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Maintenance of an appropriate microenvironment for rodents used in research is of paramount importance because changes in environmental parameters such as O2 and humidity can influence animal health and welfare and potentially alter research results. Here we evaluated the microenvironment of mouse and rat disposable cages after removal from mechanical ventilation in order to guide recommendations for their use. Cages with sealed IVC lids, unsealed lids (partially ajar), and lids without the exhaust filter (for rats) or static lids (for mice) were removed from the ventilated rack and were thereafter monitored CO2, O2, and NH3 levels. For mice, effects were investigated under both standard (set point of 72°F/22°C) and thermoneutral (set point of 82°F/28°C) temperatures. When IVC with sealed lids and group-housed C57BL/6J male mice were removed from ventilation under standard temperatures, CO2 started at 6,600 ± 265 ppm at 0 h and rose to 42,500 ± 7,263 ppm at 1 h, with mice showing a visibly elevated respiratory rate in 1 of the 3 cages; CO2 stabilized at 26,150 ± 3,323 ppm at 8 h. In contrast, CO2 levels in cages with single mice were stable after 1 h (1,350 ± 409 ppm at 0 h, 9,367 ± 802 ppm at 1 h, and 8,333 ± 1,115 ppm at 8 h). Findings were similar at thermoneutral temperatures: sealed group-housed mice cages started at 3,617 ± 475 ppm at 0 h and rose to 39,333 ± at 5,058 ppm at 1 h, whereas sealed cages with 1 mouse started at 1,117 ± 247 ppm at 0 h and were 7,500 ± 1,997 ppm at 8 h. IVC with sealed lids and pair-housed Crl:CD(SD) female rats rose to 48,000 ± 2,828 ppm CO2 and over 70% humidity within 1 h. By 3 h, IVC with sealed lids and singly housed rats had 40,167 ± 5,132 ppm CO2, and rats were displaying a visually elevated respiratory rate. O2 levels had an inverse relationship with CO2 levels. Removing the rat lid exhaust filter was not helpful. However, leaving the IVC lid ajar ameliorated the rise in CO2 and fall in O2 for both species. Therefore, IVC with sealed lids and group-housed mice should not be removed from ventilation more than 1 to 2 h; IVC containing pair- or singly-housed rats IVC should not be removed for more than 1 or 3 h, respectively. Whenever possible, such cages should be fitted with static lids, left partially ajar and monitored, or replaced on ventilation.

Vivarium husbandry practices are based on performance data and adhere to applicable regulatory guidelines. Refinements in husbandry and optimization of sanitization protocols improve animal wellbeing and help standardize the microenvironment, contributing to research reproducibility. The objective of this study was to evaluate the microenvironment to establish performance standards for mouse husbandry and sanitization, including housing at standard and thermoneutral temperatures. Male C57BL/6J mice were housed singly and in groups in disposable IVCs on α-cellulose or corncob bedding and microenvironmental indicators (ammonia, carbon dioxide) were evaluated. In addition, microbial bioburden tests (ATP and RODAC) were performed on cages and cage accessories on days 0, 7, 14 and, 28 to 30 after cage change. Water testing and aerobic culture of the waterspout of bottles containing chlorinated water were performed to determine acceptable replacement schedules. Ammonia levels remained below the National Institute of Occupational Safety and Health 8-h recommended exposure limit for humans (25 ppm) at all time points for all housing conditions through day 21 for group-housed mice, and through day 30 for singly housed mice. Microbial bioburden results for cage accessories and water testing were acceptable up to 28 d after cage change (RODAC less than 50 CFU; ATP less than 100,000 RLU) at both standard and thermoneutral housing temperatures. Mice remained clinically healthy throughout the studies. These results support site operating practices and verify extended sanitization recommendations per the Guide of the Care and Use of Laboratory Animals in this disposable IVC environment: group-housed mice receive bottom cage and water bottle change up to every 14 d with full cage change (including lid and accessories) every 28 d, and singly housed mice receive full cage change every 28 to 30 d or sooner.

