Enhancing Rodent Welfare, Cost Savings, and Efficiency: Implementation and Review of Data-Driven Improvements

Abstract

At the University of Chicago, we have implemented multiple practical, data-driven 3Rs (replacement, reduction, refinement) strategies to improve rodent welfare over the past 5 years. As there are limited publications that outline an institutional approach, this work reviews the enhancements in animal welfare and operational practices implemented in our rodent program, offering a resource and reference for other institutions. These measures encompass the use of environmental health monitoring for both colony and quarantine surveillance and housing modifications, as well as advancements in enrichment, including nesting and structural options, adoption of mouse-preferred bedding substrate, and housing room temperature modifications to better support the thermoregulatory needs of the mice. In addition, we implemented cage change modifications, which include a scent transfer and reduced cage change frequency. Finally, the use of animal handling improvements at our institution includes refined handling techniques and transport cart modifications. Ultimately, we have been successful at improving animal welfare while also implementing cost savings and efficiency improvements.

Introduction

The University of Chicago (UChicago) Animal Resources Center is a midsized animal research program with ∼20,000 rodent cages that expand across 3 vivaria. Rodent work is performed by ∼160 active principal investigators. Research activities span a multitude of scientific interests with significant areas focusing on oncology, neurosciences, immunology, infectious disease, diabetes, microbiome, and translational research models. At any institution, advancing animal welfare and operation practices is essential; however, this can be challenging given the needs of multiple stakeholders. As there are limited publications that outline an institutional approach, this work reviews the data-driven enhancements in animal welfare and operational practices implemented in our rodent program over the past 5 years, offering a resource and reference for other institutions.

Animal welfare is a well-established field of study, but there is currently no commonly accepted definition of this term.^1–3 Welfare experts do agree that the term refers to the actual state of the individual animal^3,4 and includes the animal’s overall condition, including physical health and functioning, ability to perform species-specific, highly motivated behaviors, and emotional state.¹ To provide optimal welfare, it is essential to support all 3 of these aspects. When assessing welfare, however, veterinarians may tend to overemphasize animal health and, therefore, animals in good health may automatically be considered to have good welfare.^2,3 In addition, when assessing animal welfare, it is important to measure the behavioral and physiologic responses of animals, not relying solely on inputs (eg, what humans provide to animals).^1,2

In addition to animal welfare–based improvements, many of the implemented changes at our institution have also provided improved operational efficiency and cost-saving opportunities. Improvements in operational efficiencies have allowed staff to dedicate more time to programmatic and animal welfare initiatives, while also enabling resources to be redirected toward additional enhancements in these areas. Human well-being and animal welfare are actively connected as described in the One Health, One Welfare initiative. Therefore, improvements to animal welfare may enhance human well-being and vice versa.^5–7 Based on surveys, it is shown that increased compassion satisfaction in laboratory animal personnel is associated with decreased animal stress and pain, utilization of more enrichment, and increased positive human–animal interactions.^5,6 Overall, those who believe that the institution at which they work is trying to improve animal welfare have increased compassion satisfaction.^5,6 Ultimately, these enhancements at our institution have not only benefited animal welfare but have also provided the opportunity to benefit staff well-being and the institution as a whole.

Overall, our commitment to continual improvement, with an emphasis on the advancement of animal welfare, is at the forefront of our research and animal care programs. By continually seeking and implementing improvements within the vivarium, we have not only enhanced the welfare of the animals in our care but have also realized significant benefits for the institution and the animal care team. These initiatives frequently lead to increased operational efficiency and meaningful cost savings while also improving staff morale, illustrating that progress in animal welfare can create value at multiple levels.

Programmatic welfare/3Rs improvements.

The 3Rs—replacement, reduction, refinement—are the ethical principles that have had the greatest impact within laboratory animal care and medicine. Ultimately, the 3Rs have become the international standard for humane animal research, and they appear in multiple regulations. Due to their common use in the laboratory animal care field, this work uses the 3Rs framework to discuss multiple data-driven welfare improvements we have been able to achieve (Table 1).

