide humidity-consistent air supply for the cages (especially for dry geographic regions) where there has been increased awareness of inconsistent microenvironmental relative humidity being a study variable that can affect water consumption, general physiology concerns, nociceptive response [19, 20], and other possible parameters (breeding) that are still under study. Individual air supply and exhaust port locations vary by IVC manufacture, and there has been debate about whether there are study effects related to how air has been supplied and at what rates, especially for behavioral mouse studies and anxiety outcomes in mice. One study showed increased anxiety in mice with lower cage air supply [21]. Reproduction rates and pup mortalities [22] have been evaluated, and there appears to be no real significant differences related to where air is supplied. Another study evaluated the impact of change intervals and ventilation rates on the health of breeding pairs and trios of mice [14]. Based on ammonia and carbon dioxide concentrations and physiological parameters (breeding performance, weanling weight and growth, plasma corticosterone levels, immunological function, and select histology), the researchers concluded that cage changes once every 14 days and ventilation rates of 60 Air Changes per Hour (ACH) provided the most optimal conditions for housing. Systems from different manufacturers have been shown to provide different conditions of uniformity and balance of air [23]. Most studied systems minimized ammonia concentration and maintained dry bedding conditions for up to 2 weeks. A study evaluated the possible stress issues in transferring group-housed, long-term telemetered (6 months) Guinea pigs into clean ventilated cages and monitoring for 4 days. There was no change in heart rate, and a transient change in locomotion, with a slight decrease in food consumption [24]. There was an indication that longer in-life studies are needed to fully evaluate chronic potential effects. Since the first IVC caging system developed in 1979, numerous systems with varying types of airflow mechanics have been manufactured. Due to the wide variation of ways these are reported in the literature, all using the term “individually ventilated caging,” an IVC classification system is being proposed to allow better comparisons and functions [25]. Minimally, manufacture and model number as well as airflow rates should be provided. Temperature, Humidity, and Ventilation A significant benefit to using ventilated caging is HEPA-filter air coming from the cages or directly exhausting the cage air from the room, which reduces animal allergens and odors in the environment for both animals and staff [26]. Some filtration of discharged air is desirable to reduce the amount of dust deposited into the exhaust ducts and heat recovery systems, which cause temperature maintenance issues and impact study conductance. If the discharged air is directed into the room exhaust system, odors and potential pheromones generated in the cage are also discharged. Another advantage of ventilated caging is that it allows for high-density housing. Typical mobile, double-sided, ventilated mouse cage racks commonly hold up to 140 cages (7 cages wide × 10 rows high × 2 sides); each cage has the capacity for up to 5 mice. Fixed racks (without wheels) may be stacked even higher, further increasing housing density. The environment is highly regulated for animal cage density but poorly controlled or regulated for caging density and potential animal effects. It is important to report the animal space allocations as well as room caging densities on studies. Many studies have shown that the thermoneutral zones as well as ambient temperatures selected by mice and rats in temperature gradients are considerably higher than those recommended in regulatory guidelines. Rodents must make anatomical, physiological, or behavioral adjustments in order to maintain body temperatures in their thermoneutral zone or preferred range when they are housed at temperatures comfortable for humans (eg, 22°C or standard temperatures). These adjustments are measurable and have been described in studies as consequences of altered ambient temperature. In 1 study [27], cardiovascular parameters were measured with radiotelemetry in mice and rats that were housed in temperature-controlled environments. Small changes in ambient temperature within the range specified in the guide (18–26°C) and slightly higher (30°C) had a significant impact on cardiovascular parameters. As ambient temperature decreased, mean blood pressure, heart rate, and pulse pressure increased significantly for both mice and rats, with mice demonstrating a greater sensitivity to these ambient temperature changes. The use of adequate bedding or nesting materials that allow rodents to burrow into or create nests with the materials has been shown to provide a thermal compensating mechanism to achieve ambient temperatures that approach thermoneutrality [28–30]. These data may provide an alternate explanation to psychological enrichment for rodents’ use of these materials and their preference for them. Similarly, as noted above, the thermal preferences and effective ambient temperatures may differ between single- and group-housed animals, which may also help to explain their preference for social housing [31]. Work done by Hylander [29] has shown differences in rodent cancer models and tumor growth rates due to differences in thermoneutral zone and standard temperature differences. This work cites an underlying physiological mechanism of norepinephrine and epinephrine increase to maintain animal temperature, which in turn stimulates cell proliferation and VEGF and tumor growth as well as metastasis. For more information, see the chapter on macro- and microenvironments. Space Animal cage space is highly regulated, whereas caging room density is not. At present, species-specific guidelines for animal cage space and animal cage sizes are primarily based on the weight of the animals and the number of animals per cage. The 8th version of the Guide for the Care and Use of Laboratory Animals also encourages readers to take the performance indices (age, phenotype, gender, social behavior of different rodent strains, quality of space (eg, vertical access), and structures placed within the cages) into consideration when utilizing these species-specific guidelines. In the past, absence of scientific data to the contrary, many people have anthropomorphized and believed that providing more space is desirable, has no adverse consequences, and would facilitate adding enrichment devices. As noted with animal cage space and the need for determining the optimal cage housing density that supports a physiologically normalized animal (and by extrapolation its well-being) for study conductance would greatly benefit the scientific community. However, given the many variables and responses noted above, a single set of recommendations is unlikely [32]. The main concerns with room caging density is that cages should be accessible, and animals should be able to be visualized. There are similar to animal cage space and caging room density (caging types, cage and room ventilations) where scientific investigation may give a greater understanding of room caging limitations. Currently, cage density is limited by husbandry efficiencies. What is often not published or indicated is room caging density, which has possible effects on study outcomes. Studies have evaluated