Journal of the American Association for Laboratory Animal Science : JAALAS最新文献

There are limited evidence-based husbandry recommendations for laboratory zebra finches (Taeniopygia guttata), including appropriate light sources. Light-emitting diode (LED) technology has been shown to improve circadian regulation and reduce stress in some laboratory animal species, such as mice and rats, when compared with cool-white fluorescent (CWF) lighting, but the effects of LED lighting on zebra finches have not been published. We compared the effects of broad-spectrum, blue-enriched (6,500 Kelvin) CWF and flicker-free LED lighting on the behavior, stress, and reproductive outcomes of indoor-housed zebra finches. Using breeding pairs housed in cubicles illuminated with either CWF or LED lighting, we compared the reproductive output as determined by clutch size, hatching rate, and hatchling survival rate. We also compared the behavior of group-housed adult males, first housed under CWF followed by LED lighting, using video recordings and an ethogram. Fecal samples were collected from these males at the end of each recording period, and basal fecal corticosterone metabolite (FCM) levels were compared. A FCM assay for adult male zebra finches was validated for efficacy and accuracy using a capture-restraint acute stress response and parallelism analysis, respectively. The breeding pairs had no significant difference in the clutch size or percent hatching rate, but percent hatchling survival improved under LED with an increased proportion achieving 100% survival. There was no significant difference in FCM between the lighting treatments. However, the activity budgets of the birds were altered, with a reduction in flighted movement and an increase in enrichment manipulation under LED. Overall, these results support the use of blue-enriched, broad-spectrum flicker-free LED as a safe alternative to CWF lighting for breeding and nonbreeding indoor-housed zebra finches.

The use of soiled-bedded sentinels (SBSs) has historically been the standard for colony health surveillance monitoring at our institution. With the advent of newer technologies in which dust collected from filters is tested by PCR, we compared traditional SBS with PCR testing of both exhaust air dust collected from a filter in the downstream vertical plenum (exhaust dust test [EDT]) and the SBS cage-level exhaust filter (SCEF). Our hypothesis was that both methods of filter testing would identify more pathogens than SBS testing. Twenty-five individually ventilated mouse racks that used disposable caging were sanitized and placed into rotation. Rack plenums were tested by PCR to verify negative results before the study start. Exhaust dust collection media were placed in the exhaust plenum (n = 25). SBS cages were placed on each side of the rack with 2 mice per cage (n = 42 mice), with the remaining cage slots occupied by research animals. At each triweekly cage change, the exhaust air filters were carefully removed from the cage top, placed in sterile 50-mL conical tubes, and pooled for submission. After 3mo, the SBS mice were tested via serology for bacterial and viral agents and by PCR for Helicobacter species, pinworms, and ectoparasites. In addition, the EDT filter and SCEF were collected for PCR to evaluate for the same agents. Our results indicate that the SCEF consistently detected agents more frequently than the EDT filter placed in the plenum and that the EDT filter media detected agents more frequently than did the SBS mice. Our data suggest that both PCR methods of detection are superior to SBS for individually ventilated disposable rodent cages and that the SCEF is superior to EDT. These data supported our movement of institution toward environmental monitoring as a method of rodent colony health surveillance.

Animal research facilities are noisy environments. The high air change rates required in animal housing spaces tend to create higher noise levels from the heating and cooling systems. Housing rooms are typically constructed of hard wall material that is easily cleaned but simultaneously highly reverberant, meaning that the sound cannot be controlled/attenuated so the sounds that are generated bounce around the room uncontrolled. (Soft, sound-absorbing surfaces generally cannot be used in animal facilities because they collect microbes; various wall surface features and sound control panel options are available, although rarely used.) In addition, many of our husbandry tasks such as cage changing, animal health checks, cleaning, and transporting animals produce high levels of noise. Finally, much of the equipment we have increasingly employed in animal housing spaces, such as ventilated caging motors, biosafety, or procedure cabinets, can generate high levels of background noise, including ultrasound. These and many additional factors conspire to create an acoustic environment that is neither naturalistic nor conducive to a stress-free environment. The acoustic variability both within and between institutions can serve as an enormous confounder for research studies and a threat to our ability to reproduce studies over time and between research laboratories. By gaining a better appreciation for the acoustic variables, paired with transparency in reporting the levels, we might be able to gain a better understanding of their impacts and thereby gain some level of control over such acoustic variables in the animal housing space. The result of this could improve both animal welfare and study reproducibility, helping to address our 3Rs goals of Replacement, Reduction, and Refinement in the animal biomedical research enterprise.