Table 1.

Overview of the Welfare/3Rs, Presumed Staff, and Operational Impacts of the Practical, Data-Driven Programmatic Rodent Improvements Implemented at UChicago

Category	Type	Welfare/3Rs impact	Presumed staff impact	Operational impact
Environmental health monitoring	Colony health monitoring	Replacement	Time savings and decreased compassion fatigue	$203,901 (transition to EHM)11 + $10,625 (updated testing panel)15 = $214,526/ya cost savings
Environmental health monitoring	Quarantine health monitoring	Refinement	Time savings	5 h/y time savings10
Housing	Nesting enrichment	Refinement	Additional indicator in mouse health assessment	No additional cost
Housing	Structural enrichment	Refinement	Improved/faster response to tunnel handling	N/A
Housing	Bedding	Refinement	Preferred due to improved operational efficiency and ergonomics51	Decreased personnel labor cost savings not calculated
Housing	Housing room temperatures	Refinement	N/A	N/A
Cage change modifications	Scent transfer	Refinement	Time savings due to extended frequency of accessory cage change	$106,454/ya cost savings66
Cage change modifications	Cage change frequency	Refinement	Time savings	Decreased cost savings for the following were not calculated: •Personnel labor •Caging wear and tear •Bedding and enrichment purchased •Utilities
Animal handling	Refined handling	Refinement	Decreased repetitive motion injuries	N/A
Animal handling	Transport cart modifications	Refinement	N/A	N/A
Operational cost-savings and efficiencies	Discontinue autoclaving of cage components	N/A	Time savings	Decreased cost savings for the following were not calculated: •Personnel labor •Caging wear and tear •Utilities
Operational cost-savings and efficiencies	Diet conservation	N/A	N/A	$170,343/ya cost savings

^a Inflation calculator used to standardize data to September 2025 costs: https://data.bls.gov/cgi-bin/cpicalc.pl.

Environmental Health Monitoring

At our institution, we replaced live animal sentinel health monitoring with environmental health monitoring, using exhaust dust testing (EDT) for IVC systems and sentinel-free soiled bedding (SFSB) for static caging and quarantine.

Colony health monitoring.

Environmental health monitoring (EHM) is any type of health surveillance that does not require the use of live animal sentinels, which has been shown to be more sensitive than the use of soiled bedding sentinels, and which was recently addressed in a systematic literature review.⁸ It is therefore considered a replacement. There are 2 main forms of EHM for routine colony health surveillance, EDT and SFSB. EDT involves the use of swabs or media to collect dust that accumulates in IVC exhaust systems and can be used only with certain types of IVC racks.⁸ SFSB procedurally resembles a soiled bedding sentinel program in that it involves serial pooling of soiled bedding from rodent colony cages into a cage or bin, but no live animals are used. Rather, the soiled bedding is then sampled with media. SFSB can be used with all caging types.⁸

After assessing the sensitivity of the EHM for a year,⁹ we switched to EDT for IVC systems and SFSB for static caging and the quarantine program.¹⁰ Due to this process, we decreased our programmatic animal use by >1,600 animals each year.¹¹ In addition, this change decreased labor and costs as documented at other institutions.¹² We found that EHM is 26% less expensive than the previous soiled bedding sentinel program, which resulted in a savings of $160,800 annually in 2019.¹¹ It decreased the time required by the veterinary technician overseeing the program by 1.5 h/wk per every 10,000 cages, which is ∼3 h/wk at our institution.¹¹ Finally, this welfare improvement involved a beneficial mental health component for personnel, as it removed the need for euthanasia of the sentinel animals, thus potentially mitigating compassion fatigue.¹³

For implementation of EHM, we worked closely with our commercial diagnostic laboratory. We created a system to record when racks are moved and sanitized as a way to determine what needs to occur with the sampling media in that rack. Overall, we used multiple resources, including those available online, to assist with this change such as a guide on how to switch, SOPs, frequently asked questions, and an editable slide deck used to convince stakeholders of the value to be realized.¹⁴