Molecular-based methods have shown potential for improving pathogen detection and reducing animal use. While increasing evidence supports rodent-free environmental health PCR pathogen detection, limited information is available regarding efficacy for disposable individually ventilated caging systems. In such systems, testing of plenum exhaust air dust is ineffective, and the use of collection media is optimal. We performed a series of studies to compare PCR infectious agent detection with dust collected on media placed in a mouse-free soiled bedding cage, the cage exhaust filter of an occupied sentinel cage, and direct sampling from colony and sentinel mice with traditional soiled bedding mouse sentinels. We hypothesized that after a 3-mo period, testing of filter media agitated in a soiled bedding cage would be equal to or more sensitive than more traditional methods. Agitated media detected Astrovirus-1, segmented filamentous bacteria and Helicobacter ganmani to a degree comparable to testing lid exhaust filter PCR from a sentinel mouse cage, but opportunists such as Staphylococcus aureus and Proteus mirabilis were not detected consistently, and H. hepaticus was not detected at all. Direct sampling of pooled fecal pellets and body swabs from sentinel mice and testing using PCR also failed to reliably detect opportunists and Helicobacter spp. While further work is needed to refine use of filter media in soiled bedding for detection of lower prevalence opportunists, this report provides evidence that a rodent-free method of reliably detecting murine agents in a disposable individually ventilated cage system with cage-level filtration outperforms direct sampling of soiled bedding sentinel mice.

In oncology research, while xenograft tumor models are easily visualized and humane endpoints can be clearly defined, metastatic tumor models are often based on more subjective clinical observations as endpoints. This study aimed at identifying objective non-invasive criteria for predicting imminent distress and mortality in metastatic lung tumor-bearing mice. BALB/c and C57BL/6 mice were inoculated with CT26 or B16F10 cells, respectively. The mice were housed in Vium smart cages to continuously monitor and stream respiratory rate and locomotion for up to 28 days until scheduled euthanasia or humane endpoint criteria were met. Body weight and body temperature were measured during the study. On days 11, 14, 17 and 28, lungs of subsets of animals were microCT imaged in vivo to assess lung metastasis progression and then euthanized for lung microscopic evaluations. Beginning at day 21, most tumor-bearing animals developed increased respiratory rates followed by decreased locomotion 1–2 days later, compared with the baseline values. Increases in respiratory rate did not correlate to surface tumor nodule counts or lung weight. Body weight measurement did not show significant changes from days 14–28 in either tumor-bearing or control animals. We propose that increases in respiratory rate (1.3–1.5 X) can be used to provide an objective benchmark to signal the need for increased clinical observations or euthanasia. Adoption of this novel humane endpoint criterion would allow investigators time to collect tissue samples prior to spontaneous morbidity or death and significantly reduce the distress of mice in the terminal stages of these metastatic lung tumor models.

The typical daily water intake of common marmosets (Callithrix jacchus) in a research setting has not been well characterized. Because these New World primates are in demand as animal models for neurobehavioral experiments, which can include the potential use of fluid regulation for training, veterinary and research staff need to understand how marmosets keep hydrated under normal circumstances. In the current study, we measured the water consumption of older (age, 5 to 12 y; n = 11) and younger (age, 1 to 2 y; n = 11) marmosets every 3 h during the 12-h light phase in 2 different months (January and July). The overall daily water intake (mean ± 1 SD) was 61.3 ± 20.4 mL/kg (range, 36.3 to 99.0 mL/kg); water intake by an individual marmoset or cohoused pair was fairly consistent from day to day. Water intake did not change across the four 3-h periods measured during the day, and minimal water was consumed overnight when the room lights were off. In addition, daily water intake did not differ between the 2 mo of measurements. Older animals drank significantly more than the younger group, and weight was directly correlated with water intake. Water intake was not affected by body condition score or housing status. The variation in water consumption among marmosets underscores the need for individualization of fluid regulation guidelines.