Lighting

Appropriate lighting is crucial to an animal’s physiology and behavior (Dauchy et al. 2013a, 2013b), and alterations in light cycle, intensity, or spectrum may lead to stress (Stoskopf 1983). In addition to duration, intensity, and wavelength, the species and strain of the animals are also factors to consider (NRC 2011) for the macroenvironment.

Lighting should be diffused evenly throughout the macroenvironment to provide for the animal’s specific lighting requirements and provide sufficient light to perform husbandry, sanitation, and animal health evaluation procedures. Lighting should be at appropriate levels regardless of the animal’s location in the room (NRC 2011). In the United States, most animal research facilities follow the recommendation of 325 lux (30 foot-candles) approximately 1 m (3.3 ft) above the floor for routine animal housing (NRC 2011). This lux level has been deemed appropriate for accommodating husbandry, veterinary, and research activities without causing retinal damage to albino rodents (Schofield and Brown 1996; Faith and Huerkamp 2009). European guidelines further stipulate that darker areas for withdrawal should be available within the animal enclosures (Council of Europe 2006).

To control light intensity and provide appropriate lighting for a variety of species, most modern animal research facilities are designed to include two-stage lighting systems. With two-stage lighting, lux can be set at a low level to accommodate albino rodents or at higher levels for animals with normally pigmented eyes. The lighting can also be adjusted for specific research needs or to accommodate husbandry activities. Some of the more sophisticated systems that control two-stage lighting are computer based and can be administered from the desktop or by remote access by way of a variety of mobile devices. Preprogrammed illumination override switches are often available at the room level for short-term use. These systems help to eliminate the possibility of unauthorized room-level alteration of light intensity and may even generate alarms if lighting parameters are out of prescribed ranges.

Research facilities are designed to provide a controlled light environment that prevents outside environmental influences, such as natural lighting, from creating variances from programmed photoperiods. Light cycles should ensure that animal health is not compromised and should provide for normal diurnal and circadian rhythms. Light cycles that provide 12–14 hours of light daily are appropriate for most laboratory animals (Lipman 2007; Pritchett-Corning et al. 2011).

Since most human activity in a research facility normally occurs during the day, the most widely used photoperiod in animal research facilities is illumination during the day and dark at night. During this cycle, animal holding room lights are on during the vivarium’s normal business hours and at an acceptable illumination level to permit routine husbandry, veterinary care, and research activities. However, in some situations researchers choose reverse light cycles, where lights are off during the day and on during the night. Since rodents are nocturnal animals, their highest level of activity is during the day when housed under a reverse light cycle. This type of photoperiod is convenient for researchers to observe nocturnal activities and behavior during daytime hours, but presents challenges for the daily operation of the animal facility. Red spectrum lighting and night vision goggles can alleviate some of the human vision challenges and allow staff to safely carry on with daily activities. Red spectrum lighting works because of a difference in the cells that make up the retina of the rodent and human eye (Faith and Huerkamp 2009; Pritchett-Corning et al. 2011; NIGMS 2012); rodents cannot perceive red light, while humans can see this wavelength of light. Wavelength filtering materials, such as commercially available red film over light sources, will provide a perceived dark environment for rodents, but humans can continue daily operations in the red light illuminated environment. Red wavelength illumination can be generated by door window tinting, fluorescent bulb sleeves, or tube guards. Although red spectrum lighting is widely recognized for allowing a safe working environment for humans, emerging literature points to evidence that continuous exposure to red spectrum illumination is not without negative implications for laboratory rodents. A recent study showed evidence of marked changes in circadian hormone rhythms and plasma corticosterone in nude rats that were housed in red-tinted cages (Dauchy et al. 2013b). As an alternative to using red spectrum lighting, Faith and Huerkamp described the benefits of sodium (vapor) lamps, which have wavelengths at the margins of what rodents see, but in the visual field of humans. They provide sufficient light levels for humans but dull the light for rodents to a level that allows nocturnal behavior (Faith and Huerkamp 2009).

Advanced electronic lighting systems, such as the Lutron System™ (Lutron Lighting Systems, Inc., Cooperstown, Pennsylvania), are capable of phototransition. This is the gradual change in a photoperiod by slowly increasing or decreasing light levels, creating a dusk-to-dawn effect instead of a sudden change in illumination. Sudden changes in illumination intensity or light cycle can elicit a startle response in animals. For example, birds not accustomed to programmed photoperiods may take flight and collide with walls or other obstacles when unexpectedly subjected to darkness (Faith and Huerkamp 2009). Some aquatic species require transitional lighting to stimulate feeding, cleaning, and comfort behaviors (Stoskopf 1983).