More recently, in 2023, we reassessed the routine health monitoring PCR testing panel based on recent guidance¹⁵ and saved >$10,000 annually by removing multiple infectious agents with an ∼0% prevalence rate from the routine health monitoring panel.¹⁶ We continue to screen for these low prevalence/eradicated agents during quarantine and in biologic materials testing panels since these areas are the main points of potential entry for pathogens.¹⁵ We occasionally encounter resistance during exports to other institutions due to this change; however, we have been able to resolve this through education on agent prevalence and reassurance that testing at quarantine and of biologic materials targets risk. If the accepting institution still requires additional testing, we do this at their expense.

Quarantine health monitoring.

With the introduction of PCR testing, direct colony sampling (DCS) is the most commonly used methodology for diagnostic testing of quarantined animals according to a 2021 survey.¹³ This involves “[t]he collection of feces, fur swabs, oral swabs, or blood to noninvasively (or minimally invasive) sample specific animals or their cage microenvironment.”⁸ We wanted to refine this method and assess whether SFSB could be used for quarantine diagnostic testing instead, as there are multiple publications on its use for routine colony health monitoring.¹⁰ Therefore, we compared the sensitivity of DCS of quarantine animals to SFSB for quarantine health monitoring. During the 2-week quarantine period, we concluded that SFSB had comparable sensitivity to DCS for the quarantined animals, which is consistent with a similar study.^10,17 Most importantly for animal welfare, SFSB is a refinement because it does not involve handling the animal, as the SFSB diagnostic sample is collected from the soiled bedding collected at the standard cage change.¹⁰ Alternatively, if no cage change is needed during quarantine, a known amount of bedding from the latrine area of the cage can be removed for diagnostic testing. This is in comparison to DCS, which requires handling and restraint of the animal to acquire the pelt, as well as oral swabs and fecal samples. In addition, we found a substantial decrease in the time it takes to perform SFSB versus DCS, thus improving efficiency.¹⁰

Housing

At our institution, we transitioned to providing shredded paper strips as a default nesting enrichment for all mice, avoiding cotton nestlets, and prioritizing its use over structures such as igloos. Instead of other structural enrichment items, we began implementation of home-cage tunnels for handling that also serve as a form of structural enrichment. In addition, we standardized paper pulp cellulose bedding for its welfare and operational benefits. Finally, we increased room temperatures in rodent housing rooms to better support the thermoregulatory needs of the mice.

Nesting enrichment.

Mice have a strong preference for nesting material and prefer nesting material over every other form of enrichment, including social companions.^18–20 This preference is highly conserved from wild mice.^18,21 Nesting material is important for breeding, but also for nonbreeders, as it allows for protection from aggressive companions, shelter from external stimuli, and thermoregulation.^21,22 These nests are especially important to ease the cold stress that may result from using the recommended room set point temperatures in the Guide for the Care and Use of Laboratory Animals²³ and the multiple air changes per hour that are inherent to IVC racks.^21,22 In fact, the ability to build a nest reduces stress for mice as it allows them an opportunity to alter their surroundings, offers some control over their environment, and reduces stereotypical and abnormal behaviors.^18,24 Providing mice with the proper nesting substrate is important. The literature shows that long fiber material such as shredded paper strips is the most structurally useful and naturalistic material for mice to build high-quality nests.²⁵ Compressed cotton is not an ideal substrate, as it is difficult for poor nest building strains to process, and it can cause eye lesions in nude mice from dust as well as constricting injuries to mouse pups.^26,27 Finally, the paper should be provided in amounts that allow mice to build a biologically relevant nest. At least 8–10 g is recommended, and mice will use more if provided.²⁸ To put this into context, this would be the same as providing 4–5 cotton nestlets, as each nestlet is ∼2 g.^29,30

