TY - JOUR
AU - Huerkamp, Michael J
AB - Abstract Environmental variables can have profound effects on the biological responses of research animals and the outcomes of experiments dependent on them. Some of these influences are both predictable and unpredictable in effect, many are challenging to standardize, and all are influenced by the planning and conduct of experiments and the design and operation of the vivarium. Others are not yet known. Within the immediate environment where the research animal resides, in the vivarium and in transit, the most notable of these factors are ambient temperature, relative humidity, gaseous pollutant by-products of animal metabolism and physiology, dust and particulates, barometric pressure, electromagnetic fields, and illumination. Ambient temperatures in the animal housing environment, in particular those experienced by rodents below the thermoneutral zone, may introduce degrees of stress and thermoregulatory compensative responses that may complicate or invalidate study measurements across a broad array of disciplines. Other factors may have more subtle and specific effects. It is incumbent on scientists designing and executing experiments and staff responsible for animal husbandry to be aware of, understand, measure, systematically record, control, and account for the impact of these factors on sensitive animal model systems to ensure the quality and reproducibility of scientific studies. animals, reproducibility of results, research design, stress, physiological, environment, quality control, animal models, husbandry INTRODUCTION For more than 50 years, it has been recognized that animal experiments are influenced by environmental factors, including components of research animal housing systems [1]. Environmental standardization and protection of experiments from unwanted variability has been the raison d’etre for research animal husbandry programs [2]. Early assertions have been verified across an array of environmental variables, some of which were not previously appreciated (eg, vibration) [3]. The most impactful parameters in animal research facilities are environmental temperature and illumination characteristics, and some have suggested that differences in these and other environmental parameters may contribute to the difficulties between laboratories to reproduce experimental findings [4]. The scientific community has become concerned that “preclinical research, especially work that uses animal models, seems to be the area that is currently most susceptible to reproducibility issues” [5, 6]. This growing concern has matured into a “reproducibility crisis,” culminating in animal experimentation taking a central role in the National Institutes of Health’s (NIH) Plans to Enhance Reproducibility and in ILAR’s work on the Reproducibility Issues in Research with Animals and Animal Models [5]. Paralleling efforts in the United States, the United Kingdom’s National Centre for the Replacement, Refinement and Reduction of Animals in Research (NC3Rs) recognizes the importance of animal environment factors in experimental designs. The NC3Rs’ Animal Research: Reporting of In Vivo Experiments (ARRIVE) defined which environmental details should be included in study design and methods descriptions: when (eg, time of day), where (eg, home cage or laboratory), housing type, bedding material, number of cage companions, husbandry conditions, light/dark cycle, temperature, and availability of feed [7]. These are not new concerns; a study published in 1976 covering 4080 articles published across 8 prominent journals showed that <10% of studies disclosed the ambient temperature, relative humidity (RH), room ventilation rate, photoperiod characteristics, caging system, housing density, bedding material, or sanitation schedule, among other features [8]. The American College of Laboratory Animal Medicine’s (ACLAM) Position Statement on Reproducibility is that “no detail of [animal care] should be deemed too small for sharing.” [9] ACLAM considers the documentation and reporting of these environmental parameters to be critical to the reproducibility of animal experimentation and therefore a key aspect of our collective dedication to Russell and Burch’s 3Rs [2]. Proper environmental controls have the potential to contribute to 2 of the 3Rs: (1) enhanced reproducibility reduces animals used in experimentation to the lowest number of animals needed, and (2) refinement of housing conditions to minimize stress, prevent distress, promote and sustain normal behavior (eg, reducing disruptions in the living space), standardize research variables to the extent possible, and contain costs in the conduct of research. These esteemed and varied organizations have taken the posture that rigor and attention to detail is critical. This position has been illustrated by repeated failed attempts to reproduce breakthrough papers: in a wide-ranging systematic effort, Bayer Healthcare in Germany was able to reproduce only 25% of tested publications [6] and Begley and Ellis at Amgen similarly were able to reproduce only 11% of “landmark” findings [10]. The findings of Bayer and Amgen appear to be generalizable: a survey of scientists reported that 70% were unable to reproduce the work of other groups [11, 12]. On a positive note, Begley and Ellis subsequently observed they could increase the reproducibility of some initial failed experiments where the “authors had paid close attention” to details [10], reinforcing the position that rigor is critical to the scientific process. Expanding on the importance of Begley and Ellis’ observation, NIH often found “variables that affect the results are unknown or not recognized and therefore not reported or described in sufficient detail. These confounding variables can prevent other investigators from reproducing the results if the unknown variables happen to be different between research groups” [13]. In animal experimentation, many of these unknown or unmeasured variables are environmental. Some variables are unknown because they have not been recognized and/or measured in a given experimental design. More vexing, there are undoubtedly variables that have yet to be discovered or have yet to be fully defined, and even new variables will emerge as new technologies alter environmental conditions of laboratories (eg, light-emitting diode lighting) [14]. The micro-environmental compartments within the macro-environment can vary from each other, between animal housing rooms, and from other sites such as laboratories in attributes such as ambient dry-bulb temperature and RH, gases produced or emitted in the vicinity of animals, and the concentration of particulates. Conditions whose values in and around the micro-environment are typically greater than the macro-environment are ambient temperature, RH, ammonia, carbon dioxide, and allergens [15, 16]. In a room supplied with a range of 8–15 air changes per hour (ACH) of fresh air, the ventilation rate may be about the same for a chain link pen enclosure; variable in open-top, solid-bottom cages depending on orientation of the cage enclosure with respect to the room air supply and exhaust sites; <1 ACH for a rodent cage covered by a filter top [17] (FT); and 30+ ACH where individual cage ventilation exists. Parameters that are identical between the macro- and micro-environments are barometric pressure and magnetic field effects. The closer a research institution is located to the equator, the greater the reliance on indoor cooling, whereas increasing proximity to a geographic pole puts greater design and performance emphasis of the building systems on heating. In a modern facility, the building system is designed to maintain temperature to The Guide for the Care and Use of Laboratory Animals (the Guide) standards and generally in the vicinity of 22°C, depending on occupant choice and species. To enable this result, fresh, outside air is usually heated or chilled to a specification programmed in the range of 10°C–16°C by the air handling system for distribution via the air supply ducts to individual rooms. At the point of delivery to each room, heat is usually supplemented via a terminal, hydronic reheat coil. These are sized to supply heated air up to 29°C–35°C, allowing for response to the thermostat when room temperature drops below the programmed set point. Humidification and dehumidification typically are managed at the central system or on a zonal basis; if not, there can be profound seasonal variation in indoor RH. A properly designed and functioning heating, ventilation, and air conditioning (HVAC) system will, ideally, fully and equally ventilate all areas of the room and respond to variations in climatic conditions outdoors or changes in the number and kind of animals and equipment indoors to maintain dry-bulb temperatures of ±1°C from individual room thermostatic set-point across the span of 18°C –29°C. An adequately functioning system will control RH within ±10% of a set-point in the range of 30%–70% year round [18]. Most HVAC systems are designed for average high and low temperatures and RH experienced in a geographic area within ±5% variation [19]. Moderate fluctuations in temperature and RH outside suggested ranges are generally well tolerated by most species commonly used in research as long as the fluctuations are brief and infrequent [18]. Effective control of all environmental variables at all times in many locations is a difficult goal, making monitoring systems designed into the facility requisite in the prompt identification of excessive drift or alarm conditions and in assuring quick response and restoration of environmental control. In line with expectations of the facilities to accommodate just about any species, type of study, and electrically powered scientific device, the HVAC system must be configured to be flexible and adaptable to the changing types and numbers of animals and equipment encountered during the life of the facility and should allow upgrades, renovation, and expansion with minimal disruption to activities. The ideal design prevents disturbances from physical plant maintenance by enabling routine or emergency maintenance and access to valves, filters, dampers, connections, and power to be accomplished outside of the animal housing and procedure rooms and will exclude pests so that studies are not impacted by using pesticides around animals or adding risks of infection or infestation introduced by vermin. The Guide [18] is a time-tested and enduring primer for animal research facility design and high-function operations. Readers are referred to it for review of the salient design features and operational fundamentals of air supply and exhaust volumes, exchange rates, flow patterns, the application of computational fluid dynamics, energy conservation possibilities, filtration, heating, cooling, differential pressurization of adjacencies to manage pathogens, odors, allergens and other hazards, and the value of system redundancy and preventive maintenance [18]. If one seeks to build on the wisdom of that foundation, there is the timeless, seminal contribution of Ruys [20] and more recent publications [21–23]. We embrace ACLAM’s position that it is “incumbent on laboratory animal veterinarians and the scientific community to define elements of study design that affect experimental reproducibility” [9]. In the spirit of the ARRIVE guidelines, [7] the Guide, [18] the ACLAM Position Statement on Reproducibility [9], and NIH’s multi-front effort to improve reproducibility and transparency [5, 13], we have established 4 aims for this manuscript: (1) define environmental parameters as core experimental variables, (2) arm the reader with the knowledge of which environmental parameters are most important, (3) report how these parameters influence or are likely to impact animal research, and (4) review methods used