cage space needs and the effects of social housing, group size, and density and housing conditions for many different species and strains of rodents and have reported varying effects on behavior (such as aggression) and experimental outcomes (ILAR, Guide for the Care and Use of Laboratory Animals) [33]. Study design and experimental variables make comparisons between studies very difficult. Variables include but are not limited to species, strain (and social behavior of the strain), phenotype, age, gender, quality of the space (eg, vertical access), and structures placed in the cage. These issues remain complex and should be carefully considered when housing rodents. Whittaker et al [34] has an excellent review of the effects of space allocation and housing density on measures of laboratory mouse well-being, which includes study data outcome concerns. They conclude that mouse well-being, in regard to space for biological functions, has little effect on reproductive parameters and increasing individual space may actually increase aggressive encounters. Animal density effects on welfare parameters have shown increasing trends of effects on the HPA axis, including a stress response and reduced immune function and increases in aggressive behavior, and may have effects on reproductive parameters particularly in male animals. The diversity of strains used in research and the well-known differing predispositions to aggression among them, strain may be a far greater influence on space deliberations than size, weight, or age of the animal. The varying study designs of the research they reviewed did not allow any conclusions to be drawn on the effects of sex, age, or strain on space requirements and that unfortunately is the same now. Additional well-controlled experiments in this area are needed. Also, male Hsd: Sprague Dawley rat findings suggest that social rank is a significantly greater modifier of affective state than either housing density or space allocation. Dominant animals responded with significantly more optimistic behavioral decisions compared with subordinates for both the housing density and space allocation experiments. This finding suggests that studies should consider animal social status in addition to floor space requirements [35]. Environmental Enrichment Caging composition and functionality have been concerns as a general study data confounder, but far more studies have looked at direct microenvironmental (cage complexity) concerns. The body of information regarding the influence of cage complexities on the mouse has grown considerably, and new findings continue to be published. These findings can generally be categorized into effects on the behavior or biology of the animals, often described in the context of changes in a specific animal model. The value of enrichment for the well-being of rodents is becoming well documented and important for better data outcomes, especially for behavioral-based studies [36, 37] involving mice housed in “nonenriched” cages. They have impaired brain development, stereotypies, and an anxious behavioral profile compared with enriched caged mice. An extensive review by Bayne including neurological effects in mouse models indicates extensive specific morphological changes and altered gene expressions that regulate neuronal structures, synaptic signaling, and brain plasticity in enriched vs nonenriched rodents. Also, rodent models of Rett syndrome, Alzheimer’s, streptozotocin-induced diabetes, and MPTP-induced Parkinson’s that were enriched during the studies resulted in varied experimental outcomes compared with those that were not enriched [38]. Behaviorally a standard mouse cage provides limited scope for the expression of species-appropriate behaviors for the mouse. Allowing the animals control over a complex environment should elicit behavioral changes. This is the actual desired outcome when concerned with abnormal behaviors in unenriched environments. The introduction of 1 or more objects is followed up by monitoring general activity levels (or more specifically, exploration, locomotion, sleep, stress, or anxiety-related behaviors; social, appetitive, and grooming) in some studies. Age at introduction also has an influence in that exposure to enriched environments early in the postweaning period may offset the expression of some abnormal behaviors, such as stereotypy, in animals subsequently housed in more limited environments [39]. Results vary among strains, gender, and type of object(s) introduced. The data focus on increased activities often expressed as an increased exploratory behavior in the home cage; there have been inhibitions of exploration in experimental settings, with some gender differences, such as an open-field test or elevated T-maze [40, 41]. There also has been a trend of decreased cancer incidence with environmental enrichment, which appears to be linked to downregulation of leptin in various tissues (mitochondria, adipocytes, serum, breast, pancreatic) and is mostly proposed through the upregulation of increased brain derived neurotropic factor in the hypothalamus with cage enrichment [42, 43]. This offers new opportunities in research and therapy for cancer [38]. Ways to increase the complexity of the primary environment for the animals are currently being performed and expected at most research facilities. The challenge is to balance seemingly conflicting requirements to provide for animal well-being through cage enrichment with features required to provide for routine animal care and cage sanitation while assuring animal health and achieving the research goals successfully. Some of the greatest success in this has been with using the cage enrichment as part of the study especially with cognitive components, which is mostly occurring in NHP species [43, 44], and the use of tablets for interactions. Commercial window-enhanced tablets that are encased in protective plastic and mounted on the cage are available, where animals can choose video, music, and lighting, or express themselves with finger paints. An optional remote device panel allows the NHP to remotely operate a set of lights and/or other devices. Technicians can control and add features as the animals learn the applications. Other examples [45] include a smart chair designed to automatically train macaques to enter an NHP chair for water reward without a human in the animal room, and a helmet-mounted eye camera apparatus allowing telemetric infrared video gaze tracking in 2 ring-tailed lemurs (Lemur catta) while pair housed and navigating their naturalistic home pen. Rodent automated home cage systems are also incorporating more “in-cage” devices where behavioral testing is being conducted in the home cage [46, 47]. In large animal holding areas, spaces for animals to hide or seclude from others have become more standard in design, where these items have been added into the cage for rodents but have been questioned if the addition may in fact cause more fighting due to the hiding and rank reorders needed from the hiding. Van Loo [48] determined that the type of enrichment strongly affected both aggressive behavior as well as a variety of physiological parameters in male BALB/c mice. The addition of a rigid shelter in the cage elevated stress indicators—increased intermale aggression, increased urine corticosterone/creatinine ratios, and decreased weight gain. Providing nesting material decreased intermale aggression, but urine/corticosterone/creatinine ratios did not change