It is widely recognized that appropriate light cycles are extremely important for maintaining natural circadian rhythm, which determines when the body should be awake or asleep. Deviations from natural photoperiods cause disruptions in circadian rhythm and can have detrimental effects on the brain, body function, and behavior (Lipman 2007). When mice were subjected to a 20-hour photoperiod (10 hours light and 10 hours dark) for 6–8 weeks, they exhibited profound changes in cognition and physiology, increased body temperature, disruption of normal hormone levels, and increased weight gain (Karatsoreos 2009). Other studies have shown that photoperiod changes resulted in behavioral alterations (e.g., increased aggression), as well as changes in reproduction and increased susceptibility to cancer and infectious disease (Kennaway 2005; Dauchy et al. 2011). Since unscheduled disruptions in photoperiods can be devastating to research outcomes, lighting should be electronically controlled and regularly monitored to ensure that cycles remain consistent. If cycle timers fail or override switches are not functioning properly, lights can remain on or off for long periods of time outside the scheduled photoperiod. The lighting parameters should be controlled and documented by the building environmental monitoring system and alarm if room parameters deviate outside of a specified range. During the light “on” period, task lighting in rodent rooms should be provided by override for a time-limited period before returning to the programmed lighting levels and schedule. During the lighting “off” period, the room lighting should be provided via override in the red spectrum for a time-limited period. Care should also be taken to eliminate disruption caused by light contamination during the dark period, caused by light leaking around and under door frames or unprotected windows. Digital displays on ventilated rack blowers or other equipment maintained in the animal room may also emit enough light to disrupt a dark cycle (Dauchy et al. 2011).

Historically, animal research facilities have used overhead fluorescent lighting fixtures to provide illumination in corridors, as well as in animal holding rooms. Generally, these are ballast–lamp systems holding cool white fluorescent (CWF) tubes that emit sufficient light levels in an appropriate wavelength to enable the human eye to adequately evaluate and work with the animals. In addition, most CWF fixtures produce the wavelength and spectral pattern of natural light (Hessler 2009). Fluorescent fixtures are also available in a wide variety of colors, and there is growing research interest in evaluating the effects of different light spectrums on animal physiology. In addition to color variety, the environmental impact of lighting fixtures is now a consideration in vivarium lighting. One of the newest environmentally friendly concepts is the introduction of light-emitting diodes (LEDs), which, at appropriate levels, have been shown not to interfere with circadian rhythms or cause phototoxic effects (Hessler 2009). Although LED lighting is usually more expensive initially, it consumes less power and operates longer than traditional light fixtures. Another option for room lighting is augmentation with natural lighting through windows or sky lights. This is infrequently recommended in animal research facilities because of the inability to strictly control photoperiods, but can be part of an environmental enrichment program for higher-level species, such as nonhuman primates or farm animals housed in research facilities.

Heating, Ventilation, and Air-Conditioning

Ensuring consistent, species-specific temperature and humidity in an animal room is critical for animal thermoregulation, to maintain normal physiology, and to promote natural behavioral tendencies. Species-specific room-level, dry-bulb temperature tolerance ranges for common laboratory species are listed in the Guide for the Care and Use of Laboratory Animals (Guide) (NRC 2011). (The dry-bulb temperature is the temperature of the air measured by an ordinary thermometer.) Temperatures in the Guide for rodents are typically set lower than their thermoneutral zone (the temperature range where the animals do not need to gain or lose heat) to prevent heat stress during periods of activity. The addition of nesting material is important to keep mice warm when at rest. Since wide fluctuations in temperatures can be detrimental to animal health and well-being, temperatures should be maintained at a set point toward the middle of the range and within 2° of the set point. For example, if the desired temperature for a rodent room is 70°F, the room temperature should remain between 68°F and 72°F. Wide fluctuations in temperature can produce metabolic and behavioral responses that can confound research results or alter an animal’s response to research procedures (Hessler 2009). Temperature extremes can cause a reduction in breeding, a decrease in food intake, increased disease susceptibility, and disturbances in sleep–wake patterns (Laber and Gonder 2007; Lipman 2007; Faith and Huerkamp 2009). Macroenvironmental fluctuations and high rates of intracage ventilation can be mitigated at the cage level by providing appropriate nesting materials or nesting boxes that can aid in thermoregulation. These materials provide a choice for animals to build nests, take shelter in a box, or burrow into bedding in response to environmental conditions inside the cage (Baumans et al. 2002; Gaskill et al. 2009, 2012).