to measure these parameters so that the reader can carry the mantle at their respective institution to improve reproducibility of animal experimentation by ensuring environmental control. This submission is rodent-centric by necessity as the preponderance of what is known relates to rodents; however, readers will recognize that the general principles presented are applicable across species unless established, and noted herein, to be different. OVERVIEW OF PRIMARY ENVIRONMENTAL VARIABLES Temperature The temperature supplied to the room by the HVAC is generally slightly to significantly lower than that in proximity to the animal. A number of factors influence this relationship, most importantly caging type, population density, type of contact bedding, and the presence of nesting material and/or shelters. In general, these factors add 1°C–2°C. Importantly, there are temperature gradients across all 3 dimensions of most rooms, with the macro-environmental temperature generally warmer toward the ceiling due to the relative buoyancy of hot air relative to cold and any heat emitted from ceiling light fixtures. The design of the air supply diffuser and exhaust register, as well as their locations and the positioning of equipment, may impede complete and homogeneous mixing of the air and create subtle effects in temperature exposure. The interplay of ambient temperatures and physiology is complex. Endotherms (“warm-blooded” animals) strive to maintain their core body temperature irrespective of the surrounding environment through a variety of behaviors, peripheral vessel constriction and dilation, and body postures [24, 25] The temperature at which energy-conserving behaviors and physiological changes are in equilibrium with the heat loss to the environment (entropy) is referred to as the thermal neutral zone (TNZ) [24]., [26, 27] Generally, animals that are larger, well insulated (eg, thick fur and ample subcutaneous fat), and with low surface area to volume ratios (ie, “rounder,” which is also a function of posture) [28] have a lower TNZ [25, 29]. Conversely, smaller, low insulated, and high–surface area animals have a higher TNZ (Figs 1 and 2), which was described in the seminal text by Scholander et al, 1950 [26]. For example, the TNZ of dogs and cats is 20°C–25°C, whereas rabbits—with their thick coats and relatively round morphology—have a lower TNZ of 15°C–20°C [30] (though auricular vascular constriction and dilation provides considerable ability to dissipate excess heat) [31], and small rodents such as mice and gerbils have a TNZ of approximately 30°C [24]. Table 1 Definitions Allostasis: The process of restoring homeostasis after perturbation or recalibrating physiology or behavior in compensation to an inability to return to the original steady state. Critical temperature, lower: The ambient temperature below which the rate of metabolic heat production of a resting thermoregulating tachymetabolic (“warm-blooded”) animal must be increased by shivering and/or nonshivering thermogenesis to maintain thermal balance and prevent hypothermia [25]. Dew point: A measure of atmospheric moisture defined as the temperature to which air must be cooled at constant pressure to be fully saturated with water vapor such that it will condense into water in the liquid state (ie, form dew). Homeostasis: The regulation of an organism to remain in balance. Homeothermy: State of maintaining a stable body temperature regardless of external influence. Humidity, relative (RH): Ratio of saturated vapor pressure at dew-point temperature of the enclosure to the saturated vapor pressure at dry-bulb temperature [25]; often expressed as a percentage. Macro-environment: The physical environment of the secondary enclosure (eg, a room, a barn, or an outdoor habitat) [16]. The preponderance of modern environmental standards used to guide operation of the vivarium is based on room conditions. Micro-environment: The immediate physical environment surrounding the animal (ie, the environment in the primary enclosure such as the cage, pen, or stall) [16]. In many cases under indoor conditions, but not all, the characteristics of the micro-environment closely resemble and are directly linked to the macro-environment. Temperature, dry-bulb: Temperature measured by a thermometer, whereas the thermodynamic wet-bulb temperature of a sample of air is the lowest temperature to which it can be cooled by evaporating water adiabatically. This measurement is compared with an ordinary thermometer, or a dry bulb, to determine estimate ambient humidity [25]. Thermogenesis, diet-induced: Increase of nonshivering thermogenesis occurring especially in rodents when transferred from standard food to high-palatable cafeteria diet, of which the animals consume more but dissipate part of the surplus caloric intake by enhanced heat production [25]. Thermoneutral zone (TNZ): The range of ambient temperature at which temperature regulation is achieved without regulatory changes in metabolic heat production or evaporative heat loss [25]. TNZ will therefore be different when insulation, posture, or basal metabolic rate varies. Allostasis: The process of restoring homeostasis after perturbation or recalibrating physiology or behavior in compensation to an inability to return to the original steady state. Critical temperature, lower: The ambient temperature below which the rate of metabolic heat production of a resting thermoregulating tachymetabolic (“warm-blooded”) animal must be increased by shivering and/or nonshivering thermogenesis to maintain thermal balance and prevent hypothermia [25]. Dew point: A measure of atmospheric moisture defined as the temperature to which air must be cooled at constant pressure to be fully saturated with water vapor such that it will condense into water in the liquid state (ie, form dew). Homeostasis: The regulation of an organism to remain in balance. Homeothermy: State of maintaining a stable body temperature regardless of external influence. Humidity, relative (RH): Ratio of saturated vapor pressure at dew-point temperature of the enclosure to the saturated vapor pressure at dry-bulb temperature [25]; often expressed as a percentage. Macro-environment: The physical environment of the secondary enclosure (eg, a room, a barn, or an outdoor habitat) [16]. The preponderance of modern environmental standards used to guide operation of the vivarium is based on room conditions. Micro-environment: The immediate physical environment surrounding the animal (ie, the environment in the primary enclosure such as the cage, pen, or stall) [16]. In many cases under indoor conditions, but not all, the characteristics of the micro-environment closely resemble and are directly linked to the macro-environment. Temperature, dry-bulb: Temperature measured by a thermometer, whereas the thermodynamic wet-bulb temperature of a sample of air is the lowest temperature to which it can be cooled by evaporating water adiabatically. This measurement is compared with an ordinary thermometer, or a dry bulb, to determine estimate ambient humidity [25]. Thermogenesis, diet-induced: Increase of nonshivering thermogenesis occurring especially in rodents when transferred from standard food to high-palatable cafeteria diet, of which the animals consume more but dissipate part of the surplus caloric intake by enhanced heat production [25]. Thermoneutral zone (TNZ): The range of ambient temperature at which temperature regulation is achieved without regulatory changes in metabolic heat production or evaporative heat loss [25]. TNZ will therefore be different when insulation, posture, or basal metabolic rate varies. Open in new tab Table 1 Definitions Allostasis: The process of restoring homeostasis after perturbation or recalibrating physiology or behavior in compensation to an inability to return to the original steady state. Critical temperature, lower: The ambient temperature below which the rate of metabolic heat production of a resting thermoregulating tachymetabolic (“warm-blooded”) animal must be increased by shivering and/or nonshivering thermogenesis to maintain thermal balance and prevent hypothermia [25]. Dew point: A measure of atmospheric moisture defined as the temperature to which air must be cooled at constant pressure to be fully saturated with water vapor such that it will condense into water in the liquid state (ie, form dew). Homeostasis: The regulation of an organism to remain in balance. Homeothermy: State of maintaining a stable body temperature regardless of external influence. Humidity, relative (RH): Ratio of saturated vapor pressure at dew-point temperature of the enclosure to the saturated vapor pressure at dry-bulb temperature [25]; often expressed as a percentage. Macro-environment: The physical environment of the secondary enclosure (eg, a room, a barn, or an outdoor habitat) [16]. The preponderance of modern environmental standards used to guide operation of the vivarium is based on room conditions. Micro-environment: The immediate physical environment surrounding the animal (ie, the environment in the primary enclosure such as the cage, pen, or stall) [16]. In many cases under indoor conditions, but not all, the characteristics of the micro-environment closely resemble and are directly linked to the macro-environment. Temperature, dry-bulb: Temperature measured by a thermometer, whereas the thermodynamic wet-bulb temperature of a sample of air is the lowest temperature to which it can be cooled by evaporating water adiabatically. This measurement is compared with an ordinary thermometer, or a dry bulb, to determine estimate ambient humidity [25]. Thermogenesis, diet-induced: Increase of nonshivering thermogenesis occurring especially in rodents when transferred from standard food to high-palatable cafeteria diet, of which the animals consume more but dissipate part of the surplus caloric intake by enhanced heat production [25]. Thermoneutral zone (TNZ): The range of ambient temperature at which temperature regulation is achieved without regulatory changes in metabolic heat production or evaporative heat loss [25]. TNZ will therefore be different when insulation, posture, or basal metabolic rate varies. Allostasis: The process of restoring homeostasis after perturbation or recalibrating physiology or behavior in compensation to an inability to return to the original steady state. Critical temperature, lower: The ambient temperature below which the rate of metabolic heat production of a resting thermoregulating tachymetabolic (“warm-blooded”) animal must be increased by shivering and/or nonshivering thermogenesis to maintain thermal balance and prevent hypothermia [25]. Dew point: A measure of atmospheric moisture defined as the temperature to which air must be cooled at constant pressure to be fully saturated with water vapor such that it will condense into water in the liquid state (ie, form dew). Homeostasis: The regulation of an organism to remain in balance. Homeothermy: State of maintaining a stable body temperature regardless of external influence. Humidity, relative (RH): Ratio of saturated vapor pressure at dew-point temperature of the enclosure to the saturated vapor pressure at dry-bulb temperature [25]; often expressed as a percentage. Macro-environment: The physical environment of the secondary enclosure (eg, a room, a barn, or an outdoor habitat) [16]. The preponderance of modern environmental standards used to guide operation of the vivarium is based on room conditions. Micro-environment: The immediate physical environment surrounding the animal (ie, the environment in the primary enclosure such as the cage, pen, or stall) [16]. In many cases under indoor conditions, but not all, the characteristics of the micro-environment closely resemble and are directly linked to the macro-environment. Temperature, dry-bulb: Temperature measured by a