from the control group. The ability of the mice to control their environment and allow normal nesting behavior was credited for this change. It was also noted that the quality of space (vertical complexity) was increased as was the quantity of available space with the rigid shelter. Alternative data indicated enriched weaned male and female C57BL/6 mice housed 6 to a cage in enriched large cages showed signs of increased anxiety with a plus maze test compared with similar groups in smaller unenriched cages [41]. A difference was the maze study was conducted in the dark phase of the light cycle whereas other studies had been conducted during the light phase of the light cycle. Other studies [49] used C57BL/6 female mice to evaluate how an enriched environment would affect the immune system. It was demonstrated that cage toys buffered the reactivity of the immune system in response to distress, and long-term environmental enrichment lowered the murine splenic proliferative response to acute stress compared with nonenriched mice. The references cited above help to underscore that although environmental enrichment for mice in general appears to have a positive impact on the animals, it can create a scientific variable that has not been well defined to date. It is important to have a good understanding of the term enrichment and that it should provide “beneficial” enrichment to the animals, distinguishing this as a species-relevant approach that improves welfare rather than simply putting any item into a cage and referring to it as enrichment [38]. After looking at multiple laboratories assessing the effect of enrichment on variation in behavioral endpoints and reproducibility of behavioral differences in 3 strains of mice, it has been determined that within-group variability contributed an average of 60% of total variability and was unaffected by enrichment. Thus, nonenriched cage environments fail to reduce individual variability in behavioral endpoints [50]. The environmental complexity chapter in this journal has a very thorough review of this area. Cage and Equipment Sanitation Practices Routine cage changing is important for maintaining the health and hygiene of animals. This husbandry procedure is associated with several behavioral and physiologic effects in rodents that can alter metabolism and study data results. There is growing interest to not only understand the effects and repercussions of animal responses to cage changes but also to reduce these responses. With the introduction of automated home cage systems, there is a developing knowledge base for the length of activity disruption and associated factors that contribute to cage changes [51, 52]. These reports have indicated that there is a minimal 48 to 96 hours disruption of normal cage activity (primarily daytime) dependent on room size and procedures, strain, sex, and time of day of cage changing. Importantly, age-dependent changes were not seen. Comparisons of cage changing conducted later during the day (12:00) or near the beginning of the dark cycle (17:00) altered the patterned responses but did not reduce night time motion for either sex, making this alternative not as attractive. Study behavioral and metabolic data that are collected post cage change may have inadvertent data collection and interpretation effects. Another study indicated that disruption predictability (cage changing) was not stressful [53], but the authors also indicated that the disruption period(s) were not long enough to induce chronic stress. Study reports should include cage change outs and relationship to study data collections. Many different procedures of caging and equipment sanitization are practiced in facilities. The initial step is handling the cages and putting animals into sanitized clean cages. Human-associated factors of study disruptions in this area historically were related to the wearing of fragrances, which has been addressed in regulatory guidelines. A recent report indicating staff gender differences [54] has indicated male research staff and other male animals have a significant stress influence on behavioral study outcomes related to olfactory stimulation of stress hormones. This report focused on research staff who had intermittent contact with the study animals, and it went on to indicate this same concern was noted with other male animals (excluding male cage mates). The exposure to shirts worn overnight by men, bedding material from sexually intact and unfamiliar male mammals, and the presentation of compounds secreted from the human axilla also reproduced this effect. It has not been shown that there is the same effect with care staff interactions, but it would seem likely. These findings encourage researchers to report the gender of experimenters/care staff in their publications, and if the experimenters/care staff change mid-stream to include their genders as a variables in the analysis. The adoption of statistical methods that compensate for a greater range of variability is supportive for the appropriate analysis of the data. The periods of cage changing are variable and as such the indicators for cage change (calendar based with criteria limits, or criteria based with calendar time limits) if there is partial cage change and/or nest removals done at the time of cage changing all have impacts on the study outcomes and should be indicated in the study descriptions. Animal handling for movement to behavioral study trials has been shown to affect anxiety in animals that causes variability in study outcomes. The non-adverse handling tunnel method (that has been properly trained and utilized) in high-anxiety male mice significantly improved performance reliability in habituation-dishabituation behavioral testing [55]. The study handling methods of the mice, especially in behavioral work, should be reported. Different chemicals are used in cleaning processes. Cage residues post rinsing are of the greatest concern in the use of these products due to study interference. There is sparse documentation of this subject in the literature. Agents designed to mask animal odors may have volatile compounds and as such should not be used in animal housing facilities due to alterations of basic physiological and metabolic processes. Monitoring of sanitation practices should fit the process and materials being cleaned and may include visual inspection and microbiologic and water temperature monitoring. As indicated earlier, barriers usually employ the use of autoclaves to effectively eliminate unwanted adventitious agents. Repeated cage washing and autoclaving has raised concerns regarding bisphenol A even with newer polysulfone cages once they have become damaged [4], which has caused study reproducibility issues [56, 57]. Autoclave cycle type and times should also be included in study descriptions as appropriate. The greater risk for barrier areas is food and bedding, which should either be autoclaved or irradiated. Parvo virus concerns and barrier breaks have indicated that using irradiation for food sterilization has been very effective [58]. Use of irradiated feed and bedding eliminates the need for autoclaving them into the barrier. Barrier facilities are designed and managed at various levels of microbiological control, affecting the degree of control over the ways in which supplies and personnel enter the facility. The barrier facilities may have 1 or more double-door, pass-through autoclaves and 1 or more ventilated entry and exit vestibules with