The Guide for the Care and Use of Laboratory Animals (Guide) recommends minimum floor space per mouse based on weight, with no other factors considered. We conducted a randomized experiment to evaluate the effect of housing density on reproductive indices and corticosterone levels in lactating mice. Female mice matched for age, strain, and date-of-pregnancy were housed individually. At parturition the dams were randomly allocated to have litters culled or remain intact. The experimental group had litters culled to meet the Guide's space density requirement. Litters of the second group were maintained as the numbers born to each dam. Fecal corticosterone levels (first-generation mice only), growth, and weaning weights were measured for mice in all cages; in addition, the reproductive behavior of progeny generated under both housing conditions was assessed to determine whether a space×litter size interaction affected subsequent reproduction. The growth rates for pups from culled litters were significantly greater than those from intact litters. The first-generation pups showed no statistically significant differences in fecal corticosterone or reproductive parameters. The second-generation pups showed no statistically significant differences in growth rates. The results of the study suggest that a strict interpretation of space requirements as listed in Table 2.1 of the Guide is not warranted for lactating dams with litters.

The Guide for the Care and Use of Laboratory Animals contains recommended housing densities for rodent species that are commonly used by the scientific community. However, at the time of the Guide's publication, housing density recommendations were based heavily on the professional judgment of qualified scientists. Some scientists therefore question whether rodents can be housed at greater densities, whereas others wonder whether the space currently provided for rodents is sufficient. The present study was designed to determine the effect of housing adult female BALB/c- and C57BL/6-mice in standard 75-in² (484-cm²) ventilated cages at various housing densities (n = 2, 5, and 10 mice/cage). Measures of weight gain, plasma corticosterone, behavior, and immune parameters were evaluated at 7, 28, and 70 d after housing allocation. Housing BALB/c mice at 10/cage had negative effects on weight gain, corticosterone, behavior, and immune parameters. Housing C57BL/6 mice at 10/cage did not affect immune function or weight gain, although behavior and corticosterone showed statistical trends implying a negative effect Differences associated with housing densities of 2 and 5 mice/cage were less robust for all variables measured. We conclude that housing female BALB/c mice at 10 mice/cage (that is, at twice the Guide-recommended density) affects their physiology. We also conclude that mice vary in their responses in the parameters measured. These observations support the conclusion that it will be extremely challenging to scientifically determine an optimal cage density standard that can be uniformly applied across all mouse strains.

We evaluated the effect of an enrichment device (that is, a polyurethane bone) on the voluntary consumption of ethanol-containing gel by single-housed rats. Male Sprague–Dawley rats (n = 5 per group) were exposed for 4 d to each of the following 3 treatments: access to a new synthetic bone and ethanol gel for 1 h daily (treatment 1); a new bone was left in the cage for 24 h, with access to ethanol gel for 1 h daily (treatment 2); and both the bone and ethanol gel remained in the cage for 24 h (treatment 3). Average alcohol consumption over 4 d was 0.86 ± 0.13, 0.99 ± 0.13, and 5.19 ± 0.37 g/kg in the absence of the bone for treatments 1, 2, and 3, respectively, and 1.00 ± 0.13, 0.620 ± 0.07, and 5.55 ± 0.38 g/kg with the bone for treatments 1, 2 and 3, respectively; none of these values differed significantly with regard to presence of the bone. During treatment 1, time spent with the synthetic bone was highest on the first 2 d, which altered the rate of ethanol consumption but not the total amount of ethanol consumed. During treatments 2 and 3, the rate and amount of ethanol consumption were comparable to basal levels. We conclude that adding an enrichment device that rats can chew and manipulate does not alter ethanol gel consumption. If used, environmental enrichment techniques should be evaluated during the research planning stages to avoid unintended alterations in the response to variables of interest.

Environmental enrichment for laboratory animals is a widely accepted practice for many species, but few studies address the periods of preadolescence and adolescence. Provision of igloos, tunnels, nesting materials, and social or communal housing are commonly used enrichment strategies in rat cages. In the present study, the effects of individual, pair, and trio housing and the presence or absence of physical cage enrichment on the growth rate, food consumption, and locomotor behavior of juvenile male rats through adolescence were examined. The results indicated that social and physical enrichment decreased the growth and feeding rates and locomotor activity of developing rats as compared with rats living in an impoverished environment. The results show that the growth rates are dependent predominantly on environmental enrichment and that social enrichment alone has no effect. These results demonstrate that enrichment can have significant effects on growth and behavior of male rats.

Rabbits used in the production of antibodies can be housed individually or in groups. This study compared the serum chemistries, antibody production, physiologic plasma cortisol levels, and white blood cell (WBC) counts of female New Zealand White rabbits housed in 2 different housing systems. The control group was housed individually in stainless steel cages, and the experimental group was group-housed on aspen shavings spread on the floor of the animal room. Plastic crates were placed in the group-housing area to provide opportunities for rabbits to hide, and a litter box was available at all times. Both groups received the same food and water and similar environmental enrichment devices. Clinical pathology laboratory evaluations of serum chemistries, immune responses, physiologic parameters such as plasma cortisol, and WBC counts were compared. The group-housed animals had lower WBC counts and higher levels of plasma cortisol than did rabbits individually housed. In addition, the group-housed animals had significantly less weight gain during the first week. Antibody production did not differ between the 2 groups. Group housing appeared to be an appropriate method of housing rabbits for use in research.