thermometer, whereas the thermodynamic wet-bulb temperature of a sample of air is the lowest temperature to which it can be cooled by evaporating water adiabatically. This measurement is compared with an ordinary thermometer, or a dry bulb, to determine estimate ambient humidity [25]. Thermogenesis, diet-induced: Increase of nonshivering thermogenesis occurring especially in rodents when transferred from standard food to high-palatable cafeteria diet, of which the animals consume more but dissipate part of the surplus caloric intake by enhanced heat production [25]. Thermoneutral zone (TNZ): The range of ambient temperature at which temperature regulation is achieved without regulatory changes in metabolic heat production or evaporative heat loss [25]. TNZ will therefore be different when insulation, posture, or basal metabolic rate varies. Open in new tab Figure 1 Open in new tabDownload slide Metabolic rate relative to basal metabolism as a function of air temperature for variety of mammals. Reprinted with permission from Scholander et al, 1950 [26]. Figure 2 Open in new tabDownload slide Simulated lower critical air temperature (ie, the lower bound of the thermal neutral zone) for different size endotherms as a function of posture and wind speed. Results are presented for animals with and without fur. Notice the smaller animals have much stronger response to lower wind speeds, including speeds common to ventilated caging, than larger animals because of their thinner insulation. Reprinted from Porter and Kearney, 2009 [25] under conditions of approved use of the Proceedings of the National Academies of Sciences of the United States of America. Below the TNZ, endotherms commonly generate additional biological heat via thermogenesis at a great energy cost to maintain core body temperatures [32]. Thermogenesis occurs predominantly by shivering of skeletal muscle in larger species, providing there is adequate muscle mass, or nonshivering by brown adipose tissue in small (notably rodents), artic, and neonatal mammals [32]. In an alternate strategy to thermogenesis, a number of endothermic species may strategically lower their core body temperature to balance caloric inflow and outflow, a characteristic termed heterothermy [33]. In the extreme, heterothermy can result in a reduced resting metabolism along a spectrum of hibernation, torpor, or torpor-like states with reduced energy demand [34]. Especially important to the laboratory setting, mice will exhibit these phenotypes below their TNZ and are best characterized as “facultative daily heterotherms” rather than true endotherms, that is, the thermal physiology of the mouse is dependent on the context of their environment (eg, bedding, social huddling, environmental temperature, etc), time of day (circadian), and not fixed at a specific body temperature [34, 35]. In fact, a torpor-like state is only avoided in laboratory settings due to the common ad lib feed availability [34, 35]. Songbirds exhibit gonadal regression at cold temperatures [36]. Above the TNZ, behavioral and morphophysiological adaptations allow for cooling. The tail, making up 7% of the mass of a rat, is a site of up to 25% heat dissipation [37]. A hallmark of being at or above thermo-neutrality for mice is that social huddling is absent [37]. The TNZ varies considerably both within and between individuals, species, and seasonality, which is largely mitigated in controlled laboratory environments. Therefore, the physiological state of a laboratory animal is not only a function of temperature, RH and ventilation, housing system, and husbandry program [38], but also phenotype (eg, hirsute or nude) [25, 39], body condition [40], physiological state (eg, circadian rhythm), [24] recovery from anesthesia/surgery, life stage (including age, pregnancy, and lactation status [32, 41]), available insulation (eg, nesting and bedding materials [42]), the ability to huddle to reduce collective exposed surface area [39], and the ability to select ambient temperature (thermotaxis) [43–45]. Attempts to characterize the physiological state of laboratory animals can be performed by a variety of methods: telemetry to measure core temperatures and cardiovascular parameters [46], thermography to measure ocular proxies of core temperature in rodents and rabbits [47], indirect calorimetry to measure metabolic expenditures [48], and thermography to measure non-shivering thermogenesis of brown fat [47, 49], vasodilation of the rodent tail [44, 50], or the ears of rabbits [31]. Body temperature–monitoring methods were extensively reviewed by Meyer, Ootsuka, and Romanovsky [51]. In the macro-environment, temperature is measured using any of different types of thermometers, RH with a hygrometer or any one of a variety of types of digital recorders, and each in combination with a thermohygrometer (sling psychrometer). Ventilation is calculated by measuring the room, or exhaust, in units of volume per unit of time using an anemometer and dividing that value into the cubic volume of the site. Differential air pressure can be measured using smoke sticks or various types of manometers. Impacts of Environmental Temperature. From the perspective of the experimentalist, the control of variability is best realized during study planning and execution, while for the animal resources program it is ensured operationally. Temperature, RH, and ventilation in the 2 environmental compartments together determine relative heat loss or retention and chilling [52] and ultimately contribute substantially to metabolic rate and other biological processes as previously noted. Failure to adequately control these variables during any 1 experiment or across a range of them leads to the possibility of disparate results and risks of faulty conclusions in studies across many disciplines. The Guide recommends vivarium temperatures range from 20°C to 26°C [18]. As a practical matter, most vivaria are maintained at a “room” temperature of approximately 20°C–21°C, which is a comfortable environment for human work [53] and with associated utility costs that institutions have been willing to bear. This means large animal species are often housed at or near their TNZ whereas rodents are housed markedly below. Cold stress is especially acute in mice, whose diminutive size and high surface area determines their higher TNZ. For rats, their TNZ is slightly lower at 28°C–30°C [37]. There are species-related differences in thermoregulatory responses between mice and rats. Rats, in particular, have a larger mass and lower basal metabolic rate (as normalized to body mass) and exhibit relatively smaller metabolic compensatory changes compared with mice [37]. Mice have a core body temperature approximately 1°C lower than rats and a lower critical temperature of 18°C compared with 12°C for rats [37]. The lower critical temperature is the minimum body temperature that can be tolerated by an organism, below which it becomes hypothermic [54]. Laboratory rodents kept at 20°C–22°C and unable to sufficiently adapt behaviorally, such as in huddling, building nests, and increasing energy consumption, and physiologically are chronically cold stressed, which is exacerbated without the provision of nesting material [55–57] and air flow in some individually ventilated cages [28, 49, 58] However, laboratory mice compensate with nonshivering thermogenesis [32, 49] and offset the energy costs with increased feed consumption [34, 35], which has consequences for certain experiments. The thermal physiology of marmosets, tamarins, and other similarly sized platyrrhines (New World monkeys) generally matches projected models [26] based on surface area to volume ratios, and their TNZs can be reasonably approximated [59, 60]. Behaviors play a significant role in adaptability to environmental temperature variations [61]. Platyrrhine behavioral adaptations, such as avoiding excessive cold and heat (thermotaxis), are exhibited prior to physiological changes [62, 63]. Once behavioral thermal regulation options are exhausted, platyrrhines utilize typical mammalian physiological strategies to minimize energy and water loss: piloerection [64], vasoconstriction of the feet and tail [65], sweat (though more limited than humans with eccrine glands only being found in the palms and soles) [66–69], and thermogenesis [62]. Despite the general similarities to other mammals, the reader should appreciate that although at the whole organism level platyrrhines are typical, species-specific behavioral and physiological adaptations exist [65, 70]. There is considerable thermal cellular physiology and enzymatic variation between primates and rodents [71, 72] that cannot be appreciated at the macro-thermoregulatory level by techniques such as calorimetry. For example, cold-acclimated rodents process evaluated levels of renal and hepatic mitochondrial succinoxidase and nicotinamide-adenine dinucleotide phosphate (NADPH)-cytochrome; in contrast, cold-acclimated squirrel monkeys only increase renal mitochondrial succinoxidase (and not hepatic) and have no environmental-dependent variation in NADPH-cytochrome in either tissue [72]. These findings highlight the biochemical and tissue-specific difference in thermoregulation across species. The reader seeking more details is referred to the “Thermoregulation in the Squirrel monkey” chapter of the Handbook of Squirrel Monkey Research [67]. Ambient temperatures are known to modify a wide variety of animal models of disease. Some prime examples of sub-TNZ temperature effects include increased sympathetic outflow and catecholamines [73], increased blood pressures and heart rate (602 ± 5 beats/m at Ta 23°C compared with 351 ± 11 beats/m at Ta 30°C) [74], alteration of immune function [75] and infectious disease pathophysiology [76–80], variation in tumor growth and immune-microenviroment [81], changes in metabolism, [49, 82] influences on muscle and bone physiology and anatomical development [83], modified reaction to atherogenic [84] and pyrogenic agents [37], insensitivity of blood cells to irradiation [85], modulation of the response to toxins, [86] and impaired rodent reproduction and lactation [82]. The translational impact is largely unknown. Several authors have speculated that standard sub-TNZ temperatures have a negative impact on the translational value of animal models and may be a contributor of the irreproducibility crisis, especially of rodent models [87, 88]; however, dissenting opinions exist [89]. Another risk to all thermal environmental compartments is the failing of the terminal reheat system (TRS). By design, these components of the HVAC system regulating room temperature will fail open, closed, or in place with varying ramifications. If they fail closed (the Guide preference) [18], the consequence is that fresh air is supplied to the rooms at 10°C–16°C exiting the air handling unit (AHU) unresponsive to the thermostat. Animals then are exposed to these temperatures indoors, possibly requiring animal care program intervention (eg, relocation of the census to a warmer location) until the system is repaired. In the experience of the authors (M.J.H., V.L.) with mice in this scenario, providing there is group housing for huddling and ample nesting material, passive observation suggests that the animals behave normally and are not obviously affected. Given the importance of temperature on biological processes, it is imperative that scientists be informed of these sorts of failures so that they can account for the variability in the conduct of any experiment. For TRS that fail in place, providing it did not occur at or near an extreme of a fully opened or closed valve, the mean temperature will vary from the programmed set-point, and there may be greater variability around that mean due to the effect of the inflexible TRS. The real hazard, however, is in the TRS that fail open with 82°C