interlocking doors. Packaged sterile supplies and animals in filtered containers can be passed into the barrier through these vestibules after having the exterior surface of the package chemically sanitized. In some circumstances, a pass-through dip tank, filled with high-level disinfectants, may be used to pass sterile items packaged in watertight containers into the barrier. As with equipment and supplies, a personnel barrier entry is variable. Water showers, air showers, or a vestibule with interlocking doors are some of the variable procedural means of access. Again, barrier types and procedural processes should be included in report/manuscript write-ups. Substrate Types and Effects Bedding is used to absorb moisture, minimize the growth of microorganisms, and dilute and limit animals’ contact with excreta. No type of bedding is ideal for all species under all management and experimental conditions. Untreated softwood shavings can affect metabolism [59, 60] and should be avoided. Cedar shavings emit aromatic hydrocarbons that induce hepatic microsomal enzymes and cytotoxicity [61–63] and have been reported to increase the incidence of cancer [64, 65]. Bedding can also influence mucosal immunity [66] and endocytosis [67]. Corncob bedding (primary or mixed) indicated in study disruptions related to phytoestrogen effects [68], feed conversion efficiency in prediabetic mouse model [69], and effects on slow wave sleep [70] are examples of study data effects. Zahorsky-Reeves et al [71] reported that changing to the use of paper-based (vs hardwood) bedding decreased the incidence of blepharitis in a population of nude rats. In nude or hairless mice that lack eyelashes, paper bedding with fines can cause abscesses [72]. Species-specific behaviors and thermoregulation are encouraged and exhibited with appropriate bedding and nesting materials [32]. Nesting materials provision is usually considered an enrichment, but it has become an integral part of the substrate for reasons mentioned earlier. Not all studies should be on nestlets or certain types of nestlets. As an example, tissue nesting material can be a confounding variable for studies of allergic asthma in BALB/c mice, resulting in increased total cell number, eosinophil number, and IL-13 concentration in bronchoalveolar lavage fluid compared with nonenriched control animals [73]. Cotton nestlets can cause conjunctivitis [74], and some types of nesting material can entangle the limbs of pups [75]. More recently, the provision of a nest-building material was considered an important element of the mouse’s cage environment. The manifestation of nest-building behavior by mice is considered a reliable indicator of health and welfare, with its absence reflecting pain or discomfort in the mice. The cancer field has been exploring how environmental enrichment negatively impacts several types of cancer growth (adipocytes, breast, pancreatic, murine, and transplanted human glioma cells) related mostly to upregulation of hypothalamic brain derived neurotropic factor in environmentally enriched mouse studies and then it is downregulation effect on leptin levels, mitochondrial genes, or elevated interleukin-15 [38]. The scientific evidence overwhelmingly supports enrichment to improve the welfare of mice used in research. However, the type of enrichment must be biologically relevant, safe for the animal, improve the animal’s welfare, and not interfere with the scientific measures taken from the animals. When these criteria are met, the data produced by the animal will be more valid and reliable. Interestingly, the use of environmental enrichment has led to new areas of scientific enquiry [38]. Unlike mice, adult laboratory rats do not spontaneously exhibit nest-building behavior when nesting material is offered. Van Loo and Baumans [76] have shown that nest building in rats is an acquired behavior. When rats were provided nesting material from birth, they learned to utilize the materials, as has been observed also in wild rats. Similarly, rats have been shown to prefer the type of bedding on which they were raised [77]. The choice of bedding or nesting material may also have a significant impact on physiological parameters and therefore on experimental results. Noise vibration and lighting concerns are each covered in detail in other chapters in this journal. Water Quality and Supply Water supply means are varied throughout the industry and are covered in detail in another chapter in this journal (Water Quality and Stability). Water quality is usually part of a QA program with appropriate periodic monitoring for pH, hardness, and microbial or chemical contamination. Given localities will influence the results and understanding of the primary influences, and those changes can be a trigger for when to monitor. Water can be treated or purified to minimize or eliminate contamination when protocols require highly purified water. Reverse osmosis water is commonly used at most facilities. The question of acidification, chlorination, or no water treatment is dependent on the facility and should be guided by its water quality findings. The selection of water treatments should be carefully considered because many forms of water treatment have the potential to cause physiologic alterations, reduction in water consumption, changes in microflora, or effects on experimental results [78–80]. In a report with water acidified with either hydrochloric or sulfuric acid to pH levels of 2.0 or 2.5 given for 6 weeks to normal and immuno-suppressed male, random-bred mice, the pH of the drinking water remained stable for periods up to 7 days. When water was acidified to a pH of 2.0, all mice exhibited decreased water consumption and weight declines, with immunosuppressed animals more affected. Also noted was the number of bacterial species isolated from the terminal ileum was reduced [81]. Using conditional adenomatous polyposis coli (Apc) knockout mouse models, a chromosomal knockout Apc line was given chlorinated drinking water (10.0 mg/L chlorine) and tap water (0.6 mg/L chlorine). In Apc mice given chlorinated drinking water, tumors tended to develop in the colon, whereas in those that drank tap water, tumors were mostly observed in the small intestine. Chlorinated water altered the enteric environment by reducing several fecal populations of the obligatory anaerobes, including species belonging to the Atopobium cluster (Enterobacteriaceae and Staphylococcus sp.), which was associated with colon tumorigenesis in CPC;Apc mice [82]. Water supply and treatments need to be clearly defined in study reports. Developing Areas of Specialized Equipment Isolators The past 2 decades have seen an explosion of research investigating the role of microbiota in health and disease. A major component of this drive has been related to the National Institutes of Health-acknowledged issues of poor reproducibility in biomedical research, particularly in preclinical studies using animal models [1]. Earlier in this chapter, several areas to improve reproducibility were discussed, primarily focusing on the thorough documentation of all husbandry variables and associated equipment used. Another publicized area suggests the need for reporting, and possibly standardizing, the gut microbiota of research animals [83]. This area is developing rapidly and has great merit, especially with the development of fecal banks. Isolators have been a main-stay in maintaining germ-free or gnotobiotic animals that are essential in