Relative humidity (RH) is another component of the macroenvironment that could have detrimental effects on animal health if not maintained within tolerable ranges. There is a wider range of control for humidity (30%–70%) than temperature, but extreme variations from recommended RH should be avoided (NRC 2011). Although these values are based on tolerable ranges for most mammalian species, maintaining consistency in RH is important for all animals used in biomedical research. Extreme fluctuations have been shown to impede an animal’s rate of heat loss, impact normal activity, and cause changes in normal amounts of food intake (Faith and Huerkamp 2009; Rosenbaum et al. 2010; NRC 2011). Certain species can develop dermatitis or flaky skin, and rats can develop ringtail from prolonged periods of exposure to humidity below 30% (Schofield and Brown 1996; NRC 2011). RH that is extremely high can also influence conditions in the microenvironment, such as increased moisture levels in the bedding, cage wall condensation, higher cage temperatures, food spoilage, and bacterial generation of ammonia (Burn and Mason 2005; NRC 2011).

Considering the ramifications of uncontrolled environmental conditions, temperature and humidity should be monitored and recorded on a regular basis. Equipment as simple as a room-level hygrothermometer or a sophisticated, facility-wide electronic environmental monitoring system can ensure that animals are not subjected to extended periods of extreme temperature and humidity fluctuations.

Air Quality

The quality of the air at the room level not only impacts animal health but also is an important component of creating a safe environment for personnel working in animal facilities. One of the most common occupational hazards for animal care staff or others working in animal facilities is the development of animal-related allergies or increased sensitivity to allergens (NRC 1997; Harrison 2001). The quality of the animal room air is affected by the particulate load of a variety of particles, such as animal dander and dust generated from animal bedding, by gases such as ammonia vapors and by infectious agents from both animals and humans. Most research institutions control the quality of room air by exchanging the entire volume of the air with 100% outside air at a set rate per hour, based on the type of caging or species housed within the room. The number of times the room air is replaced per hour is referred to as air changes per hour (ACH). Exchanging the volume of the room air can also aid in reducing heat load, decreasing RH, and replacing carbon dioxide generated by animals and humans with oxygen. Ten to fifteen constant volume ACH are recommended in most animal housing rooms, but these recommendations do not take into account the variations in heat load; the size, species, and numbers of animals; the room design; and the microenvironment (NRC 2011). Setting of ACH at the higher end of the suggested range may be unnecessary and a waste of energy. Conversely, at the low end, a room that is heavily populated or housing a significant number of large animals could be underventilated at 10–15 ACH, leading to increases in temperature, RH, particulate counts, and noxious odors. In order to ensure a higher level of air quality, modern facilities frequently consult with engineering specialists to include computational fluid dynamics (CFD) analysis when designing heating, ventilation, and air-conditioning (HVAC) systems for new construction. Computerized CFD software effectively evaluates how air will enter, circulate, and exit the room based on volume of air and dynamics created by the air entering the room, in conjunction with the placement of supply and exhaust registers (Norton and Brouwer 2009). This analysis utilizes various types and amounts of caging planned for each room, as well as variations in species. Obvious flaws will be identified, and adjustments can be made to provide the best possible macroenvironment for animals and humans.

Noise and Vibration

Noise and vibration have been shown to affect many behavioral and physiological parameters in animals and can be a confounding variable in research studies (Anthony and Harclerode 1959; Buckley and Smooker 1970; Turner et al. 2005, 2007; Small and Dietrich 2007). As a result, the current edition of the Guide has placed emphasis on taking steps to control the effects of these factors (NCR 2011). Although both sound and vibration can adversely affect animals, it is not a simple matter to determine what magnitudes and frequencies of sound or vibration are problematic. The information below summarizes the relevant scientific properties of these factors and what is currently known about the effects of sound or vibration on laboratory animals.