Therefore, at UChicago, the default nesting enrichment for mice is shredded paper strips, and we do not provide cotton nestlets. Shredded paper is provided to every rodent cage as a default. Due to education on the need for nesting enrichment, we currently have no colonies of mice on campus with an IACUC-approved exemption to not provide this type of enrichment. In addition, we allow for adequate nesting material to build robust nests by providing 4 g of paper strips (Bed-r’Nest; Lab Supply, Fort Worth, TX) in addition to the twisted paper nesting material provided in the paper pulp cellulose bedding (ALPHA-dri PLUS; Shepherd Specialty Paper, Watertown, TN) (Figure 1). Finally, the ability to build a nest is used as a sensitive indicator of health and welfare at our institution, as nest building activity is altered when mice are unwell, in pain, or when aggression is present in the cage.³¹

Figure 1.

Example of Standard Mouse Cage with Enrichment. (A) A robust nest is created using the shredded paper strip enrichment (brown paper) with incorporation of the twisted paper nesting material from the paper pulp cellulose bedding (white paper). (B) Bird’s-eye view of the mouse cage with a robust nest and tunnel for handling.

Structural enrichment.

In general, increased structural cage complexity is beneficial and preferred by mice.^20,32 However, when adding resources to cages, it is important to ensure that the items meet the animal’s needs and preferences. This is especially important as each item requires money to purchase and time to deploy. Therefore, institutions should prioritize using a data-driven approach with animal-based outcome measures when selecting enrichment items. That being said, the success of using shelters (igloos, huts, and shacks) as enrichment tools is mixed.^18,33 It has been shown that some of these shelters may even cause negative consequences, including aggression, especially in male mice, as they become a resource to guard and defend.^34,35 At some institutions, these shelters may be used instead of nesting material as they are easier to use, standardized, and commercially available.¹⁹ However, at UChicago, we prioritize the use of nesting material, as mice have a strong preference for nesting material over shelters.²⁰ This is most likely because it allows the animals to have control over their environment by providing the ability to build their own nest.^18,20 In addition, nesting material does not typically result in the increased aggression that is sometimes seen with shelters.³⁵

At UChicago, we leave the tunnel that is used to perform refined handling in the cage as a home cage tunnel for structural enrichment. Tunnels, as structural enrichment, have been shown to not increase aggression.³⁶ The home cage tunnels provide more complexity to the cage as the animals can choose to go through the tunnel, walk across the top, or perch on top. Tunnels also work as a retreat in the event of an aggressive interaction, as well as allowing self-rescue if a cage floods. The increase in complexity of the cage allows a shelter for rodents to use, which is preferred by mice.²⁰ In addition, the home cage tunnel provides an improved and faster response to refined handling of some mice.³⁷

Bedding type.

Considerations for choice of contact bedding include animal preference and welfare, adverse health effects, breeding and research impact, absorbency and cage ammonia concentration, ability to visualize the animals, known chemical and microbiologic contaminates, and cost.^30,38,39 Corncob bedding has multiple limitations even though it is one of the most commonly used bedding types, as it disrupts sleep cycles and increases aggression levels since it contains estrogen disruptors.^40–42 The microbial contamination rate of the raw product is high, so it needs to be autoclaved or irradiated prior to use.^38,43,44 Molds can grow from this bedding, which may produce mycotoxins that cannot be removed by autoclaving or irradiation.⁴⁰ In addition, it may be ingested by animals, so it should be avoided when fasting is required or when being used for animals involved in microbiome research.⁴⁵ Wood-based bedding products are commonly used and also have multiple drawbacks including the risk of contamination by pesticides or heavy metals, variations from seasonal changes, and the fact that softwood shavings release aromatic hydrocarbons and phenols, which can cause liver toxicity.^38,39 There are also respiratory health concerns, especially due to the elevated dust content of this bedding.⁴⁴ This is in contrast to paper pulp cellulose, which has fewer adverse effects on rodents. More importantly, it is actually preferred by mice, which is presumed to be due to increased comfort since it is soft.^46,47 Paper pulp cellulose is associated the with the fewest deleterious health effects and contaminants, which leads to decreased lung pathology; and it has the lowest endotoxin and mycotoxin levels.^44,48 Furthermore, it improves breeding performance,⁴⁹ and many varieties of this bedding have additional twisted paper nesting material incorporated, which allows for additional species-specific nesting and foraging behaviors.