this microbiome research. There are 2 basic types: flexible film isolators and semi-rigid wall isolators. Currently, most flexible film isolators are made from transparent poly-vinyl-chloride flexible plastic sheeting supported by a metal or plastic frame and positive pressure air, and semi-rigid isolators from polycarbonate and poly-vinyl-chloride flexible plastic sheeting. HEPA-filtered air is supplied, thus minimizing the risk of contaminating animals with infectious agents from outside the isolator. Personnel work inside the isolators through portholes fitted with sleeves and gloves made of latex or other similar material. Well-established systems involving equipment, ports, and procedures are used for moving sterile materials into the isolator and soiled materials and trash out of the isolator. Many of the issues that were mentioned for IVC caging have similar issues with the isolators (either flexible film or semi-rigid) in that air rates are variable for the units and there are inherent vibration and noise concerns with design of some of the units that can impact breeding especially of sensitive transgenic lines. In addition to traditional isolators, which generally have a large foot print related to holding animal holding capacity and are primarily used to maintain breeding animals, there has been an increased use of sealed positive-pressure IVC caging systems to allow expanded research studies of smaller cohorts of animals more efficiently for shorter periods of study times [84]. The type of isolators, settings that the isolators are maintained at, and methods of how they are maintained should be reported in study methods to allow better study replications [85]. Automated Home Cage Systems The NIH’s continued acknowledgment of the issues of poor reproducibility and translation has indicated there is great experimental variability in studies regarding differences in animal caging, substrates used, husbandry practices, and animal handling procedures as mentioned earlier in this chapter. Many variables can influence animal behavior and physiology, potentially affecting scientific study outcomes. Laboratory and husbandry procedures—including handling, cage cleaning, injections, blood collection, and animal identification—can produce a multitude of effects. Previous ways to study the effects of these procedures was by making behavioral and physiologic measurements at specific time points. This approach can be disruptive and limits the frequency or duration of observations. Because these procedures can have both acute and long-term effects, the behavior and physiology of animals would best be monitored continuously. What are considered incidental mandatory procedures such as cage changes may seem to be innocuous due to control animal comparisons but may indirectly have impacts on study data related to types of studies being conducted [52]. The ability to gather longitudinal data on a large sample population of animals would allow researchers to explore how these environmental manipulations interact with age and genetics and to understand the magnitude and duration of procedure-related effects. There are several types of automated home cage systems [86], with the majority using video systems and/or Radio Frequency Identification (RFID) for capture of animal activity, but other modalities are used as well (electromagnetic field (EMF) disruptions, vibration, infrared beam breaks). Many of the systems require animals to be moved into customized environments (not a standard housing cage or major alterations to a standard housing cage [altered wire bar lid and top]) and acclimated and thus are significantly altered from the home cage environment. Other systems utilize the standard housing cage (currently Actual Analytics, CleverSys Inc, Tecniplast DVC, and Vium) and augment the rack to allow animals to remain in a standard housing cage, which by-passes acclimation concerns for the general rodent population and can allow greater scale-up for study purposes. As such, true home cage has been defined as a normal rack-mounted cage where mice are born, reared, and housed within their established social groups [86]. Some of the systems only allow for single mouse evaluations (especially with side camera modalities or beam break type of systems). Removing the mouse from its cage-mates and placing it into a novel environment has been shown to affect behavior, general well-being, and metabolism [87]. Social behaviors and the ability to have social interactions are critical in display and development of normal behaviors. Paired social interactions using automated visual tracking systems are most commonly studied in laboratory environments. However, complex group social behaviors are often harder to quantify; the current challenge lies in the ability of visual systems to distinguish between animals and patterns when the mice are close to each other and in the presence of enrichment and nesting materials [88]. The use of top-down cameras allows better animal tracking and changes in bedding but has some loss in granularity, making detection of fine motor movements like grooming not detectable [89, 90]. Many of these systems can also be multiplexed with additional equipment to allow additional data capture, tracking of multiple animals, and quantification of complex social interactions free from experimenter bias. Some of these caging systems have been evaluated to determine the influences of the stand-alone system on animal behavioral and/or pathology outcomes and have not shown differences [91, 92]. This should be a critical aspect of system incorporation especially with the newer technologies (RFID and EMF as examples). In addition to animal activity, behavioral analysis, and metabolism measures, some systems also evaluate the cage conditions, allowing a standard automated evaluation of soiled caging conditions and more consistent caging bedding conditions across different animal housing densities. The primary use in all of these systems is to greatly improve study data collection by maintaining animals in the home cages and thus avoid excessive laboratory and husbandry procedures—including handling, cage movement and cleaning, animal identification, and personnel entry into the housing room—that then produce a multitude of effects on animal behavior and physiology. These effects include changes in activity patterns, stress and anxiety levels, heart rate, blood pressure, and body temperatures [9]. As with other caging systems and per the ARRIVE Guidelines, the type of automated home cage system, how they are used, and how data are derived should be reported in study methods to allow better study replications [93, 94]. Conclusion In conclusion, rodent housing systems and corresponding husbandry practices have known and potential confounders in the research environment. These practices are best documented to full extent in the methods sections of study papers allowing full understanding by other research groups of the extent and nature of study conduct and the better opportunity for study and data replications and comparisons. This area has a very high impact for reproducibility and comparability of study data outcomes. References 1. Franklin CL , Ericsson AC . Microbiota and reproducibility of rodent models . Lab Anim (NY). 2017 ; 46 ( 4 ): 114 – 22 . doi: 10.1038/laban.1222. Google Scholar Crossref Search ADS PubMed WorldCat 2. Weldon L , Abolins S , Lenzi L , et al. The gut microbiota of wild mice . PLoS One. 2015 ; 10 ( 8 ). doi: 10.30802/AALAS-CM-18-000049. OpenURL Placeholder Text WorldCat 3. Masopust D , Sivula CP , Jameson SC . Of mice, dirty mice, and men: using mice to understand human immunology . J Immunol. 