Sound and vibration are forms of energy that travel in waves, with sound being perceived by what we hear and vibration by what we feel. These waves have both amplitude and frequency (Figure 20.2). The amplitude indicates the intensity of the sound or vibration and is represented by how far the peak of the wave moves past the neutral position. The frequency is the amount of time that it takes to complete one cycle from a point on one wave to the same point on the next wave. The term hertz is used as a unit of measure for frequency and is the number of cycles per second. One hertz is one cycle per second (Crocker 2007). As discussed in the text below, both the amplitude and frequency have to be considered when determining the effects of sound or vibration on laboratory animals.

Figure 20.2

Depiction of sound or vibration wave demonstrating the amplitude (intensity) and wave cycle of sound or vibration.

Noise

The amplitude of sound can be measured in decibels. As a sound wave moves through the air, it creates pressure. The measurement of this pressure relative to the sound pressure at the hearing threshold can be expressed as “decibels (sound pressure level)” or “dB (SPL)” (Crocker 2007). In Figure 20.3, the hearing threshold curves in dB (SPL) at various frequencies are approximated for the human, dog, and mouse. The rabbit and rhesus monkey have similar hearing profiles as the dog (Turner et al. 2005). Each curve, or audiogram, has a similar shape in that it takes a higher sound pressure (“louder” sound) for hearing to occur at lower frequencies than in the middle of the frequency range, and then higher-level sound is required for hearing to occur as the frequencies continue to increase.

Figure 20.3

Approximation of audiograms for the human, dog, and mouse.

As noted in Figure 20.3, both dogs and mice hear higher-frequency sound than humans. Studies have shown that husbandry practices and common equipment found in animal facilities produce sound at frequencies above what humans can hear but within the hearing range of animals (Milligan et al. 1993; Small and Dietrich 2007). Conversely, sound at frequencies 1000 Hz would be audible to humans, but animals, such as the mouse, may not be able to hear sound at these frequencies. Although it has been shown that a ventilated rack and animal transfer station both produced sound pressure levels above the ambient level within the human hearing range, the sound pressure levels within the mouse hearing range did not increase above ambient noise from either noise source. In the same study, when various types of construction equipment were used adjacent to the ventilated rack, the sound pressure level within the mouse hearing range was increased, but to a lesser degree for each implement than were the sound pressure levels within the human hearing range. At more distant locations within the animal facility, sound pressure levels from a large jackhammer within the mouse hearing range decreased much more rapidly than did those in the human hearing range, indicating that less of the sound is perceived by mice than by humans (Reynolds et al. 2010). Therefore, it cannot be assumed that animals hear sounds that humans hear or that humans hear sounds that are audible to animals.

In controlled studies where mice are exposed to prolonged high-decibel sound at frequencies well within their hearing range, noise has caused teratogenic and reproductive effects. Noise may also cause changes in stress hormones and sleep, and behavioral disturbances in laboratory animals, as well as changes to the gastrointestinal, cardiovascular, immune, reproductive, and nervous systems (Pfaff 1974; Zakem and Alliston 1974; Fletcher 1976; Peterson 1980; Nawrot et al. 1981; Murata and Takigawa 1989; Turner et al. 2005; Rabat 2007).

Sanitation

Sanitation is the maintenance of environmental conditions in a manner that will limit microbial levels. Most animal facilities include both cleaning and disinfection as components of a sanitation program. Cleaning removes gross debris and organic materials that shield microorganisms to allow for effective disinfectant penetration (NRC 2011). Disinfection eliminates or significantly reduces microbial concentrations. Good sanitation programs start with conscientious planning to ensure that appropriate surface materials are used to allow for ease of sanitation of both the macro- and microenvironment. Surfaces should be smooth and impervious and composed of materials capable of withstanding frequent, vigorous applications of degreasers, detergents, and corrosive disinfectants. Animal rooms should not have unsealed penetrations, cracks, or seams that could house vermin or microorganisms (Rollin and Kesel 1990). To prevent cross-contamination between rooms, each room should have its own dedicated cleaning implements. Mops, buckets, brooms, and so forth, should not be shared between rooms and corridors, as they may be fomites for the spread of pathogens.