Due to these welfare and research benefits, we have transitioned to paper pulp cellulose as the standard bedding at UChicago. In addition, we have found improved operational efficiency and ergonomics for personnel since the bedding is lighter and more absorbent.^50,51 This has been particularly beneficial ergonomically for cages autoclaved out from our biosafety spaces, as the soiled paper pulp cellulose bedding does not adhere to the cage bottom as corncob bedding does, meaning cage scraping is easier for cagewash staff. Therefore, based on staff surveys, both cage wash staff and animal care technicians prefer the paper pulp cellulose bedding.⁵¹

Overall, the transition to the paper pulp cellulose bedding was straightforward; however, there are differences from the corncob bedding in practice that should be considered in the transition period. Importantly, in cages using water bottles with ∼1.5-in. sipper tubes, the level of the paper pulp cellulose bedding in the cage must be monitored. During changeover, we noted that in cages with water bottles, there was an increased risk of cage flooding due to wicking when the bedding level was too high. This was mitigated through changes to improve consistency in bedding levels when cages are set up. Cage flooding associated with the paper cellulose bedding has not been seen with automated water systems. In addition, if cages with paper pulp cellulose bedding are flooded, it rarely results in a health concern, as this bedding type is more absorbent, leaving less standing water, and dries significantly faster than with corncob bedding.⁵² Finally, while there are bedding dispensers specifically designed for paper cellulose bedding products available, we do not currently have access to these and instead chose to retrofit one of our existing bedding dispensers (model D236, BetterBuilt, Delta, BC, Canada) to accommodate the product, which has been successful. In the other facilities, we are currently hand-bedding.

Housing room temperatures.

The thermoneutral zone of mice is 26–34 °C (78.8–93.2 °F), which is the temperature at which they maintain their body temperature without needing to use energy above the normal basal metabolic rate.^30,53 Furthermore, preference testing shows that mice prefer temperatures around 25–30 °C (86 °F), which depends on the light phase and whether they are group- or single-housed.^53–56 However, the macroenvironmental temperature recommendation for rodents in the Guide for the Care and Use of Laboratory Animals is 20–26 °C (68–79 °F), and many animal facilities use the lower part of this range for their animal housing room temperatures, which is below the preferred temperature range and the thermoneutral zone.^23,57 This means that research mice are chronically subjected to cold stress, and some argue that cold stress is one of the most important extrinsic environmental conditions affecting laboratory mice.^30,55 This is due to the fact that chronic cold stress may effect changes in metabolism, immune function, and reproduction, all of which could potentially impact research.^22,30 These low recommended temperatures are particularly concerning for neonates and weanlings, nude and debilitated mice, mice housed singly, and those housed in IVC systems.³⁰

The current recommendations for housing room temperature are not based on the animal’s needs but are set to provide human comfort and avoid the high cost of maintaining facilities at relatively warm temperatures.^30,56,57 Therefore, numerous publications recommend revising room temperatures for mice to move closer to the lower limit of preferred/thermoneutral zone.^30,58–61 Based on this information, we increased the UChicago rodent housing room setpoint an additional 1.1 °C to 23.3 °C (increase of 2 °F to 74 °F) to balance personnel comfort with animal welfare. For the animal, the microenvironment is warmer, as the temperature in a cage of 5 mice with nesting and bedding material on an IVC system is usually 1 or 1.5 °C warmer than the room temperature.^30,57 In addition, it is important to provide adequate and appropriate bedding and nesting material for behavioral thermoregulation as described in the sections above.^30,55,62

Cage Change Modifications

At our institution, we transfer nesting material at every cage change, extend cage accessory change intervals (wire lids and filter tops every 3 months, watering valves every 6 months), and have switched to changing cage bottoms every 21 days using paper pulp cellulose bedding. Early cage changes are performed when specific criteria are met. This provides both welfare and operational benefits.