2017 ; 199 ( 2 ): 383 – 8 . doi: 10.4049/jimmunol.1700453. Google Scholar Crossref Search ADS PubMed WorldCat 4. Gorence GJ , Pulcastro HC , Lawson CA , et al. Chemical contaminants from plastics in the animal environment . J Am Assoc Lab Anim Sci. 2018 , 2019 ; 58 ( 2 ): 190 – 6 . doi: 10.30802/AALAS-JAALAS-18-000074 [Epub 2019 Jan 15] . Google Scholar Crossref Search ADS WorldCat 5. Reynolds RP , Kinard WL , Degraff JJ , et al. Noise in a laboratory animal facility from the human and mouse perspective . J Am Assoc Lab Anim Sci. 2010 ; 49 : 592 – 7 . Google Scholar PubMed OpenURL Placeholder Text WorldCat 6. Reynolds RP , Li Y , Garner A , et al. Vibration in mice: a review of comparative effects and use in translational research . Animal Model Exp Med. 2018 ; 1 ( 2 ): 116 – 24 . doi: 10.1002/ame2.12024. Google Scholar Crossref Search ADS PubMed WorldCat 7. Campagna MV , Faure-Kumar E , Treger JA , et al. Factors in the selection of surface disinfectants for use in a laboratory animal setting . J Am Assoc Lab Anim Sci. 2016 Mar ; 55 ( 2 ): 175 – 88 . OpenURL Placeholder Text WorldCat 8. Hershey JD , Gifford JJ , Zizza L , et al. The effect of various cleaning agents on performance of mice in common behavioral assays of anxiety . J Am Assoc Lab Anim Sci. 2018 ; 57 ( 4 ): 335 – 9 . Google Scholar Crossref Search ADS PubMed WorldCat 9. Otto G , Tolwani RJ . Use of microisolator caging in a risk-based mouse import and quarantine program: a retrospective study . Contemporary Topics in Laboratory Animal Science. 2002 ; 41 : 20 – 7 . Google Scholar PubMed OpenURL Placeholder Text WorldCat 10. Lipman NS . Microenvironmental conditions in isolator cages: an important research variable . Laboratory Animal Science. 1992 ; 21 : 23 – 7 . OpenURL Placeholder Text WorldCat 11. Lipman NS . Rodent facilities and caging systems. In Hessler JR , Lehner NDM , eds. Planning and Designing Research Animal Facilities. San Diego : CA: Elsevier Inc ; 2009 . p. 265 – 88 . Google Scholar Google Preview OpenURL Placeholder Text WorldCat COPAC 12. Corning BF , Lipman NS . 1991. A comparison of rodent caging systems based on microenvironmental parameters . Laboratory Animal Science. 1991 ; 41 : 498 – 503 . Google Scholar PubMed OpenURL Placeholder Text WorldCat 13. Hasenau JJ , Baggs RB , Kraus AL . Microenvironments in microisolation cages using BALB/c and CD-1 mice . Contemporary Topics in Laboratory Animal Science. 1993 ; 32 : 11 – 6 . OpenURL Placeholder Text WorldCat 14. Reeb-Whitaker CK , Paigen B , Beamer WG , et al. The impact of reduced frequency of cage changes on the health of mice housed in ventilated cages . Lab Anim. 2001 ; 35 : 58 – 73 . Google Scholar Crossref Search ADS PubMed WorldCat 15. Rosenbaum MD , VandeWoude S , Johnson TE . Effects of cage-change frequency and bedding volume on mice and their microenvironment . J Am Assoc Lab Anim Sci. 2009 Nov ; 48 ( 6 ): 763 – 73 . OpenURL Placeholder Text WorldCat 16. Lipman NS . Isolator rodent caging systems (state of the art): a critical view . Contemporary Topics in Laboratory Animal Science. 1999 ; 38 : 1 – 17 . OpenURL Placeholder Text WorldCat 17. Baumans V , Schlingmann F , Vonck M , et al. Individually ventilated cages: beneficial for mice and men? Contemp Top Lab Anim Sci. 2002 ; 41 : 13 – 9 . Google Scholar PubMed OpenURL Placeholder Text WorldCat 18. Krohn TC , Hansen AK , Dragsted N . 2003. The impact of cage ventilation on rats housed in IVC systems . Lab Anim. 2003 ; 37 : 85 – 93 . Google Scholar Crossref Search ADS PubMed WorldCat 19. Nicolaus ML , Bergdall VK , Davis IC , et al. Effect of ventilated caging on water intake and loss in 4 strains of laboratory mice . J Amer Assoc Lab Anim Sci. 2016 ; 55 ( 5 ): 525 – 33 . OpenURL Placeholder Text WorldCat 20. Chesler EJ , Wilson SG , Lariviere WR , et al. Identification and ranking of genetic and laboratory environment factors influencing a behavioral trait, thermal nociception, via computational analysis of a large data archive . Neurosci Biobehav Rev. 2002 Dec ; 26 ( 8 ): 907 – 23 . Google Scholar Crossref Search ADS WorldCat 21. Burman O , Buccarello L , Redaelli V , et al. The effect of two different individually ventilated cage systems on anxiety-related behaviour and welfare in two strains of laboratory mouse . Physiology and Behavior. 2014 ; 124 : 92 – 9 . Google Scholar Crossref Search ADS WorldCat 22. Marsella G , Zennaro E , Grignaschi G . Reduced air changes per hour in IVCs and the impact on breeding performance . Animal Technology and Welfare. 2012 ; 10 ( 1 ): 7 – 10 . OpenURL Placeholder Text WorldCat 23. Hoglund AU , Renstrom A . Evaluation of individually ventilated cage systems for laboratory rodents: cage environment and animal health aspects . Lab Anim. 2001 ; 35 : 51 – 7 . Google Scholar Crossref Search ADS PubMed WorldCat 24. Giral M , Armengol C , Sánchez-Gómez S . Gavaldà A. Effects of changing to individually ventilated caging on Guinea pigs (Cavia porcellus) . J Am Assoc Lab Anim Sci. 2015 ; 54 ( 3 ): 267 – 72 . Google Scholar PubMed OpenURL Placeholder Text WorldCat 25. Perkins S , Lipman NS . Properly describing individually ventilated caging systems in scientific manuscripts . J Am Assoc Lab Sci. 56 ( 5 ): 488 . OpenURL Placeholder Text WorldCat 26. Thulin H , Bjorkdahl M , Karlsson , Renstrom A-S, A . Reduction of exposure to laboratory animal allergens in a research laboratory . Ann Occup Hyg. 2002 ; 46 : 61 – 8 . Google Scholar PubMed OpenURL Placeholder Text WorldCat 27. Swoap SJ , Overton JM , Garber G . Effect of ambient temperature on cardiovascular parameters in rats and mice: a comparative approach . Am J Physiol Reg Int Comp Physiol. 2004 ; 287 : R391 – 6 . Google Scholar Crossref Search ADS WorldCat 28. Gordon CJ . Effect of cage bedding on temperature regulation and metabolism of group-housed female mice . Comp Med. 2004 ; 54 : 63 – 8 . Google Scholar PubMed OpenURL Placeholder Text WorldCat 29. Hylander B , Repasky E . Thermoneutrality, mice, and cancer: a heated opinion . Trends in Cancer. 2016 ; 2 ( 4 ): 166 – 75 . doi: 10.1016/j.trecan.2016.03.005. Google Scholar Crossref Search ADS PubMed WorldCat 30. Hankenson CF , Marx JO , Gordon CJ , et al. Effects of rodent thermoregulation on animal models in the research environment . Comp Med. 68 ( 6 ): 2018 , 425 – 438 (14) doi: 10.30802/AALAS-CM-18-000049 . OpenURL Placeholder Text WorldCat 31. Gordon CJ , Becker P , Ali JS . Behavioral thermoregulatory responses of single- and group housed mice . Physiol Behav. 1998 ; 65 : 255 – 62 . Google Scholar Crossref Search ADS PubMed WorldCat 32. Gonder JC , Laber K . A renewed look at laboratory rodent housing and management . ILAR J. 2007 ; 48 ( 1 ): 29 – 36 doi: 10.1093/ilar.48.1.29 . Google Scholar Crossref Search ADS PubMed WorldCat 33. ILAR (Institute for Laboratory Animal Research), Committee for the Update of the Guide for the Care and Use of Laboratory Animals. Guide for the Care and Use of Laboratory Animals. 