water (or steam) circulated through the coil. In a room receiving 10 ACH, where in theory the full volume of the air in the room is exchanged every 6 minutes, the room temperature rapidly escalates to 40°C–50°C with the ensuing specter of compromised experiments, animal deaths, and impaired breeding colony production [90]. The importance of this is not only in proper design, but in deploying environmental monitoring systems that detect failures and emergency conditions and alarm and immediately notify responders. From the animal care perspective, factors that are highly influential in the macro- and micro-environmental relationship can be loosely divided into those determined by the husbandry program and those that are the consequence of vivarium design and construction. The former includes the type of cage or enclosure, particularly where air exchange is influenced at the cage level (eg, by FT, individual ventilation of cages), contact bedding material, shelf location, population density, schedule of cage servicing, and adherence to regulation [18]. Products of the latter are the filtration and temperature of supply air, room ventilation patterns and rates, exhaust destination for the effluent from ventilated systems (ie, returned to the room or ducted from the rack to room exhaust), location of environmental monitoring sensors (eg, room wall vs exhaust duct for temperature and RH), redundancy of systems to ensure uninterrupted operation, and effectiveness of thermostatic response. Achieving precision across this myriad array is a complex and sometimes bewildering proposition; however, the importance of maintaining precise temperature control and reporting these values in manuscripts cannot be overemphasized given the impact of environmental temperature on metabolism, behavior, and a wide range of other biological processes. The seemingly simple solution to macro-environments, and thus micro-environments, too cold for the conduct of valid studies with mice and rats is to warm rooms to 28°C while commensurately adjusting RH and then enable the conduct of better experiments. This approach is not tenable, however, as it does not comport with sustainability goals or productive working environments [53] and may result in increased aggression in mice [91] and prohibitively increase utility bills. Each 3150-cubic foot room heated +6°C, if ventilated at 8–10 ACH, requires 1.5–1.7 KWh annually and, at an average of $0.12/KWh, adds $1600–$1700/room/year to the utility bill for the institution [92]. Instead, scientists and animal care specialists wait for a pioneering and enterprising allied industry to manufacture and sell housing systems with innovative micro-environmental warming approaches economically delivered at the rack or cage level. For those key experiments where the thermal environment is a critical variable to control, remedies beyond the nominal capabilities of the facility and its housing modalities are to consider keeping animals in groups, provide nesting material and/or shelters for those genotypes inclined to use them, offer supplementary heating (as might be done for swine or poultry), adopt warming solutions focused on the micro-environment of small enclosures [43], and use sealed, specialized chambers that allow precise environmental control. Air Quality and Ventilation Air quality standards for the macro-environment are dictated by human exposure limits set by regulatory agencies with little guidance specific to the laboratory animal community. The Occupational Safety and Health Administration (OSHA) has established mandatory permissible exposure limits [93], although states can set more stringent regulatory requirements, as is done by California [94]. Recommended human exposure limits are established by the National Institute for Occupational Health and Safety of the Center for Disease Control and Prevention and the professional organization The American Conference of Governmental Industrial Hygienists [95]. OSHA recognizes many more components of air quality than are relevant for laboratory animal housing, but ammonia, carbon dioxide, microbes, and particulates are the most likely to be significant for research animals [94, 96] Research efforts on the effects of air quality in animals have focused on levels that exceed human thresholds, but no prescribed standards have been set for laboratory animals. The Guide for the Care and Use of Laboratory Animals only recommends that air pollutants are minimized [18] but does not give more specific guidance. Depending on whether the cage is open, fully enclosed with a FT, or individually ventilated, there can be gradations in differences of air quality between that in the cage in proximity to the animals and the macro-environment [16, 97–100]. For example, a differentiating characteristic of individually ventilated caging (IVC) and gnotobiology isolators is that the supply air is cleaner than that of the room as it is highly pure, usually HEPA-filtered, and free of dust, microorganisms, and particles suspended in the air. In general and with respect to rodents, air quality is better in suspended wire cages, uncovered shoebox cages, and IVC than in static (unventilated; sometimes called isolator) FT cages [101, 102]. The micro-environment of a static FT cage is more likely to contribute to research variability than the macro-environment because of the relative accumulation of contaminants contained by the FT lid and the high variability in air quality reported. [98–101, 103–105]. Although micro-environmental conditions are affected by the room air quality, intracage levels of gases and RH can be much higher in static relative to ventilated cages [15, 102, 106]. The primary potential concerns for air quality in a FT cage are ammonia, carbon dioxide, and particulates as these are products of the animals and materials within the cage. Ammonia will arise from enzymes in the bedding or from bacteria that convert urea in urine to ammonia [107]. Carbon dioxide is the byproduct of the animal’s respiration. Particulates may originate from the macro-environment but are more often derived from components added to the cage such as food and bedding [15]. Unlike the macro-environment, intracage air quality has no generally accepted standards, and institutions are left to extrapolate from macro-environmental limits as to acceptable intracage accumulations of these parameters. Regulatory requirements for humans are often used as a reference point for evaluating intracage air quality (Table 2), but the relevance of these values to rodents has not been established [108]. These quality parameters are not measured regularly in animal cages, but familiarity with how they could affect the animals and research may be useful when establishing or changing housing and husbandry practices and for experimenters to consider regarding potential impacts to their models. Other gaseous pollutants noted to be present in rodent cages, but not demonstrated to be consequential, include methane [101, 109], acetic acid [110], hydrogen sulfide [109], and sulfur dioxide [110]. Options to minimize environmental fluctuations associated with unventilated FT cages include frequent cage changes, decreased cage population densities, individual cage ventilation, and/or using mice free of urease-producing bacteria [111]. Table 2 Regulatory Guidelines for Human Exposure to Airborne Contaminantsa . OSHA PEL . NIOSH REL 10-hour TWA . ACGIH 2018 TLV 8-hour TWA . Ammonia 50 ppm (35 mg/m3) 25 ppm STEL of 35 ppm 25 ppm STEL of 35 ppm Carbon dioxide 5000 ppm 5000 ppm STEL 30 000 ppm 5000 ppm STEL 30 000 ppm Particulates not otherwise regulated 15 mg/m3 total dust 5 mg/m3 respirable fraction . OSHA PEL . NIOSH REL 10-hour TWA . ACGIH 2018 TLV 8-hour TWA . Ammonia 50 ppm (35 mg/m3) 25 ppm STEL of 35 ppm 25 ppm STEL of 35 ppm Carbon dioxide 5000 ppm 5000 ppm STEL 30 000 ppm 5000 ppm STEL 30 000 ppm Particulates not otherwise regulated 15 mg/m3 total dust 5 mg/m3 respirable fraction ACGIH = American Conference of Government Industrial Hygienists, a private, nonprofit, nongovernmental, scientific professional organization, Cincinnati, OH; NIOSH = National Institute of Occupational Safety and Health; OSHA = Occupational Safety and Health Administration; PEL = permissible exposure limit; REL = recommended exposure limit; STEL = short-term exposure limit (usually for 15 continuous minutes); TLV = Threshold Limit Value, a registered and reserved term of the ACGIH [206]; TWA = time weighted average; defined as the maximum average concentration of an inhalable material in the workplace to which workers can be exposed during a full work day and workweek, over a lifetime, without experiencing significant adverse health effects. aExcerpted from OSHA AnnotatedTable Z-1 for limits of airborne contaminants [94]. The respirable fraction referenced by OSHA is the particle size that can penetrate into the gas exchange portion of the lung in humans [123]. The TLVs and other exposure limits are intended to be solely used by qualified industrial hygienists in promoting the health and safety of human workers and, as such, herein are provided as guidelines, not standards, to be considered in the care and use of research animals and in the safety of humans working with them [95, 96] The terms TLV and ACGIH were reprinted with permission of the ACGIH. Copyright 2019. Open in new tab Table 2 Regulatory Guidelines for Human Exposure to Airborne Contaminantsa . OSHA PEL . NIOSH REL 10-hour TWA . ACGIH 2018 TLV 8-hour TWA . Ammonia 50 ppm (35 mg/m3) 25 ppm STEL of 35 ppm 25 ppm STEL of 35 ppm Carbon dioxide 5000 ppm 5000 ppm STEL 30 000 ppm 5000 ppm STEL 30 000 ppm Particulates not otherwise regulated 15 mg/m3 total dust 5 mg/m3 respirable fraction . OSHA PEL . NIOSH REL 10-hour TWA . ACGIH 2018 TLV 8-hour TWA . Ammonia 50 ppm (35 mg/m3) 25 ppm STEL of 35 ppm 25 ppm STEL of 35 ppm Carbon dioxide 5000 ppm 5000 ppm STEL 30 000 ppm 5000 ppm STEL 30 000 ppm Particulates not otherwise regulated 15 mg/m3 total dust 5 mg/m3 respirable fraction ACGIH = American Conference of Government Industrial Hygienists, a private, nonprofit, nongovernmental, scientific professional organization, Cincinnati, OH; NIOSH = National Institute of Occupational Safety and Health; OSHA = Occupational Safety and Health Administration; PEL = permissible exposure limit; REL = recommended exposure limit; STEL = short-term exposure limit (usually for 15 continuous minutes); TLV = Threshold Limit Value, a registered and reserved term of the ACGIH [206]; TWA = time weighted average; defined as the maximum average concentration of an inhalable material in the workplace to which workers can be exposed during a full work day and workweek, over a lifetime, without experiencing significant adverse health effects. aExcerpted from OSHA AnnotatedTable Z-1 for limits of airborne contaminants [94]. The respirable fraction referenced by OSHA is the particle size that can penetrate into the gas exchange portion of the lung in humans [123]. The TLVs and other exposure limits are intended to be solely used by qualified industrial hygienists in promoting the health and safety of human workers and, as such, herein are provided as guidelines, not standards, to be considered in the care and use of research animals and in the safety of humans working with them [95, 96] The terms TLV and ACGIH were reprinted with permission of the ACGIH. Copyright 2019. Open in new tab The drafts and cooling effects created by intracage ventilation add an additional potential variable beyond the effects of ventilation on gaseous air quality. IVC have different designs for air input and exhaust, hourly ACH, and air distribution patterns depending on manufacturer [106, 108, 112]. In particular, the design and air velocity from the input valve affect the intracage air distribution [102], and airflow even for the same rack could