Scent transfer.

Olfaction is a very important sense for mice as it allows them to recognize social and mating partners, detect food, and avoid predators. The transfer of nesting material from the soiled to the clean cage allows for stabilization of olfactory cues.^63,64 This is thought to be because soiled nesting material contains hormones that inhibit aggression.^63,64 However, it is important to only transfer nesting, not soiled bedding, as scent marks in bedding may elicit aggression.⁶⁵ Therefore, at UChicago, for every cage on campus, we perform a nesting material transfer at every cage change. In fact, we also provide other scent transfers due to extending change-out of cage accessories.⁶⁶ We validated this extension of cage component change-out via an ATP luminometer, which is a measure of the amount of organic debris present and, therefore, an indicator of cleanliness. The results verified that the change-out frequency of the wire bar lid and filter top could be extended to every 3 months and the automated watering valve to 6 months.⁶⁶ Multiple other institutions also extend the change-out frequency of their cage accessories.^67–70 At our institution, this extension has also led to a decrease in personnel time of approximately 3,000 technician hours per a year for every 10,000 cages, which is a total annual labor savings of ∼$100,000 as calculated in 2023.⁶⁶

Cage change frequency.

Providing rodents with clean cages affects behavior, may cause aggression, may increase pup cannibalism, and leads to animal stress.^63,64,71,72 This is due to the fact that it disturbs scent marks, disrupts the social structure of animals and the physical nest, and decreases the social stability of groups.^64,71–73 Therefore, frequency of cage changes should be kept to a minimum to avoid disruption of olfactory cues in the cage environment.⁶⁴ At UChicago, we went one step further to investigate whether we could extend the cage change frequency from our current 14-day period to potentially decrease stress further.⁵⁰ We performed a study comparing the impact of 2 different mouse bedding types on the IVC microenvironmental parameters over a 21-day period and found that, based on ammonia levels, temperature, humidity, and an overall health score, the cage bottom change-out can be extended to 21 days for either bedding.⁵⁰ This extended cage bottom change frequency has also been noted at other institutions,^74,75 including a recent study that used machine learning/artificial intelligence to determine that cage change can be extended up to 3–6 weeks for average-sized mice, depending on the number of animals per cage.⁷⁶ Furthermore, we noted a difference in bedding type, so that the UChicago early cage change criteria may be met sooner for corncob bedding based on urine latrine size in the cages.⁵⁰ Therefore, paper pulp cellulose bedding decreases early cage change and allows for an extended change-out frequency, which may decrease animal stress.⁵⁰ Reducing the number of cage changes needed has additional benefits, as it decreases personnel labor and the cost of cage wash operations, decreases wear and tear of caging through cage wash, reduces bedding, enrichment, water and electricity use, and improves overall animal care efficiency.^50,76

Implementation of this change was mainly smooth, although a few investigators complained about the cages being overly soiled. Ultimately, we have addressed this by explaining that we have early cage change criteria adapted from the following study⁷⁷ that would prompt our staff to change the cage before 21 days if the criteria were met. Therefore, cages with 5 mice (especially a cage containing males) or breeding cages are likely to be changed before the 21 days period. In addition, we have referenced the data in our study and the known effects of cage change on rodents, as it is one of the more stressful procedures we perform on mice.^63,64,71 Finally, we have explained that in the wild, there is no cage change in underground burrows and that we have to ensure that we are not making decisions based on human cleanliness preference of a mouse’s microenvironment.

Animal Handling

At our institution, we promoted use of nonaversive handling techniques for mice through use of tunnels and cupped hands. We implemented tunnel handling for mice in ∼30% of cages and began converting unneeded water bottles into tunnels, increasing access to refined handling and reducing costs. In addition, we retrofitted transport carts with rubber wheels and gel pads, significantly reducing noise and vibration exposure during rodent transport. These changes help provide welfare benefits, while also having the potential to improve staff well-being.

