A checklist for good practice when caring for rodents in the laboratory.A checklist for good practice when caring for rodents in the laboratory.

In part due to a heightened awareness among the general public, and along with a global community of passionate ethologists and investigators concerned with animal welfare, this year the European Union (EU) legislated that, in law, animals are sentient beings. Globally, it raised awareness as well about the number of animals in use and the overall benefit of the animal studies.

The media are interested and the general milieu is one of guidelines and regulation. In many countries one must demonstrate in comprehensive project proposals the case for using animals and the methods that will keep animal suffering to the strict minimum. Most people define welfare this way: a state of balance between positive and negative experiences similar to those of the animals’ wild counterparts1.

It is encouraging, too, that guidelines for reporting out to professional journals—guidelines such as the ARRIVE Guidelines from NC3Rs2—have been powerful in drawing attention to treatment of animals used in research. The ARRIVE Guidelines are explicit as well as voluntary, calling on biomedical investigators to report details of design, experimental procedures, animals, housing, husbandry, sample size, allocation of animals to experimental groups, experimental outcomes, and statistical methods. And, reports of results, they say, must include baseline data, numbers analysed, outcomes and estimation, and adverse effects. In the UK, prominent groups have subscribed to the Guidelines and no doubt the culture will migrate. In other words, it’s impolitic to ignore the Guidelines though often inconvenient to subscribe to them, and the result is— when it comes to the welfare of animals in research—there’s nowhere to hide.

What’s important here? Animal welfare has taken its rightful place as a pivotal element in the design of biomedical investigations. All we’ve learned about the impact of emotion and environment on human health and well-being, learning, and memory will now influence, and in some places define, the way investigators and technicians manage non-human animals in research. With everything from housing to handling called into question, it would seem the industry may be in for an overhaul.

We were delighted, then, when ALN World™ invited us to set out a checklist for good practice, summarising what we know about how to provide for a rodent’s needs. What a very great pleasure indeed. We hope the items will become the animal welfare baseline for rodent care as well as a template for addressing needs of other animals in captivity for research.

Ideally the animal should feel secure in a complex, challenging, predictable environment that it can control. To help the animal achieve a sense of security, provide nestable, manipulable, and effective material plus hiding places and compatible cage mates.

Bedding Material
Provide bedding that is comfortable for resting. Sawdust is a popular choice though fine particles can cause preputial and respiratory problems. The type of wood can affect physiological parameters; for example, soft wood may affect microsomal liver enzymes3. Choose hard wood or other material with no side effects. Avoid hygroscopic material like cat litter; it may cause dehydration in a newborn.

Bedding Provision
All laboratory rodents spend considerable time manipulating bedding and often creating tunnels and building nests for shelter and warmth. Make sure there is sufficient depth as well as the right material.

Bedding and Odour
Bedding must absorb urine. Mice use bedding to deposit odour patterns. They claim territory through urinary scent marking. Proteins specific to individuals prepare a male mouse to identify his own and another male’s urine scent. However, too much urine in crowded conditions can unnaturally extend oestrous cycles.

Change the bedding about once a week. That is a general rule, though the specifics of the research design will provide a more discreet guideline. Whatever the design, though, there should be a balance between increasing the animal’s anxiety and aggression by removing familiar odour patterns on the one hand and minimising the build-up of harmful ammonia on the other. It has been shown that olfactory cues from nesting and bedding material affect aggression in a different way: transfer of nesting material reduces belligerence whereas sawdust containing urine/faeces intensifies aggression, meaning that different pheromones may be involved4.

In general, bedding and nesting material should be:

  • in accordance with the mouse’s needs;
  • nontoxic or not in any other way harmful to the animal;
  • absorbent, but not dehydrating for neonates; and
  • relatively dust free.

Cage Design
In the past, housing was something of a conundrum. Keeping cage design constant seemed a reasonable way to minimise uncontrolled environmental effects. However, we’ve since learned that the current standard rodent cages—too often translucent plastic boxes that are left barren instead of kitted out with the makings of nests, tunnels, and barriers—create stress, lead to abnormal behaviour such as stereotypies, and probably interfere with the development and function of brain and behaviour. So, what’s designed to reduce experimental bias may ironically be a primary source of pathological artefacts. The guideline? Remember that mice take charge of their micro-environment. They huddle and build nests in order to adjust the temperature—which influences such functions as reproduction, food, and water intake and blood parameters—as well as humidity and light which each have similar impact on the animal’s physiological and psychological status.

There should be enough provision in the cages for the mice to partition, as they create separate spaces for sleeping, defecating, urinating, eating, and play. Consider the number of animals in the cage when providing opportunities to climb and to hide.

In order to avoid as much as possible the frustration, suffering, and stereotypies engendered by environments that restrict natural behaviour, the cage should have a solid floor with bedding for digging, nesting, and resting. Include a wide range of stimuli including other animals and lots of opportunity to burrow.