change over time as the air inlets and outlets become occluded with debris [113]. These differences in cage ventilation can lead to variations in air quality and temperature experienced by animals because intracage ventilation creates drafts and affects humidity and the accumulation of gaseous products especially relative to static cages [97, 102, 108]. Ammonia is the most common and abundant gaseous pollutant associated with rodent cages for which there is the potential for high levels with extensively documented adverse effects [114]. Control of ammonia is particularly challenging given that many rodents, even those that are specific pathogen-free, may not be well defined regarding their inherent bacterial flora, and they may be colonized with urease-producing bacteria capable of catalyzing the conversion [107]. Factors that affect ammonia levels beyond inherent animal characteristics include RH, caging types, and bedding. Male mice excrete approximately twice as much urine as females [115], and cages with male mice will be associated with higher ammonia values than females [116–118]. Although the studies comparing caging type are limited, cage type has been shown to have an effect and could play a role both by containing ammonia and increasing RH levels [104, 119, 120]. Urine output also increases with age [115] and also possibly by strain. Bedding materials vary in their ability to control ammonia levels, with corncob having relatively high absorbency and demonstrated suppressive effects, followed by paper and then wood-based beddings [110, 121]. An animal resources program concerned about the effects of husbandry modifications on ammonia levels should measure them under their specific conditions given the wide variability in reported intracage concentrations despite seemingly similar housing conditions. Minimizing dust is a general goal within cages, but the levels and particle size relevant for rodents is unknown. Particles lodge within the respiratory system depending on size, with particles 2–10 μm more likely to reach alveoli and larger particles becoming lodged in the upper respiratory tract of humans [122]. Particulates within an enclosed rodent cage can be higher than in the macro-environment and arise primarily from sources within the cage, but particles within the room can also be found within static FT cages [15]. Bedding is a source of particulates, [15] although most common contemporary beddings are low in dust, with levels typically <0.15%. Corncob- and paper-based bedding have the lowest level of particulates, and wood-based products have the highest levels. The units and methods of measurement for gases and particulates are applicable to both the macro- and micro-environment. Gases are reported as either a mass to volume ratio, typically mg/m3, or parts per million (ppm). These units can be converted from one to another based on the molecular weight and using a specific calculation where the concentration in mg/m3 is equal to the product of 0.0409 multiplied by the molecular weight and by the concentration in parts per million [123]. For example, ammonia with a molecular weight of 17.031 Daltons calculates to 1 mg/m3 or 1.4 ppm. Particulates are described as either mass or number to volume ratio, such as milligrams per cubic meter or particles per cubic centimeter. Gases can be detected manually with commercially available gas detection tubes and bellows, with the tubes containing a chemical that reacts with the gas when air is drawn into the tube. These tubes are particularly useful for determining levels in small, enclosed environments such as rodent cages. OSHA also recognizes portable infrared spectrophotometers or a chip measurement system for gas level detection and includes the specific models and manufacturers commonly used for these measurements in the OSHA Occupation Chemical Database description for each chemical [124]. OSHA recommends a particle counting instrument for airborne particulates, most commonly an aerosol photometer, also known as a nephelometer, light scattering meter, or aerosol monitor. This device relies on dust scattering light or infrared radiation to quantify particle levels. Impacts of Air Quality. The direct effects of room air quality on research are few but have been previously summarized [107]. When research outcomes are highly dependent on RH, such as with the transmission or aerosolized infectious diseases, the levels should be specifically controlled. Although difficult to generalize, at extremes of <30% or >70% RH can notably influence food and water consumption [125], activity [82], postnatal development [126], transmission of infectious agents [82, 127–130], ocular physiology [131], and the transcutaneous absorption of drugs [82]. RH can also affect nociception [132] and the development and progression of skin conditions, most notably ringtail in rodents, a condition ranging from focal tail tip necrosis to an extensive contortive deformity of the tail, usually, but not always, associated with low extremes of RH [133] (ie, dew point) in combination with the ambient temperature, thus affecting evaporative heat loss that in turn influences temperature perception. At high RH, evaporative heat losses are retarded, creating the perception of greater heat effect [134–137]. Humans experience this as “mugginess.” Likewise, low RH (also achieved in effect by the movement of air through various means of ventilation [138, 139]) promotes evaporative heat loss [140–144] and a cooling effect, which may increase comfort at high temperatures or increase chilling at temperatures below the TNZ [137, 144, 145, 146]. The movement of air through a room or caging system thus in and of itself can have morphophysiological implications. Although the cause was not identified, in 1 report rats housed in IVC cages had lower respiratory function and higher levels of lung inflammation relative to rats housed in static cages [147]. Although the temperatures in static cages have not been found to be lower than IVC cages when directly compared, mice in IVC cages can have signs of cold stress relative to those in static cages [58, 99, 108]. The underlying reasons are unknown, but drafts and cold stress could be reasons for some of the differences seen between mice in static and IVC cages related to growth and behavior. Mice in motor-free IVC and static cages have greater growth than mice in IVC cages with 60 ACH [102, 148]. In addition, mice appear averse to direct airflow with IVC systems set at 60 and 100 AHC [148, 149]. Rats not only prefer air exchange rates of 50 ACH relative to 80 ACH and higher, but also 80 ACH and above have significant impacts on heart rate and blood pressure [150]. At least some IVC caging systems are set to lower ACH rates of closer to 30 ACH, and although gaseous air quality has been studied at that AHC, [97] preference tests for rats and mice at these levels have not been evaluated to the authors’ knowledge. Perhaps related to the aversion to high ACH rates, IVC have been associated with affecting behavioral phenotypes of mice [151, 152]. Importantly, aversion and cold stress in mice seen at the higher ACH can be mitigated with protective materials such as shelters and nesting materials [58, 149], although the ability for these materials to reduce potential impacts of IVC on behavioral phenotypes is unknown. Increasing RH, particularly that above 60%, directly and significantly promotes the production of ammonia [153–155], enables the hatching of food pests [156], and results in condensate that fosters mold growth on feed [133] and within the ducts of the HVAC system [157]. The Guide does have a general standard for the macro-environment of terrestrial species of 30%–70% humidity, but higher levels might be found within static FT cages. Even with RH levels of 44%–50%, static FT cage RH levels are routinely 70%–74% [16, 110, 119]. In general, ammonia production is inhibited by RH < 50% [110, 158], and IVC offer the advantage of tying micro-environment RH more closely with the room. The primary concern for ammonia as a research variable is its effects on the respiratory system [114]. Although these effects are well documented in humans and animals, the precise levels having specific biological effect for animals, and in particular rodents, are not well established. Short-term human exposure to high levels causes irritation to the upper and lower respiratory system and ocular tissues, and chronic exposure impairs respiratory function [114]. These effects have been duplicated in multiple animal species, and the available literature supports maintaining agricultural species below the thresholds recommended for humans; no evidence is available suggesting that airborne ammonia levels <25 ppm will affect research with larger species. The research on expected ammonia levels and how these levels affect rodents is more variable than in large animals, but enough research has been done to support considering the potential impacts of ammonia for rodents. Airborne ammonia could interfere with studies in which nasal histology is part of the evaluation or in infectious disease studies of the respiratory tract [159]. Respiratory damage is the only effect currently considered a significant risk for humans exposed to airborne ammonia, and research on impacts to other systems is not sufficient to make definitive conclusions [114]. Blood ammonia concentrations have a high capacity to regulate against inhaled environmental ammonia concentrations, and therefore systemic physiologic effects are expected to be limited if present [160, 161]. Histopathological evaluations on other organs have not demonstrated consistent effects from ammonia, particularly of clinical importance or at levels to which animals would be exposed in a normal housing environment. Studies have reported either no effects or have failed to find effects that could be repeated and reasonably attributed to ammonia exposure, unless given at near lethal levels [109, 114, 162–164]. A possible exception is corneal lesions as they have been suspected as being caused by ammonia in mice, with wide variations in strains susceptibility [165], and ammonia has caused corneal lesions in nonrodent species [114, 163]. Limited research suggests a possible effect of ammonia on the immune system. Ammonia levels of 90 ppm have been shown to suppress cell-mediated immunity in guinea pigs [166], although typical levels in guinea pig FT cages is not well established and they are often kept in open top cages where ammonia levels would be much lower. Exposure of mice to 86 ppm ammonia affected cytokine production and lymphocyte signaling [167]. In rodents chronically exposed (~3 weeks) to ammonia at consistent levels that commonly occur in the micro-environment (ie, 25–100 ppm), there was enhanced incidence and severity of respiratory infections [109], impairment of mucociliary activity [168], and altered hepatic microsomal enzyme activity [169]. No differences in immune function in mice were noted when increasing cage change from 7 to 14 days for a 5-month period in ventilated cages, although ammonia levels were not reported for this part of the study so correlations remain difficult to determine [155]. Chickens exposed to 30 ppm had no immune system effects even after 45 weeks [170], but higher doses of 70 ppm were demonstrated to affect the immune system [171]. More research is needed to determine whether ammonia levels pose a practical risk or research variable for immune parameters. The dichotomy between experimental and “real-world” findings confounded by protections for humans and an extensive body of in vitro work demonstrating ammonia to have immunosuppressive properties has been a vexation of animal care programs for years. The studies specific to rodents show wide variability in ammonia levels within routine housing conditions. With