Refined handling.

Removing mice from their home cage has traditionally occurred by lifting animals by the tail with forceps or hands. Picking up mice by the tail is aversive to the animal and increases anxiety levels.⁷⁸ Therefore, nonaversive handling involves the use of tunnel or cupped hands to pick up mice and is shown to benefit animals, personnel, and science.⁷⁹ In mice, refined handling reduces anxiety, depressive-like behaviors, chronic stress, and increases resiliency to subsequent negative stimuli.^{37,78,80–85} It has been shown to increase experimental test reliability, improve physiologic parameters and breeding, and increase voluntary interaction with humans.^{81,83,84,86,87} Ultimately, tunnel handling gives rodents a choice on when and how to enter the tunnel and be transported out of the cage, and it is known that providing choice is important for good welfare.⁸⁸ In addition, the increase in voluntary interaction with the handler is important for promoting human–animal enrichment and strengthening the human–animal bond, which can be more difficult to accomplish with mice than other animals. Furthermore, refined handling allows for psychological and emotional benefits for staff, in addition to safety benefits from fewer bite wounds.⁷⁹ It has also allowed our staff members to become proficient at a new technique, which improves their quality of life at work and employee retention. Therefore, refined handling improves human well-being, as staff members know they are using a technique known to improve animal welfare.^5,6,79

We implemented tunnel handling by adding this technique to several research colonies; those where there appeared to be an explicit need and those where the data have shown that this form of handling will improve animal welfare and research results. This involved colonies where investigators reached out for assistance due to poor breeding, high-anxiety mice, or those used in behavioral research. We used available online resources to introduce this implementation and provide training for staff.^89,90 Anecdotally, we have seen the mice in these colonies benefit from this refined handling method; for example, tunnel handling improved breeding parameters in a nonobese diabetic mouse colony and allowed for more consistent weight gain in mice fed a high-fat diet. In addition, it has provided additional opportunities for our animal care staff to learn new techniques. We have received positive feedback from the staff members using tunnel handling. In some instances, once the animal care staff are fully trained, they have trained the investigative staff on this technique. This has empowered the animal care staff and has improved morale, as they are able to explain the benefit to animals that can be achieved by use of tunnel handling. After receiving a grant to purchase additional tunnels, we have been able to increase the use of tunnel handling, as we were able to openly advertise the benefits of their use and provided the tunnels on a first-come basis. At this point, about 30% of our ∼20,000 cages of mice have tunnels present and use tunnel handling as we continue to expand use of this technique. In addition, we have recently begun the conversion of unneeded water bottles into tunnels (Figure 2). The mice use the converted water bottle tunnels in the same manner as the traditional tunnels. Reusing old water bottles as tunnels is an excellent example of recycling materials to reduce costs. This approach has enabled us to expand the implementation of tunnel handling across more cages within our institution and saved money.

Figure 2.

Tunnels for Handling Mice Were Created Using 16-oz Water Bottles That Were No Longer Being Used. (A) The tunnels were made by cutting the back of the water bottle off and smoothing the edges. (B and C) The original opening of the bottle was left intact, and mice freely move in and out of either side during tunnel handling.

Our husbandry technicians now have multiple options when handling rodents during cage change but are trained to perform tunnel handling if a tunnel is present in the cage. We use clear tunnels (mouse tunnel no. K3171; Bio-Serv, Flemington, NJ), and these allow conduct of a health check of the animals during tunnel transfer. If no tunnel is present, personnel cup the mice in their hands or pick up the mice by the tail or scruff using their hands or forceps. We have advocated for technicians to use their hands if no tunnel is present, since there is a higher incidence of repetitive motion injuries using forceps.⁹¹ Overall, we have also found that refined handling has psychologic, emotional, safety, and time benefits for personnel as seen at other institutions.^79,86,92

Transport cart modifications.