8th ed . Washington, DC : National Research Council, National Academies Press ; 2011 . Google Scholar Google Preview OpenURL Placeholder Text WorldCat COPAC 34. Whittaker AL , Howarth GS , Hickman DL . Effects of space allocation and housing density on measures of wellbeing in laboratory mice: a review . Lab Anim. 2012 ; 46 : 3 – 13 . doi: 10.1258/la.2011.011049. Google Scholar Crossref Search ADS PubMed WorldCat 35. Barker TH , George RP , Howarth GS , et al. Assessment of housing density, space allocation and social hierarchy of laboratory rats on behavioral measures of welfare . PLoS ONE. 2017 ; 12 ( 9 ): e0185135. doi: 10.1371/journal.pone.0185135. OpenURL Placeholder Text WorldCat 36. Van Praag H , Kempermann G , Gage FH . Neural consequences of environmental enrichment . Nature Rev Neurosci. 2000 ; 1 : 191 – 8 . Google Scholar Crossref Search ADS WorldCat 37. Wurbel H . Ideal homes? Housing effects on rodent brain and behavior . Trends Neurosci. 2001 ; 24 : 207 – 12 . Google Scholar Crossref Search ADS PubMed WorldCat 38. Bayne K . Environmental enrichment and mouse models: current perspectives . Animal Models and Experimental Medicine. 2018 ; 1 ( 2 ): 82 – 90 doi: 10.1002/ame2.12015 . Google Scholar Crossref Search ADS PubMed WorldCat 39. Marques JM , Olsson IAS . The effect of preweaning and postweaning housing on the behavior of the laboratory mouse (Mus musculus) . Lab Anim. 2007 ; 41 : 92 – 102 . Google Scholar Crossref Search ADS PubMed WorldCat 40. Lin E-J , Choi E , Liu X , et al. Environmental enrichment exerts sex-specific effects on emotionality in C57BL/6J mice . Behav Brain Res. 2011 ; 216 : 349 – 57 . Google Scholar Crossref Search ADS PubMed WorldCat 41. Zhu SW , Yee BK , Nyffeler M , et al. Influence of differential housing on emotional behavior and neurotrophin levels in mice . Behav Brain Res. 2006 ; 169 : 10 – 20 . Google Scholar Crossref Search ADS PubMed WorldCat 42. Cao L , Liu X , Lin E-JD , et al. Environmental and genetic activation of a brain-adipocyte BDNF/leptin axis causes cancer remission and inhibition . Cell. 2010 ; 142 : 52 – 64 . Google Scholar Crossref Search ADS PubMed WorldCat 43. Calapai A , Berger M , Niessing M , et al. A cage-based training, cognitive testing and enrichment system optimized for rhesus macaques in neuroscience research . Behav Res Methods. 2017 ; 49 ( 1 ): 35 – 45 Published online 2016 Feb 19. doi: 10.3758/s13428-016-0707-3 . Google Scholar Crossref Search ADS PubMed WorldCat 44. Coleman K , Novak MA . Environmental enrichment in the 21st century . ILAR J. 2017 Dec 1 ; 58 ( 2 ): 295 – 307 . doi: 10.1093/ilar/ilx008. Google Scholar Crossref Search ADS WorldCat 45. Wyatt JD . Large (nonagricultural) animal enclosures and housing. In Weichbrod RH , T hompson GAH , Norton JN , eds. Management of Animal Care and Use Programs in Research, Education, and Testing. 2nd edition. Boca Raton, FL: CRC Press/Taylor & Francis ; 2018 . Chapter 22, pp 16. doi: 10.1201/9781315152189-22. 46. Winter Y , Schaefers ATU . A sorting system with automated gates permits individual operant experiments with mice from a social home cage . J Neurosci Methods. 2011 ; 196 : 276 – 80 . Google Scholar Crossref Search ADS PubMed WorldCat 47. Bains RS , Wells S , Sillito RR , et al. Assessing mouse behavior throughout the light/dark cycle using automated in-cage analysis tools . J Neurosci Methods. 2018 ; 300 : 37 – 47 . doi: 10.1016/j.jneumeth.2017.04.014. Google Scholar Crossref Search ADS PubMed WorldCat 48. Van Loo PL , Mol JA , Koolhaas JM , et al. Modulation of aggression in male mice: influence of group size and cage size . Physiol Behav. 2001 ; 72 : 675 – 83 . Google Scholar Crossref Search ADS PubMed WorldCat 49. Kingston SG , Hoffman-Goetz L . Effect of environmental enrichment and housing density on immune system reactivity to acute exercise stress . Physiol Behav. 1996 ; 60 : 145 – 50 . Google Scholar Crossref Search ADS PubMed WorldCat 50. Würbel H . Environmental enrichment does not disrupt standardization of animal experiments . Altex. 2007 ; 24 : 70 – 3 . Google Scholar PubMed OpenURL Placeholder Text WorldCat 51. Pernold K , Lannello F , Low BE , et al. Towards large scale automated cage monitoring—diurnal rhythm and impact of interventions on in-cage activity of C57BL/6j mice recorded 24/7 with a non-disrupting capacitive -based technique . PLoS ONE. Feb 2019 . doi: 10.1371/journal/pone.0211063 . OpenURL Placeholder Text WorldCat 52. Schoevitz LR , et al. Retrospective analysis of the effects of identification procedures and cage changing by using data from automated, continuous monitoring . J Am Assoc Lab Anim Sci. 2019 ; 58 ( 2 ): 126 – 41 . Google Scholar Crossref Search ADS PubMed WorldCat 53. Robinson-Junker A , O’Hara B , Durkes A , et al. Sleeping through anything: the effects of unpredictable disruptions on mouse sleep, healing, and affect . PLoS ONE. 2019 ; 14 ( 1 ): e0210620. doi: 10.1371/journal.pone.0210620 . OpenURL Placeholder Text WorldCat 54. Mogil JS , Sorge RE , Martin LJ . Olfactory exposure to males, including men, causes stress and related analgesia in rodents . Nature Methods 2014 ; 11 : 629 – 32 . doi: 10.1038/nmeth.2935 . Google Scholar Crossref Search ADS PubMed WorldCat 55. Gouveia K , Hurst JL . Optimizing reliability of mouse performance in behavioural testing: the major role of non-aversive handling . Sci Rep. 2017 ; 7 : 44999. doi: 10.1038/srep44999 . Google Scholar Crossref Search ADS PubMed WorldCat 56. Howdeshell KL , Peterman PH , Judy BM , et al. Welshons WV. Bisphenol A is released from used polycarbonate animal cages into water at room temperature . Environ Health Perspect 2003 ; 111 : 1180 – 7 . doi: 10.1289/ehp.5993 . Google Scholar Crossref Search ADS PubMed WorldCat 57. Hunt PA , Koehler KE , Susiarjo M , et al. Bisphenol a exposure causes meiotic aneuploidy in the female mouse . Curr Biol. 2003 ; 13 : 546 – 53 . doi: 10.1016/S0960-9822(03)00189-1 . Google Scholar Crossref Search ADS PubMed WorldCat 58. Reuter J , Livingston R , LeBlanc M . Management strategies for controlling endemic and seasonal mouse parvovirus infection in a barrier facility . Lab Anim. 2011 ; 40 ( 5 ): 145 – 52 . doi: 10.1038/laban0511-145. Google Scholar Crossref Search ADS WorldCat 59. Vesell ES . Induction of drug-metabolizing enzymes in liver microsomes of mice and rats by softwood bedding . Science. 1967 ; 157 : 1057 – 8 . Google Scholar Crossref Search ADS PubMed WorldCat 60. Vesell ES , Lang CM , White WJ , et al. Environmental and genetic factors affecting response of laboratory animals to drugs . Fed Proc. 1976 ; 35 : 1125 – 32 . Google Scholar PubMed OpenURL Placeholder Text WorldCat 61. Torronen R , Pelkonen K , Karenlampi S . Enzyme-inducing and cytotoxic effects of woodbased materials used as bedding for laboratory animals: comparison by a cell culture study . Life Sci. 1989 ; 45 : 559 – 65 . Google Scholar Crossref Search ADS PubMed WorldCat 62. Weichbrod RH , Hall JE , Simmonds RC , et al. Selecting bedding material . Lab Anim Sci. 1986 ; 15 : 25 – 9 . OpenURL Placeholder Text WorldCat 63. Weichbrod RH , Cisar CF , Miller JG , et al. Effects of cage beddings on microsomal oxidative enzymes in rat liver . Lab Anim Sci. 1988 ; 38 : 296 – 8 . Google Scholar PubMed OpenURL Placeholder Text WorldCat 64. Jacobs BB , Dieter DK . Spontaneous hepatomas in mice inbred from ha:ICR Swiss stock: Effects of sex, cedar shavings in bedding, and immunization with fetal liver or hepatoma cells . J Natl Cancer Inst. 1978 ; 61 : 1531 – 4 . Google Scholar PubMed OpenURL Placeholder Text WorldCat 65. Vlahakis G . Possible carcinogenic effects of cedar shavings in bedding of C3H-AvyfB mice . J Natl Cancer Inst. 1977 ; 58 : 149 – 50 . Google Scholar Crossref Search ADS PubMed WorldCat 66. Sanford AN , Clark SE , Talham G , et al. Influence of bedding type on mucosal immune responses . Comp Med. 2002 ; 52 ( 5 ): 429 – 32 . Google Scholar PubMed OpenURL Placeholder Text WorldCat 67. Buddaraju AKV , Van Dyke RW . Effect of animal bedding on rat liver endosome acidification . Comp Med. 2003 ; 53 : 616 – 21 . Google Scholar PubMed OpenURL Placeholder Text WorldCat 68. Landeros RV , Morisseau C , Yoo HJ , et al. Corncob bedding alters the effects of estrogens on aggressive behavior and reduces estrogen receptor-α expression in the brain . Endocrinology. 