Avoid grid flooring unless you must use it in order to continuously collect urine and faeces for research purposes. However, allow metabolic cages only for a restricted time period and include a small solid floored box to rest and hide. As animals generally don’t soil their nests, you can still collect all urine and faeces.

Cage Size
What is the optimum cage size? We now know that what’s important about a cage is not the size but rather complexity; that is, how you equip it. Mice are thigmotaxic; that means they design their homes with barriers and walls rather than an open plan.

You can keep the cage size adequate if not generous but make sure that what’s inside—nest boxes, tubes, and partitions, for examples—will keep the animals happy.

The environmental materials the animals use to create their spaces—materials like tissues, paper, and wood wool—must be sufficient for lots of activity, economical to use, clean, disposable, and of course inedible.

The recently revised EU Directive 86/609/EEC/ Annex II, Guidelines for Accommodation and Care of Animals concerning space might come in handy here. It states “Any restriction on the extent to which an animal can satisfy its physiological and ethological needs shall be limited as far as practicable” and stressed that such limitations should be kept to a minimum.

In the U.S., the National Research Council’s Guide for the Care and Use of Laboratory Animals recommends allowing housing of mice in harmonious groups, and research demonstrates that increasing the complexity of the cage is more important than increasing floor area, as structures provide opportunity for activity and increase usable space.

Individually Ventilated Cages
Urine marking is a must for maintaining homeostasis and a rodent’s sense of harmony with the natural world. When animals mark within a caged existence, two bedding changes a week, no more, simulates a macrobiotic setting. However, urine means ammonia which can unsettle technicians, and in a crowded cage can cause respiratory tissue infection in the animals.

The ammonia matter, among other husbandry challenges, is satisfactorily addressed by individually ventilated caging. An individually ventilated cage—an IVC—is a self-contained environment where air circulates from a supply direct to each cage rather than from the ambient air. Individually ventilated cage (IVC) systems were developed to maintain low ammonia and CO2 concentrations, to support a low relative humidity, and to reduce spread of infective agents and allergenic contaminants5. An air filter, typically a HEPA filter, keeps the air relatively pure. Frequent air changes, optimally about one a minute and cage changes every fourteen days, keep the ammonia within the cage at levels satisfactory to the animals.

However, IVCs can present dilemmas and challenges.

Air source is subject to a delivery system that typically supplies a large cage rack. If the supply goes down, even for a short time, NH3 and CO2 build up affecting the animal’s heart rate, blood chemistry, and stress levels— sometimes detected only by running blood analyses and therefore disrupting both the animal and the experiment long before observable behaviour alerts you to problems.

Furthermore, the air filters can become clogged with dander and dust before it’s time for a filter change, so staff must keep vigilant.

Research shows that in an IVC system, mice avoid high intracage ventilation rates (mitigated by sufficient nesting material) and that they strongly prefer air supply in the cover. However, air supply in the cage top can result in dead air pockets or ineffective and incomplete air changing. So there is a dilemma still because the animals prefer air changed at the top while air change at the animal level exchanges air better and more completely.

To make best use of IVCs:

  • Back up the air supply source with a battery pack and keep an emergency generator.
  • Run maintenance checks weekly and service regularly.
  • Change filters frequently enough to keep out dust and dander.
  • Establish a monitoring system to detect need for filter change.
  • Keep air changes to eighty per hour maximum; rodents prefer fifty.
  • The air speed should be between 0.3 m/s and 0.5 m/s.
  • Choose IVC systems that deliver air flow above the animal level.
  • Provide nesting material and/or shelter to protect against draught.

Cage Mates
Mice are gregarious, so as a general rule keep them in groups but with the dominance hierarchy in mind. Even living alone together—two mice in one cage but separated by a grid partition—actually exacerbates the stress of solitary existence 6.

Both mixed sex and single sex groups offer opportunities and pose problems. In the wild most rodents live in mixed sex groups though breeding frequently triggers agonistic events like threat, attack, appeasement, or retreat. If the individuals are not strangers, life is calmer. Male mice prefer social contact even at rest, but beware: keep no more than three males per cage in cases of more aggressive strains such as BALB/c mice4.

Judicious use of used nesting material—though not bedding material—can reduce aggression. And keep a close watch on barbering. It can be a sign of boredom or inappropriate living conditions7.