Characterization of animal housing conditions can determine the frequency of bedding and cage changes, which are not standardized from facility to facility. Rabbits produce noticeable odors, and their excreta can scald and stain cages. Our facility wanted to document measurable airborne contaminants in a laboratory rabbit room in which excreta pans were changed weekly and cages changed biweekly. Contaminants included particulate, endotoxin, ammonia, carbon dioxide, and a rabbit salivary protein as a marker for rabbit allergen. Concentrations were measured daily over a 2-wk period in a laboratory animal facility to determine whether they increased over time and on days considered to be the dirtiest. Except for ammonia, concentrations of all airborne contaminants did not differ between clean and dirty days. Concentrations were lower than occupational health exposure guidelines for all contaminants studied, including ammonia. After measurement of concentration, a model was applied to calculate mean emission factors in this rabbit room. Examples of emission factor utilization to determine airborne contaminant concentration in rabbit rooms under various environmental conditions and housing densities are provided.

A novel environmental preference chamber (EPC) was developed and used to assess responses of laboratory mice to atmospheric ammonia. The EPC features 1) a test chamber with 4 individually ventilated, mutually accessible compartments; b) automated tracking of mouse movements by using paired infrared sensors; c) identification of individual mice by using photosensors; d) monitoring and regulation of the NH₃ concentration in each compartment; and e) personal-computer–based data acquisition. In an initial preference study with the EPC, 4 groups of 4 laboratory mice (BALB/c/Bkl; body weight, 13.4 to 18.4 g) were each given a choice among 4 NH₃ concentrations (mean ± SE) of 4 ± 2, 30 ± 2, 56 ± 4, and 110 ± 6 ppm for 2 d after a 2-d familiarization period. Once trained to use the intercompartment tunnels, the mice made extensive use of the EPC, with each group making more than 2000 intercompartment movements during 48 h. Video recording verified the results of the automatic tracking system, which detected and correctly determined mouse location for 79% of the moves. The use of photosensors proved to be ineffective in recognizing individual mice. Although the EPC would benefit from refinement and further development, it simplified analysis of locomotion behavioral data. Results of the preference study indicated that the mice exhibited no clear preference for, or aversion to, any of the experimental concentrations of ammonia and that the mice clearly preferred the upper 2 compartments of the chamber over the lower 2 compartments. Further investigation should be conducted to verify these preliminary results and explore other preferences of laboratory mice for environmental conditions and resources.

This study compares resuable and disposable individually ventilated mouse cages in terms of the formation of intracage CO₂ and NH₃. Crl:CD-1(ICR) female mice were placed in either disposable or reusable ventilated cages in a positive pressure animal rack. Intracage CO₂ and NH₃ were measured once daily for 9 d; temperature and relative humidity were monitored for the first 7 d. Results indicated higher CO₂ levels in the rear of the disposable cages and in the front of the reusable cages. This pattern corresponded to where the mice tended to congregate. However, CO₂ concentrations did not differ significantly between the 2 cage types. Average CO₂ levels in both cage types never exceeded approximately 3000 ppm. Intracage NH₃ began to rise in the reusable cages on day 4, reached approximately 50 ppm by day 5 and by day 9 was greater than 150 ppm at the cages' rear sampling port while remaining at approximately 70 ppm at the front sampling port. Intracage NH₃ levels in the disposable cages remained less than or equal to 3.2 ppm. Intracage temperature and relative humidity were approximately the same in both cage types. We concluded that the disposable ventilated cage performed satisfactorily under the conditions of the study.

The use of automated watering systems for providing drinking water to rodents has become commonplace in the research setting. Little is known regarding bacterial biofilm growth within the water piping attached to the racks (manifolds). The purposes of this project were to determine whether the mouse oral flora contributed to the aerobic bacterial component of the rack biofilm, quantify bacterial growth in rack manifolds over 6 mo, assess our rack sanitation practices, and quantify bacterial biofilm development within sections of the manifold. By using standard methods of bacterial identification, the aerobic oral flora of 8 strains and stocks of mice were determined on their arrival at our animal facility. Ten rack manifolds were sampled before, during, and after sanitation and monthly for 6 mo. Manifolds were evaluated for aerobic bacterial growth by culture on R2A and trypticase soy agar, in addition to bacterial ATP quantification by bioluminescence. In addition, 6 racks were sampled at 32 accessible sites for evaluation of biofilm distribution within the watering manifold. The identified aerobic bacteria in the oral flora were inconsistent with the bacteria from the manifold, suggesting that the mice do not contribute to the biofilm bacteria. Bacterial growth in manifolds increased while they were in service, with exponential growth of the biofilm from months 3 to 6 and a significant decrease after sanitization. Bacterial biofilm distribution was not significantly different across location quartiles of the rack manifold, but bacterial levels differed between the shelf pipe and connecting elbow pipes.

Rodent toxicology studies have historically been performed in wire-bottom cages, but the 1996 Guide for the Care and Use of Laboratory Animals recommends solid-bottom caging with bedding. Some investigators have expressed concern that changing to solid-bottom cages would interfere with technicians' ability to detect clinical signs. To test this hypothesis, rats were housed in both types of caging and given compounds to induce a variety of subtle clinical signs common to toxicology studies including chromodacryorrhea, soft stool, stereotypic behaviors, mild hypoactivity, abnormal postures, and discolored urine. For one comparison, fecal pellets were removed to simulate decreased production of feces. Technicians, blinded from knowing which animals had been treated, observed the rats and recorded the clinical signs they detected. The technicians who administered the treatments verified that clinical signs were present before and after the blinded technicians made their observations. The number of animals observed with clinical signs divided by the number of animals verified with signs was calculated for each compound and compared between the cage types by using the Fisher Exact Test. The only statistically significant difference observed was a diminished ability to detect discolored, dark urine from rats in wire-bottom cages. These results suggest that concerns about technical staff's inability to detect clinical signs in toxicity tests should not prevent investigators from using solid-bottom cages with bedding.