2012 ; 153 ( 2 ): 949 – 53 . doi: 10.1210/en.2011-1745 . Google Scholar Crossref Search ADS PubMed WorldCat 69. Ambery AG , Tackett L , Penque BA , et al. Effect of corncob bedding on feed conversion efficiency in a high-fat diet-induced prediabetic model in C57Bl/6J mice . J Am Assoc Lab Anim Sci. 2014 ; 53 ( 5 ): 449 – 51 . Google Scholar PubMed OpenURL Placeholder Text WorldCat 70. Leys LJ , McGaraughty S , Radek RJ . Rats housed on corncob bedding show less slow-wave sleep . J Am Assoc Lab Anim Sci. 2012 ; 51 ( 6 ): 764 – 8 . Google Scholar PubMed OpenURL Placeholder Text WorldCat 71. Zahorsky-Reeves JL , Schimmel AA , Suckow CE , et al. Paper-based bedding dramatically decreased the incidence of blepharitis in a nude rat population . Contemp Top Lab Anim Sci. 2005 ; 44 : 62 . Google Scholar PubMed OpenURL Placeholder Text WorldCat 72. White WJ , Hawk CT , Vasbinder MA . 2008 . The use of laboratory animals in toxicology research. In: Hays AW , ed. Principles and Methods in Toxicology , 5th ed. Boca Raton , FL: CRC Press ; 2008 : 1055 – 1101 . Google Scholar Google Preview OpenURL Placeholder Text WorldCat COPAC 73. Pasalic I , Bosnjak B , Tkalcevic VI , et al. Cage enrichment with paper tissue, but not plastic tunnels, increases variability in mouse model of asthma . Lab Anim. 2011 ; 45 : 121 – 12 . Google Scholar Crossref Search ADS PubMed WorldCat 74. Bazille PG , Walden SD , Koniar BL , et al. 2001. Commercial cotton nesting material as a predisposing factor for conjunctivitis in athymic nude mice . Lab Anim. 2001 ; 30 : 40 – 2 . OpenURL Placeholder Text WorldCat 75. Jennings M , Batchelor GR , Brain PF , et al. Refining rodent husbandry: the mouse . Lab Anim. 1998 ; 32 : 233 – 59 . Google Scholar Crossref Search ADS PubMed WorldCat 76. Van Loo PL , Baumans V . 2004. The importance of learning young: the use of nesting material in laboratory rats . Lab Anim. 2004 ; 38 : 17 – 24 . Google Scholar Crossref Search ADS PubMed WorldCat 77. Ras T , van de Ven M , Patterson Kane EG , et al. Rats’ preferences for corn vs wood-based bedding and nesting materials . Lab Anim. 2002 ; 36 : 420 – 5 . Google Scholar Crossref Search ADS PubMed WorldCat 78. Fidler IJ . Depression of macrophages in mice drinking hyperchlorinated water . Nature. 1977 ; 270 : 735 – 6 . Google Scholar Crossref Search ADS PubMed WorldCat 79. Hermann LM , White WJ , Lang CM . Prolonged exposure to acid, chlorine, or tetracycline in drinking water: effects on delayed-type hypersensitivity, hemagglutination titers, and reticuloendothelial clearance rates in mice . Lab Anim Sci. 1982 ; 32 : 603 – 8 . Google Scholar PubMed OpenURL Placeholder Text WorldCat 80. Homberger FR , Pataki Z , Thomann PE . Control of Pseudomonas aeruginosa infection in mice by chlorine treatment of drinking water . Lab Anim Sci. 1993 ; 43 : 635 – 7 . Google Scholar PubMed OpenURL Placeholder Text WorldCat 81. Hall JE , White WJ , Lang CM . Acidification of drinking water: its effects on selected biologic phenomena in male mice . Lab Anim Sci. 1980 ; 30 ( 4/1 ): 643 – 51 . Google Scholar PubMed OpenURL Placeholder Text WorldCat 82. Sasada T , Hinoi T , Saito Y , et al. Chlorinated water modulates the development of colorectal Tumors with chromosomal instability and gut microbiota in Apc-deficient mice . PLoS One. 2015 , 2015 Jul 17 ; 10 ( 7 ): e0132435 ). doi: 10.1371/journal.pone.0132435. OpenURL Placeholder Text WorldCat 83. Bleich A , Hansen AK . Time to include the gut microbiota in the hygienic standardization of laboratory rodents . Comp Immunol Microbiol Infect Dis. 2012 ; 35 : 81 – 92 . Google Scholar Crossref Search ADS PubMed WorldCat 84. Paik J , Pershutkina O , Meeker S , et al. Potential for using a hermetically-sealed, positive-pressured isocage system for studies involving germ-free mice outside a flexible-film isolator . Gut Microbes. 2015 ; 6 ( 4 ): 255 – 65 . doi: 10.1080/19490976.2015.1064576. Google Scholar Crossref Search ADS PubMed WorldCat 85. Nicklas W , Keubler L , Bleich A . Maintaining and monitoring the defined microbiota status of Gnotobiotic rodents . ILAR J. 2015 ; 56 ( 2 ): 241 – 9 . doi: 10.1093/ilar/ilv029 . Google Scholar Crossref Search ADS PubMed WorldCat 86. Bains RS , Cater HL , Sillito RR , et al. Analysis of individual mouse activity in group housed animals of different inbred strains using a novel automated home cage analysis system . Front Behav Neurosci. 2016 ; 10 : 106 . Google Scholar Crossref Search ADS PubMed WorldCat 87. Sun M , Choi EY , Magee DJ , et al. Metabolic effects of social isolation in adult C57BL/6 mice . Int Sch Res Not. 2014 ; 2014 : 690950 . OpenURL Placeholder Text WorldCat 88. Hong W , Kennedy A , Burgos-Artizzu XP , et al. Automated measurement of mouse social behaviors using depth sensing, video tracking, and machine learning . Proc Natl Acad Sci U S A. 2015 ; 112 : E5351 – 60 . Google Scholar Crossref Search ADS PubMed WorldCat 89. Shemesh Y , Sztainberg Y , Forkosh O , et al. High-order social interactions in groups of mice . Elife. 2013 : 2 e00759 . OpenURL Placeholder Text WorldCat 90. Salem GH , Dennis JU , Krynitsky J . SCORHE: a novel and practical approach to video monitoring of laboratory mice housed in vivarium cage racks . Behav Res Methods. 2015 ; 47 ( 1 ): 235 – 50 . Google Scholar Crossref Search ADS PubMed WorldCat 91. Burman O , Marsella G , Di Clemente A , Cervo L . The effect of exposure to low frequency electromagnetic fields (EMF) as an integral part of the housing system on anxiety-related behavior , cognition and welfare in two strains of laboratory mouse . PLoS ONE. May, 2018 . doi: 10.1371/journal.pone.0197054. OpenURL Placeholder Text WorldCat 92. Recordati C , De-Maglie M , Marsella G , Milite G , Rigamonti A , Paltrinieri S , Scanziani E . Long-term study on the effects of housing C57BL/6NCrl mice in cages equipped with wireless technology generating extremely low-intensity electromagnetic fields. Toxicol Pathol . May, 2019 . doi: 10.1177/0192623319852353. 93. Gerdin AK , Igosheva N , Roberson LA , et al. Experimental and husbandry procedures as potential modifiers of the results of phenotyping tests . Physiol Behav. 2012 ; 106 : 602 – 11 . doi: 10.1016/j.physbeh.2012.03.026 . Google Scholar Crossref Search ADS PubMed WorldCat 94. Kilkenny C , Browne WJ , Cuthill IC , et al. Improving bioscience research reporting: the ARRIVE guidelines for reporting animal research . PLoS Biol. 2010 ; 8 :e1000412. OpenURL Placeholder Text WorldCat © The Author(s) 2020. Published by Oxford University Press on behalf of the National Academies of Sciences, Engineering, and Medicine. All rights reserved. For permissions, please email: journals.permissions@oup.com This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)
TI - Reproducibility and Comparative aspects of Terrestrial Housing Systems and Husbandry Procedures in Animal Research Facilities on Study Data
JF - ILAR Journal
DO - 10.1093/ilar/ilz021
DA - 2020-04-10
UR - https://www.deepdyve.com/lp/oxford-university-press/reproducibility-and-comparative-aspects-of-terrestrial-housing-systems-SVYQYnY0uu
DP - DeepDyve
ER -