At many institutions, rodents are transported using plastic rolling carts between different facilities. Recordings at our institution and another institution revealed that the noise and vibration levels during cart transport are higher than the currently recommended standards in certain locations on the transport route.^93,94 Noise and vibration exposure can affect multiple welfare and research parameters in mice, such as breeding performance, immune function, cardiovascular parameters, and increase stress hormone levels.^30,93,94 To reduce animal exposure to noise and vibration, we retrofitted the plastic rolling transport cart with rubber wheels and plate casters and implemented the use of a gel pad under the shipping container or cage.⁹³ The combination of these 2 modifications reduced noise by 27% and vibration by 57% at our institution.⁹³ These are comparable modifications to those performed at another institution where they were able to reduce noise and vibration by the use of pneumatic wheels and under-cage padding with towels.⁹⁴ That study also described elevated stress hormones in mice exposed to higher noise and vibration during transport before these modifications; however, anxiety-sensitive behavioral tests displayed no influence depending on the differing transport conditions.⁹⁴

Operational Cost-Savings and Efficiencies

At our institution, we discontinued routine autoclaving of rodent cages after cage washing, now only autoclaving or irradiating cage items not processed in the cage washer (eg, bedding, nesting material), leading to reduced costs, labor, and environmental impact. In addition, we trained staff to measure and transfer unsoiled diet between cages at each change, cutting diet waste by over 50% annually.

Discontinue autoclaving of cage components.

Data show that the use of a cage washer that uses water temperatures of at least 180 °F (82.2 °C) prevents the transmission of multiple commonly excluded infectious agents to naive immunocompromised and immunodeficient mice.^95–97 These results suggest that autoclaving after sanitization is not necessary to prevent transmission of these agents to mice.^95–97 Nonetheless,the autoclaving of rodent cages after cage wash sanitization was included in the past at our institution out of an abundance of caution and due to it being a standard practice in the industry even though there was no evidence to support it. This is especially apparent since, after autoclaving, the cages are not maintained sterilely. Based on the current established prevalence data for rodent infectious diseases¹⁶ and our institution’s exclusion list,¹⁵ we determined that autoclaving of cage components was redundant. Therefore, we now only autoclave or irradiate all cage items that are not processed through the cage washer, including bedding and nesting material. This operational efficiency has decreased costs and labor, decreased wear and tear on caging, and saved on water and electricity use, which also has a positive environmental impact.⁹⁷

Diet conservation.

We performed an internal study assessing the amount of diet we discarded yearly by disposing of unconsumed diet at every cage change. We found that we wasted almost 200,000 lbs of diet annually, which accounted for >50% of the diet we purchased. This waste was an estimated $125,000 per year when the study was performed in 2014. Based on this, we trained our animal care technicians to use a scoop for measuring diet to avoid overfilling the wire bar lids. In addition, we assessed whether we could transfer the diet at cage change to the clean cage without discarding it. Based on the fact that mice consume 3–5 g of diet each day,⁹⁸ and an internal study where we assessed consumption in 8 cages of 4–5 male or female mice and calculated that mice consume 4 g of diet per day on average, we calculated that the uneaten feed would be minimal and could be reused on the new cage unless it was visibly soiled or contaminated. Even though the food would be reused, our process ensured that it would be consumed before the manufacturer’s recommended expiration date as also documented at another institution.⁹⁹

Conclusions

At UChicago, we were successful at accomplishing multiple practical, data-driven rodent welfare advancements by concentrating on improvements that go beyond basic health and allow for performance of highly motivated, species-specific behaviors and positive emotional states. Overall, these enhancements may result in improvements in animal welfare, research data, staff well-being, and efficiency, while also providing opportunities for cost-savings.

Conflict of Interest

The authors have no conflicts of interest to declare.

Funding

Work performed at the University of Chicago was internally funded by the University of Chicago Animal Resources Center RRID:SCR_021806. The tunnel handling grant was awarded by the Animal Welfare Institute.

Ethical review

All animal care and use were conducted in accordance with federal polices and guidelines and was approved by the University of Chicago’s IACUC. The University of Chicago has a PHS assurance with OLAW and is accredited by AAALAC, International.