respect to the typical cage, the ammonia concentration is 0 ppm immediately after cage change, begins to increase exponentially sometime thereafter (as influenced by cage type and environmental factors), often then plateaus, and then drops back to zero subsequent to the next cage service. Among any group of cages, there will be great variation in the rate of increase and peak measurement of ammonia at any point in time. Individual cage ventilation is the most obvious method to decrease ammonia levels [16]. Ventilated cages can be easily kept below a peak of 25 ppm even at longer cage change intervals, depending on air change frequency [15, 97, 103, 155] and as mitigated by low RH [119]. However, more surprising is the large range in ammonia levels in static cages from <25 ppm to >400 ppm by day 7, with 1 report even up to 700 ppm ammonia; the potential contributing factors to this variability were discussed above in the Overview [16, 98, 101, 103, 110, 119, 120, 172]. Nasal lesions and increased susceptibility to respiratory pathogens have been shown in rodents at levels reported in normal housing environments. Ammonia has been associated with inflammation, necrosis, and epithelial changes in both rats and mice. Historical papers have found nasal lesions in rats even at mean concentrations of 150 ppm with natural exposure and 250 ppm of experimental exposure [109], and levels as low as 100–200 ppm have been associated with tracheal lesions in rats [154]. Experimental exposure of mice and rats to 300 ppm ammonia has also resulted in nasal lesions [159, 173], and 1 study in mice suggested even levels of 93 ppm are associated with nasal necrosis and 180 ppm with rhinitis [116]. Studies with natural exposure to variable ammonia levels have found conflicting results. One study found increasing severity of nasal lesions in mice from 25 to >400 ppm ammonia [104], which is consistent with another study that found nasal lesions in mice housed in cages that reached 200 ppm ammonia [174], while other studies found no nasal pathology at similar levels [105, 117]. High individual variability and unknown factors other than ammonia contributing to nasal pathology could be leading to conflicting results, as 1 study found rhinitis in 15%–30% of mice exposed to an average of 2 ppm ammonia and 10%–25% of mice exposed to an average of 240–350 ppm [155], and mice exposed to relatively low levels of 25 ppm had mild lesions in another study [104]. Strain variability in nasal pathology response to ammonia exposure could also be a factor [175]. In addition, study duration may play a role since nasal pathology associated with ammonia exposure might be attributable to chronic effects that worsen with prolonged exposure. In addition to nasal lesions, ammonia levels can increase the severity of experimental respiratory infections in a dose-dependent fashion from 25 to 250 ppm in rats with Mycoplasma pulmonis [109, 176], and ammonia levels of 500 ppm exacerbate Pasteurella multocida infections in mice [177]. Fewer studies have evaluated the pulmonary effects of exposure to ammonia, but none have been found with normal exposure levels [98]. None of the studies noting abnormalities in rodents with ammonia exposure have reported any notable clinical signs or abnormal behavioral changes even with the highest ammonia levels encountered under practical conditions. The lethal concentration 50 (LC50) for rats is 16 000 ppm at 60 minutes of exposure [178]. Very limited information is available on the effect of ammonia on behavior, with 1 study suggesting mice have no aversion to ammonia even up to 110 ppm [179] and 1 study showing that ammonia at 300 ppm decreasing running wheel activity in mice and for rats both at 100 ppm and 300 ppm with much greater reduction at 300 ppm [180]. However, changes in activity might not be appreciable on casual observation. Traditional agricultural species have similar and variable respiratory effects from ammonia as rodents, although they may be more prone to effects at lower concentrations than rodents. Pigs have no adverse effects of respiratory disease or lesions in the upper or lower respiratory system with ammonia levels <75 ppm [181, 182], whereas other studies have found lesions in the nasal turbinates in pigs from 25 to 100 ppm [183, 184]. Ammonia levels of 50 ppm exacerbate the pathology of experimental P. multocida infections [185] and increase the incidence of P. multocida and Mycoplasma hyopneumoniae in pigs in a commercial population [186]. Chickens have been shown to have adverse responses to environmental ammonia concentrations as low as 25 ppm [187], and 70–75 ppm has repeatedly been shown to reduce weight gain [171, 188–191]. Carnivores, livestock, rabbits, and other species larger than rodents kept in biomedical animal research facilities are unlikely to be exposed to ammonia concentrations >25 ppm where the conditions must meet OSHA indoor air–quality standards for human industrial workers [157]. Mouse intracage carbon dioxide levels range from 1000 to 6000 ppm in static FT cages [16, 101, 110, 119, 192], 900 to 2000 ppm in open cages, [16, 101, 193] and <5000 ppm in IVC [16, 101, 117, 155]. These values are significantly greater than the macro-environmental carbon dioxide level of approximately 400 ppm. In contrast to ammonia, CO2 levels measured in the macro-environment are generally below the threshold for human exposure, and no research has shown experimental effects on rodents exposed to naturally occurring levels. The Threshold Limit Value [95] for continuous safe human exposure to carbon dioxide for 40 h/wk has been established at 5000 ppm with a maximal 30 000 ppm safe exposure for 15 minutes [93, 95]. However, 1 study did find high carbon dioxide levels routinely >8000 ppm and individual cages even up to 20 000 ppm [120]. Not surprisingly, that study also had the highest reported ammonia levels naturally occurring in mouse cages [120], so carbon dioxide could be worth measuring across types of cages, particularly nonventilated versions, as an indirect indicator of adequacy of ventilation. The effect of chronic exposure to carbon dioxide concentrations in excess of 400 ppm on rodent biological processes and the implication for research results is not known. In nature, tunneling rodents may be acclimated to ambient carbon dioxide concentrations of up to 14 000 ppm [194]. Rodents exposed to 30 000 ppm CO2 have been shown to have increased corticosterone levels consistent with distress [192]. However, true aversive effects for rats and mice are in excess of a threefold increase of that concentration [195] and unlikely to be encountered in the micro-environment. Poultry are highly tolerant of even higher concentrations [196, 197]. Carbon dioxide is a progressive respiratory stimulant at concentrations up to at least 100 000 ppm [198]. At chronic carbon dioxide exposures of 6000 ppm or less, such as those found routinely in mouse cages, the risk of asphyxiation from oxygen displacement is nonexistent, but any effect on mouse physiology is not clear. In the case of power failures for ventilated caging systems, carbon dioxide levels in mouse cages may increase to 80 000 ppm in 1–2 hours [192, 199]. In a controlled power study of a rat ventilated caging system, oxygen levels dropped to lethal levels of <10% in 30–60 minutes in cages with dams and nursing pups, [200] making prompt detection of power failures essential and connection to back-up power highly desirable. Contact Bedding Laboratory animal beddings vary in material and shape, creating the potential to add variability to animal models, with a potential for a range of effects on reproduction, behavior, physiology, dietary studies, the microbiome, and exposure to undesirable microorganisms. These effects could be particularly profound for species that ingest the bedding, such as rats and mice. No specific type of bedding is ideal for all species under all management or experimental conditions, but optimally contact bedding should be absorbent, free of toxic or aromatic chemicals, safe, nonabrasive, disposable, comfortable, nonstaining, inexpensive, and readily available, but not readily eaten [18]. The effects of contact bedding on rodents are the focus of this article as little information is available for larger animals. Corncob-, wood-, and cellulose-based products are the most commonly used in current facilities for rodents [108], with contact bedding less commonly used for larger animals. Corncob is typically used at one-eighth- and one-fourth-inch sizes, with smaller particles, pelleted form, and mixtures also available. Cellulose products range from pure cellulose to recycled newspaper with a wide range in shapes, including shreds, rolls, square chips, and pellets. Hardwood beddings are available in chips and shavings as single-tree source options, such as aspen or maple, or mixes of multiple hardwood species that could also include beech, birch, and poplar. Other materials that have been evaluated for use in rodents but without sufficient evidence to advocate as a contact bedding include rice and wheat straw, [201] perlite [202], rice hulls [203], and synthetic materials [204]. Rodent preferences for bedding cannot be disentangled from enrichment, which is discussed elsewhere within this publication and was reviewed previously [99]. Mice prefer softer materials and those that allow burrowing and nesting, such as paper-based beddings, and both rats and mice prefer larger particles and fibrous materials relative to small particles [205–211]. However, bedding material can be supplemented with enrichment that supports these activities, and these preferences might not be as strong given additional enrichment options. Volume of bedding is a factor mitigating warm adaptation [212]. Impacts of Contact Bedding. Contact bedding, in general, offers many advantages for animal comfort, and some, such as corncob, virgin cellulose, and eucalyptus pulp, inhibit ammonia production [105, 110, 213]. Softwoods such as pine have been used historically and are still sometimes used following heat treatment that minimizes its impact on liver function [99]. Pine and cedar have long been recognized as affecting drug metabolism in rodents by inducing cytochrome P450 enzymes with effects that can last for weeks, even after removing the animal from the bedding [169, 214, 215]. Softwood beddings have also been shown to induce rat liver endocytosis [216] and possibly affect other liver enzymes as well [201]. These effects have not been found in rabbits exposed to softwood bedding, although the rabbits did not have direct contact, and bedding treatment was not described in this study [217]. Contact beddings made from heat-treated hardwoods such as maple, birch, beech, aspen, or poplar may have residual vitamin C, which can confound studies of scurvy using guinea pigs [218]. Bedding materials are a source of microorganisms and are often sterilized to reduce the microbial burden by either irradiation or autoclaving. Multiple types of unsterilized bedding have been found as a source of fungal spores, bacteria, and nonpathogenic nematodes [103, 219–222]. These agents may be ingested or aerosolized, presenting a risk of mucosal contact or inhalation, depending on the bedding’s characteristics such as dustiness [220], and can be influenced by the caging system utilized. Although even the same type of bedding can vary widely in bacterial content and no beddings are sterile after normal manufacturing processes, paper beddings are reported to have lower bacterial content and endotoxin levels compared with corncob and wood [220, 221]. Autoclaving not only reduces the microorganism loads to undetectable levels but also may be associated with ammonia suppression within the cage [103, 219, 220]. Although extensive analysis of the bacterial flora associated with specific bedding types is limited