Frequency of sanitation is usually determined by the species housed, the type of animal enclosures, and the amount of debris generated by the animals (NRC 2011). Rooms housing large species such as dogs, pigs, and nonhuman primates in open enclosures or kennels require more frequent room sanitation intervals than rooms with rodents housed in microisolator caging (NRC 2011). In general, floors are usually mopped or hosed on a daily basis in the large animal rooms and complete room sanitation is performed every 1–2 weeks, depending on the institutional sanitation program. For small animal rooms, surfaces are cleaned daily and mopped at intervals ranging from daily to once a week (Casebolt 2009). Based on institutional performance standards, rodent rooms are usually sanitized at least monthly and when an empty room is reopened for occupation. In rooms where detergents and disinfectants are applied by a hose or a foamer, proper protective gear, such as goggles or a face shield and a mask, should be worn to protect against aerosolized chemicals, fecal material, or infectious agents. Eye protection should also be worn when applying detergents or disinfectants on walls and ceilings. Care should be taken to ensure that all surfaces are thoroughly rinsed of detergents or disinfectants and that the animals are not sprayed with water or cleaning chemicals during sanitization activities.

Pest Control

A pest control program should prevent, control, or eliminate pest infestations in the animal environment (NRC 2011) due to their potential for disease transmission and contamination of feed and bedding. Pest control plans should include prevention of attraction and access to the facility by insects and vermin from the outside, as well as a vigorous sanitation program on the inside. The implementation, control, and monitoring of the plan should be documented.

How the exterior of a facility is designed can play a major role in preventing vermin excursions into the interior of the facility. For example, the type of landscaping and type of light fixtures, doors, and finishes, if chosen properly, can enhance an external pest control program. The National Institutes of Health (NIH) Design Requirements Manual recommends using landscaping plans that do not include ground cover–type plantings in close proximity to the building that could provide nesting and harborage for pests (NIHOM 2013). Conduit and lighting fixtures should be sealed and flush with exterior surfaces to prevent a nesting area for insects or wild birds. All exterior wall penetrations should be sealed, and doors should have door sweeps. The use of chemical pesticides around the facility exterior should be carefully considered for safety and effectiveness and be in compliance with state and federal regulations. If exterior trapping devices are used, they should be regularly checked and documented.

Like the exterior of the building, pest control inside the facility begins with good design features and a robust sanitation program. Interior surfaces should be smooth and of a material that can withstand repeated exposure to cleaners and disinfectants. All junctions and cracks should be sealed and drains fitted with screens or covered in order to prevent infestation by insects or vermin. Light fixtures should be sealed and water resistant to enable adequate sanitation. If “drop-down” ceilings are used in animal areas, the grid system should be tightly gasketed and the tiles made of a washable material. Routine sanitization of animal areas at a frequency appropriate to the species housed is key to the deterrent of pests. The use of chemical pesticides should be avoided whenever possible in animal housing areas due to potential adverse effects on animals (NRC 1997, 2011). Chemical pesticides should not be used prior to review by the veterinarian, operations manager, and investigators. Only humane rodent traps should be used, and they should be checked daily. “Sticky boards” that could entrap rodents should not be used for humane reasons.

Environmental Monitoring

In addition to monitoring the disease prevention and sanitation programs in the animal facility, other environmental influences, such as physical plant functionality, must be monitored and documented as well. Consistency in temperature, ACH, and humidity and light cycles are critical to maintaining normal metabolism and normal behavioral activities in research animals. Automated monitoring systems are currently available that are capable of sampling, recording, and with enhanced options, controlling room conditions. Preferably, an automated system should trigger alarms and send alerts via e-mail, pager, or phone when excursions outside of set parameters occur or when malfunctions are detected in automatic watering systems. In many cases, automated systems designed for laboratory animal facilities serve as redundant systems for building automation systems.