and likely highly variable depending on material and vendor processing, the primary bacterial flora identified in corncob bedding is considered normal flora for immunocompetent mice [103]. The risks of unsterilized bedding should still be considered given that it has been associated with fungal rhinitis in rats [219], and feed and bedding in shipping containers have been implicated as a source of rotavirus infection in mice [223]. The presence of endotoxins and bacteria has been suggested as a potential confounder for respiratory-related research, and endotoxins are not as sensitive to autoclaving as bacteria [103, 221, 224]. Bedding can affect the microbiome and immune system of rodents, likely both by direct exposure to microorganisms but also because of the substrate provided to bacteria from bedding ingestion. Although bedding effects cannot be definitively attributed to direct exposure vs ingestion, mice and rats will eat at least some bedding materials, and impacts are likely direct results of this ingestion [225]. Bedding type has influenced dietary studies in mice and [226] the outcome of periodontal studies and disease in rats [227, 228], and it has had variable effects on rat body weight [229, 230]. The effect of microbial exposure vs ingestion of the bedding material can be difficult to distinguish if scientific studies do not specifically report whether the bedding was sterilized. Extensive studies have shown that bedding affects gut microbiota [225, 231–234], which is only briefly addressed here as it is the subject of another article in this issue. Bedding influences on the microbiome and immune system are likely dependent on direct exposure to the bedding, as the impact is reduced if rodents are housed on a grid floor suspended over the bedding [225]. Bedding can specifically affect intestinal immune responses in mice. In a comparison of irradiated wood vs cotton bedding, the former was associated with the proliferation of Peyer’s patches and increased total and virus-specific Peyer’s patches and mesenteric lymph node IgA responses in rotavirus-challenged mice [235]. The authors speculated this effect could be attributed to immunostimulatory properties in hardwoods or to the bedding material directly affecting microflora. The primary concern related to the direct effect of bedding on rodent behavior is the potential of estrogen-like compounds in corncob bedding. Phytoestrogen and xenoestrogen exposure can result from diet as well as other husbandry-related sources. Reviews are available on general sources of estrogen-like compounds [236] and specifically from corncob bedding [237]. Corn and corncob bedding have been found to contain an estrogenic mycotoxin not present in other common bedding materials [236, 238]. This mycotoxin was attributed as the cause for shortened time to vaginal opening in mice housed on corncob, and experimental administration of this mycotoxin increased mouse uterine weight and affected reproduction [236, 239]. A corn mitogen unrelated to a mycotoxin was then discovered in corncob bedding that was not reduced by autoclaving [240, 241]. This mitogen was found to disrupt endocrine function [242], affect male sexual behavior [243], female sexual behavior [240], and estrous cycles [241] in rats when administered orally in a controlled study. Male nude mice implanted with human prostate cancer xenografts have accelerated tumor growth when housed on corncob as opposed to cellulose bedding [240]. Corncob bedding relative to wood bedding has also been shown to affect social behavior and aggression in Peromyscus californicus [244, 245]. In contrast to these previous studies, Jones found that housing on corncob bedding had variable inhibition on ovariectomized rat female sexual behaviors depending on the hormones they received and that even when female inhibition was noted it could be overcome by cues from male rats [246]. Jones proposed that the contradiction was possibly because of differing hormone doses or underappreciated aspects of handling and husbandry, although the rat strain was also different than previously reported [246]. Many rodent breeding colonies have been successfully maintained on corncob bedding, so the practical aspects of these effects are likely minimal. In addition, commercial rodent diets typically include corn as part of the diet, so eliminating exposure to all corn products would be challenging. However, the potential effects of corncob bedding on more sensitive behavioral and hormonal studies should be considered given the aforementioned findings. Although the effects of bedding on other aspects of rodent behavior are limited, research suggests possible effects. The use of contact bedding is well established for rodents, with a lack of access to bedding providing the basis for an established model of neonatal stress in rats and mice [247–249]. The absence of bedding has affected open field activity in rats [229], but the effects of any 1 particular bedding type are less clear. One study found no difference in wood chips vs shavings in mice on anxiety, depression, or social behaviors, even though breeding success was better on shavings [211], and conversely another study found no difference between shavings and chips for mice in breeding outcomes; however, the 2 bedding types did differ in tests of memory, activity, and anxiety, particularly in males [250]. Rats housed on corncob bedding had less anxiety-like behaviors relative to animals raised on wood pulp, independent of levels of maternal care [251]. Rats spent less time in slow wave sleep when housed on corncob as opposed to aspen chip bedding [252]. Mice housed on a mix of shredded aspen and cellulose displayed excessive grooming behaviors relative to mice housed only on aspen or cellulose [230]. In 1 study, aspen shavings vs chips even led to increased breeding fecundity, but this was associated with improved nests, so the possibility exists that the effect was from enhanced nest-building [211]. In a study comparing wood flakes with chips, there was no difference on fecundity [250]. Bedding type has also been shown to affect responsiveness to a noxious needle assay as well as heat sensitivity and temperature preferences in mice [206] and to directly affect the progression of neuropathic pain models in rats [210]. Because the bedding rats are raised on can affect preferences [253], the impact neonatal bedding exposure could have on behavior studies in adults could be under-appreciated. Researchers should be aware that bedding varies in size, shape, and material and could have differing effects on behavior. Few studies have evaluated the direct impact of bedding on rodent health. In rats, aspen shavings have been associated with higher rates of sneezing irrespective of ammonia levels relative to paper bedding, suggesting that differences in dust levels could be relevant [254]. Particulates from cotton enrichment can be a source of conjunctivitis and blepharitis in nude mice [99, 255]. Although there is very limited research evaluating effects on the lower respiratory system, at least 1 study found no difference in oxygen consumption when comparing cedar, pine, and natural fiber bedding in mice [256]. Paper, wood, and corncob bedding have been used successfully for many years in laboratory rodents, and generally the concerns regarding bedding principally relate to subtle research effects as described rather than posing a direct health hazard. Experimenters should be aware that it is likely that the wide variety in bedding material texture and size could affect various aspects of behavior directly or indirectly from health effects. Barometric Pressure Biological effects may be altered in animals as a consequence of changes in barometric pressure associated with local weather patterns, in transportation (particularly by air freight), and when relocated between low- and high-altitude sites. Air pressure is lower at high altitude; for example, at Fort Collins, Colorado, at 1550m above sea level, it is 83% of the value of a coastal location [257] and can drop up to 5% in any locale in the advance of stormy weather. Decreases in barometric pressure in the range associated with advancing storms aggravate neuropathic pain in animal models [258, 259] and influence behavior [260, 261]. Humans who are not acclimated are at risk of altitude sickness when rapidly ascending to high elevation where inspired oxygen concentrations and partial pressure of oxygen in the bloodstream substantially decrease [262]. Acute altitude sickness effects and signs have not been documented in nonadapted research animals but should be presumed. Studies using simulated atmospheric pressures suggest that it may be a variable in hemodynamic experiments where the animal subjects are reared at one elevation and then transferred without sufficient acclimation for study at a different altitude [263]. Changes in air pressure are measured in a variety of units standardized to sea level using a barometer. An atmosphere is a unit of measurement equal to the average air pressure at sea level at a temperature of 15°C. One atmosphere is equivalent to 1013 millibars, 101.3 kilopascals, 760 mm of mercury, 29.9 inches Hg, or 14.7 psi. Impacts of Barometric Pressure. In light of potential influence on pain, behavioral, and cardiopulmonary studies, investigators are advised to record and take into account barometric pressure changes as an independent variable possibly influencing biological processes, as a risk of using historical controls, and in stabilizing newly acquired animals in advance of experimentation. In a study of adaptation of rats to hypobaric and hypoxic conditions, allostasis of respiratory-associated metabolism required 7–14 days [264]. Electrical and Magnetic Fields Other than in imaging and medical treatment and encounters with airport security systems, biological entities encounter magnetic and electromagnetic fields naturally and unavoidably as a consequence of living on earth, when in proximity to sources and transmitters of electrical power, and in association with communications devices and systems. Some animals have been shown to use the direction and intensity of the earth’s magnetic field as an aid to orientation and navigation. The perception of magnetic fields, so far, has been demonstrated to be used more prominently by nonmammals, particularly migratory birds, but also some fishes, reptiles, amphibians, and invertebrates [265]. Among mammals, bats, and naked mole rats have demonstrated magnetic compass orientation [265]. The phenomenon of magneto-reception is retained in captivity by migratory birds, sedentary chickens, and zebra finches, all of which are occasionally used in research [265]. The visual system is integrally involved with magneto-reception influenced by light intensity and wavelength in wild birds [265]. The strength of a magnetic field is measured using a magnetometer in units of Tesla or microTesla and also Gauss or milliGauss, where 1 G is equivalent to 10−4 T (or 1 milliGauss = 0.1 microTesla). Images of magnetic fields can be recorded using flux-detector film. The direction of a magnetic field also can be determined using a compass. Impacts of Electric and Magnetic Fields. The overwhelming preponderance of what is known about magneto-reception is in the study of migration [266], hypo-magnetism in models of space travel [267], and medicinal effects of static magnetic fields [268]. Given the fact that only a few species have been studied in detail, and none that are used in abundance within the confines of a vivarium, the biological effects of natural magnetic fields on common research animals, if any exist, remain conjectural and require more research. Unless expensively shielded, there is no distinction between macro- and micro-environment with respect to exposure to the earth’s magnetic field because there