Terrestrial Animals

For terrestrial animals, the immediate surrounding area serves as the primary enclosure or microenvironment and may be a cage, pen, stall, or kennel. Enclosures should be provided that are designed to fit the needs of the species occupying the space. Modern technology offers the ability to provide environments that significantly stabilize the conditions within the microenvironment. Until the advent of individually ventilated caging (IVC) systems, the static microisolator cage was the predominant enclosure for microisolation rodent housing. When compared with static caging, mechanically ventilated mouse isolator cages at approximately 60 ACH have been shown to have dramatically lower levels of ammonia, carbon dioxide, and RH (Memarzadeh et al. 2004). Although IVC improves air quality within the cage, there are aspects of IVC systems that may have an unintended negative impact. Baumans et al. (2002) illustrated that high air velocities (60 ACH and higher) may cause stress to the animals, which was manifested in the study through avoidance behavior. In their study, when given the freedom to move between adjoining mechanically ventilated cages and static caging, mice showed a preference for the static cages and avoided air inlets that were placed low in the IVC systems. The study reinforced the importance of providing appropriate amounts of nesting material in IVC since when provided with huts or nesting material, avoidance behavior decreased and nests were built by mice in IVC. Mice in this study also demonstrated a preference for the type of IVC systems where forced air entered from the cage lid instead of the cage wall (Baumans et al. 2002). Along with controlling the flow dynamics inside the cage, most modern IVC systems allow for changing the directional flow of air through the cages from positive to negative and vice versa relative to the room. Caging in a barrier setting would normally have a positive directional flow in relation to the macroenvironment, and in most situations, caging being used for control of pathogens or experimental infectious agents would incorporate a negative airflow setting (Lipman 2009). Advancing technology also provides methods for monitoring the microenvironment in IVC. Many vendors can provide optional specialized caging features that will sample and report on environmental conditions, such as temperature, humidity, and ammonia concentration inside of the cage. Newer models of exhaust blowers on IVC systems will display cage exhaust temperature digitally. Caging systems can also be equipped with wireless technology capable of monitoring, reporting, and storing data on environmental conditions. This technology can also generate alarms for excursions outside of desired parameters or power loss. An emergency power supply, to ensure that the cages are properly ventilated in the event of a power failure, is common when using IVC systems. IVC racks are portable equipment, so it is important that consideration be given to using twist-lock plugs or some other method of preventing inadvertent disconnection from the electrical source in the event that the IVC rack is moved (NRC 2011).

In direct contrast to a controlled, predictable micro- and macroenvironment, research animals such as dogs, nonhuman primates, and agricultural animals have often been housed in outdoor or indoor–outdoor enclosures. Frequently, animals with access to indoor–outdoor housing are group housed in an effort to provide added enrichment. Although animals can benefit from this type of setting, group housing could present species-specific challenges in regard to enclosure density and temperament or social rank of the animals. This type of environment can be distressful to some animals if not carefully controlled (Novak and Suomi 1989; Overhall and Dyer 2005). Indoor–outdoor and outdoor environments can also confound research results due to natural influences, such as seasonal changes in temperature, humidity and light cycles, an inability to control nutrient intake in grazing animals, and the possible presence of naturally occurring pathogens (NRC 2011).

Along with controlling and monitoring the microenvironment, we must also consider sanitation practices as an important component of providing for the health and well-being of the animals. Cage changing frequency and effective sanitation of the primary enclosure both contribute to the quality of the microenvironment. Enclosures and accessories should generally be sanitized every 2 weeks. Some types of caging systems or enclosures may require more or less frequent sanitization based on the cage’s size, animal holding density, or type of air ventilation system (NRC 2011; Horn et al. 2012). The process of sanitation or disinfection may necessitate the use of chemical cleaners or disinfectants. Products used for sanitizing the microenvironment should be free of perfumes or odors and should be thoroughly rinsed from all surfaces. Studies evaluating routine distressful environmental conditions in the animal facility report that the presence of perfumes or odors from volatile chemicals can elicit physiological reactions (Castelhano-Carlos and Baumans 2009). A variety of cleaning methods may be effective, such as mechanical cage washers or manual sanitization; however, the method should be appropriate for the materials being used (NRC 2011). Recommended methods for ensuring and validating the effectiveness of the sanitization process can include visual inspection, monitoring water temperature, and microbial and ATP testing. These methods are appropriate for both the macro- and the microenvironment.

Lighting is another component of the microenvironment that usually differs significantly from the macroenvironment. Type of caging and location of the cage in the room or on the rack can influence the amount of light reaching the animals within the enclosure. Studies have shown that light intensity can vary as much as 80-fold from the top to the bottom of the rack in transparent cages and up to 20-fold inside the cage (NRC 2011). Cage material can affect the amount of light that penetrates the enclosure, with clear translucent caging allowing the highest light intensity. There is evidence that light levels that are too low can negatively impact normal physiological responses in some strains of rodents (Faith and Huerkamp 2009). Room lighting should be such that consideration is given to the well-being of the species to be housed at the cage level, in addition to the ability of staff to safely visualize the animals (NRC 2011).