are no common building materials that block it; however, Faraday shielding can be used to stop electrical and radio waves. The benign neglect accorded to the presence of magnetic fields that are not sensed by humans, but may be perceived by research animals, gives pause to wonder if geographic location and the arrangement of equipment relative to magnetic field direction, a factor resistant to geographical standardization across, might be a source of the difficulties chronicled in replicating behavioral experiments [269–271]. In pursuit of improved reproducibility through the most thorough description of methods [272], investigators concerned about magnetic fields as a variable in their studies are advised to record the GPS coordinates of study locations and describe the orientation of phenotyping devices, particularly those that record locomotion and positioning, relative to compass bearings. Transportation One setting where the potential exists for rapid macro-environmental-coupled-to-micro-environmental change, risking perturbation (or worse) of the micro-environment, is in the process of transportation. Whether it be the domestic or international shipment of experimentally naïve animals from a producer to a research destination, as a matter of exchange between research institutions worldwide or regional travel back and forth involving a unique modality, to small trips within the confines of an institution between the housing site, a laboratory, or a specialized service center, transportation can perturb results. Transportation across broad distances may take 1 or more days [273], is an unquestioned stressor of animals, is difficult to precisely measure, affects a broad array of physiologic processes [274], and requires a period of adaptation or of restoration to homeostasis [18]. The associated variation in handling; mixing of unfamiliar animals at high density; novel environs and the micro-environment of the shipping container; provision of food and water possibly by different means to those accustomed; differences in macro-environment between conveyances and depots, and associated shipping-related stresses; barometric, electromagnetic, time zone, and thermohygromic stresses; and other factors all can be impactful and influence the conduct, quality, and veracity of experiments. These steps and processes are in the hands of purveyors and stations, often multiple, along the journey. Air transportation in North America has been shown to expose animals to temperature extremes either above [273] or below the acceptable range [273, 274] or with >10°C ranges during transport [273] over 60% of the time. Flights with stopovers, commercial flights (rather than dedicated couriers), and international shipments were most likely to experience temperature extremes and large variations in temperature [273]. An additional variable associated with long-distance shipments is unforeseen weather that can worsen extremes, delay shipments, and prolong transport time. Not even trips of only a few hours or even room-to-room transport within a facility or transfer to a different cage type are free of the likelihood of disequilibrium [275–277]. Impacts of Transportation. Historical data have suggested that at least 48–72 hours are needed for most laboratory animals to return to normal physiological and immune function following the stress of shipping [278–285]; therefore, many research institutions recommend this period for acclimation. This metric has been challenged in some cases. Acclimation periods of at least 3 weeks have been recommended when measuring blood pressure or markers of stress in mice [286, 287] and, in some cases, up to 4 weeks may be required [288] even after short traverses of only a few hours [289]. Although a default acclimation period of 72 hours after receipt of animals may be appropriate in many cases before starting an experiment, model systems unique to different research purposes and variables of species, gender, age, and shipping characteristics dictate that investigators know the key homeostatic attributes of their model system and that acclimation periods are sufficient to ensure they are achieved. The time to achieve stability after transportation can be reduced through the application of the common sense principles of availing dedicated, expert couriers using climate-controlled vehicles to transport animals; specifying ground transportation over air and, where the latter is the only choice, direct flights; avoiding flights during periods of temperature extremes, eg, mid-summer or mid-winter; shipping at night during hot weather; providing nesting material in shipping crates; and when crafting shipment orders to specify aircraft cargo-hold temperatures that are suitable for the species [273]. ENVIRONMENTAL VARIABLES AND JOHAIR WINDOW KNOWNS AND UNKNOWNS Designing and executing animal experiments is a difficult and resource-intensive endeavor. The onus of the NIH, NC3Rs, and ACLAM positions and guidelines add another layer of complexity, but, as we have explored throughout this manuscript, documenting them is critical to achieving experimental reproducibility and can have major implications on the state of animal models. On a pragmatic level, investigators will find it helpful to view the myriad of environmental factors through the lens of the Johair window: environmental parameters can be assigned a known or unknown state to self and others [290]. Known to Self-known to Others Well-known critical and/or regulatory mandated environmental parameters are best measured, controlled, and reported to ensure rigor and transparency are brought to each animal experiment [5, 7, 9, 13]. These are well defined by the ARRIVE guidelines [7]. Known to Self to Others Investigators sometimes unveil important environmental factors that influence the pathophysiology of animal models and/or welfare but were heretofore unknown or underappreciated by the larger community. A recent example is the influence of ambient temperatures on tumor immune-micro-environments with profound implications on therapeutic response to check-point inhibitors [81]. In this case, responsibility lies with the investigator to share this knowledge with their respective field (eg, by publication) and with the institutional laboratory animal group [5]. Unknown to Self-known to Others Many environmental parameters are well documented per regulatory and animal welfare mandates [18, 291] and by institutional laboratory animal and facility departments. Investigators, laboratory animal specialists, and facilities operations units should freely share and report this data in the spirit of ACLAM’s position statement [9] and NIH’s transparency stance [5]. Unknown to Self-unknown to Others The existence of unknown unknowns is especially vexing. New data may emerge that highlights an unappreciated environmental parameter that puts into question experiments already completed. In recent years, this includes vibration [3], availability of nesting materials [57], and drafty ventilated caging [58]. In the face of unknown unknowns, confirmatory studies reproducing the results of another group, preferably at another institution, becomes the strategy of default. Should environmental parameters be uncovered as a source of variability, data should be shared across institutions as encouraged by NIH’s transparency effort [5] to encourage refinement and enhance reproducibility of animal models. Another strategy is to introduce controlled variation a priori as an opportunity to uncover new facets of contextual biology [4, 292–294]. This approach is analogous to efforts to expand the population enrolled in clinical trials [295]. Mo et al [292] speculated this approach offers that “opportunities for clinical translation will be guided by the ‘environmental construct validity’ of the preclinical data” and could be generalized by adding systemically included environmental context variables (eg, with and without enrichments) to animal models, especially if combined with advanced statistical nonlinear mixed-effects modeling to interrogate multiple factors [296]. Conclusions Numerous environmental variables can have profound effects on biological responses of research animals beginning before zygote formation and extending on a continuum to death. Throughout this progression, the environment can influence the organism, its biological processes, and experiments. Some environmental factors remain unpredictable in effect and any number, not known or granted great weight now, could become significant. In some cases, variables associated with the environment may be difficult to standardize geographically and seasonally. Given that the quality and consistency of the environment and its myriad components are critical elements in research, it has been a continuing charge extending back to the nascence of research animal care and support programs 100+ years ago to focus on it. This was understood in 1959 by W.M.S. Russell and R.L. Burch in their observation that “physiological uniformity is likely to be one of the great rewards of good husbandry”[2]. The conduct of valid and reproducible experiments requires the management of any variable that can be identified, measured, and recorded, where applicable, and controlled to promote the animal existing in harmony with its environment. Although some have criticized this traditional allegiance to standardization in the care and use of experimental animals, [4] it remains the seminal charge of the animal care program up until the time the circumstances of a specific experiment warrant otherwise. It is imperative that animal care programs understand that they do more than husbandry. The best ones understand that they participate in every experiment from the time of conception or receipt and first housing of the animals on through the life of the study. They do this by providing a consistent and wholesome environment where the animals reside, experiments are run, and data are collected. This begins with and is made possible by a well-designed facility. Likewise, given that facility design and operations do not homogenize to 1 national or international standard in terms of animal care, it is important for investigators to know and understand the resiliency, dependence, or sensitivity of their model systems to effects of temperature; moisture in the environment; and other environmental factors in the vivarium, in their laboratories, or other study areas; in transportation; or when relocating to another institution. Correspondingly, those who care for the animals and those who use them should collaborate to identify the degree of deviations in environmental conditions that are significant so these can be recorded in consideration of the interpretation of any data and their reporting along the lines of the ARRIVE guidelines [7]. Through understanding these factors and concepts and with those involved in animal care and use integrated as an informed team, the conduct of valid reproducible experiments, the generation of new knowledge, and improvements in human and animal health are likely to be the benefit. Acknowledgments We thank the reviewers and editors for the careful attention devoted to our manuscript. The American Conference of Governmental Industrial Hygienists, the Copyright Clearance Center, and National Academy of Sciences of the United States graciously granted use of copyrighted material. Emory University paid all copyright and color image fees. Potential conflicts of interest. 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TI - Micro- and Macroenvironmental Conditions and Stability of Terrestrial Models
JF - ILAR Journal
DO - 10.1093/ilar/ilaa013
DA - 2019-12-31
UR - https://www.deepdyve.com/lp/oxford-university-press/micro-and-macroenvironmental-conditions-and-stability-of-terrestrial-zJjrJWITx2
SP - 120
EP - 140
VL - 60
IS - 2
DP - DeepDyve
ER -