Here are some tips and tricks for feeding:

  • Mice are omnivores. Their incisors and molars wear down and regrow with use. Check that each mouse has the required teeth in good order; anything less and you risk under-nutrition or malnutrition.
  • Mice eat most of their daily ration in the dark. Disturb their routine and you risk unwanted variation in experimental results.
  • Keep the food rack pretty full as mice like to eat from a full plate. However, mice will eat whenever there’s food around so you want to pace delivery in order to control weight. Unless it interferes with research protocol, scatter food such as grain through the bedding. This will help meet the mouse’s need to forage and also will prevent boredom. Foraging this way plus lots of bedding may show increase in weight over animals in less enriched environments.
  • The food hopper should be between 3 mm and 5 mm; mice can escape through space in a food hopper that is greater than 5 mm. A diet board can give rodents the opportunity to forage, which will preempt boredom and obesity8.

Try to avoid handling a postpartum female or her litter for several days after birth. Handling and transporting at this time could cause the mother to reject her young or eat them. Fortunately, you can reduce the risk of such trauma by taking note of these hints:

  • Wear plastic gloves, to eliminate human scent.
  • Place the dam in a separate cage while the litter is handled.
  • Rub the pups with bedding material from the home cage.

Taking a mouse by the tail is a common way of lifting the animal. It is acceptable though not optimal because it is very stressful and creates antagonism toward the handler. While the mouse is inside the cage, take the base of the tail between your thumb and index finger, then pick the animal up by the tail. Support the animal’s weight on your arm or the cage floor; never allow the animal to dangle.

Other methods for routine handling are to:

  • Cup the mouse in your open hand, but note: the mouse will try to escape (and will sometimes succeed).
  • Handle while inside a home cage tunnel.
  • Collect the mouse using the tunnel and transfer to your open hand.

To restrain the mouse, place the animal on a rough surface such as a cage lid. Hold the loose neck skin between your thumb and index finger, and then lift the mouse securing the tail between the fourth or fifth finger and palm of the same hand. Your other hand is free for injections or other procedures. The colour of the nose and mouth mucous membranes will let you know whether you’re holding the skin too tight. To pick up newborn or mice less than two weeks old, take the loose neck/shoulder skin into thumb and index finger or cup the pup in your open hand.

To reduce the animal’s anxiety, be sure you are confident and competent, and equipped to work efficiently.

In time the offspring of captive animals adapt to the lab environment. However, were they free to do so, they would behave much as their conspecifics do in the wild. They would forage for a wide variety of food. They would build nests designed to accommodate the specifics of their broad behavioural repertoire and the intricacies of their complex social structure. The drive to live out their natural behavioural patterns is never far from the surface.

In order to care for animals and provide for their welfare, consider all elements of the lab animal’s environment and benchmark against what they would probably do in the wild. The elements of housing are also stimuli, affecting not only their observable behaviour but the feelings that motivate it.

Paper towels, tissues, and wood wool are good; they allow the animal to create shade as well as the chance to regulate microclimates, such as temperature. They allow the animal to create a shelter in order to hide, and generally to control the social environment. Provide enough nesting material for all laboratory mice, not only for breeding females, to build a complete covered nest. Nest boxes are popular alternatives to nesting material but mice always prefer nesting material, like paper.

Upon release, even animals raised in a cage seamlessly take up life in the wild. That fact helps to explain why environmental factors play a major role in helping to mimic freewheeling existence within a cage—with noise and light near the top of the influence list. For example, even hushed sounds can have a considerable impact on animal physiology and behaviour, because they engage limbic structures and higher brain centres involved in determining context and meaning. Sounds that startle us may not startle the animals, and in day to day lab activity we may create sound we can’t hear that causes the animal serious distress.

Early exposure at critical developmental times can influence sensitivity. Key times are opening of the ear’s meatus at twelve to fourteen days after birth, and ten to sixteen days after birth, when proper development of several auditory system parts require acoustic stimulation.

Ultrasound wavelength is short and does not easily pass through barriers such as plastic cages though it will pass through the grid on top. However, during cage changing there might be inter-cage communication and alarm calls to conspecific animals attempting to communicate a stressful or frightening situation and ultrasound noise from rattling metal, running water, and other sounds of the vivarium environment.

Be aware that the everyday quiet of lab life may be a noisy place for the animals, where even outdoor noise may raise the levels of stress hormones and interrupt all basic behaviours. The influences will usually be subtle and beyond our own detection, so it’s essential to consider what you don’t see or hear.

The optimal temperature range for mouse rooms is 20–26°C. Choose the upper end of this range, 22–24°C if the mice are hairless or nudes, or if the demands of your research design mean the mice have only little bedding or nesting material. Different strains show some variation in temperature preference, but typically the deciding factor is the nature of the floor.

Stress engendered by transport may endure for up to five days and affects physiologic processes such as immunity and reproduction.

To move a mouse a short distance within the facility, set the animal in a clean cage or cardboard carton. For longer trips, use a cardboard container that is coated with moisture proofing material. There should be two ventilation openings, one on either side of the container. Cover the opening with steel wire mesh and filters. Provision the container with bedding, as usual, and with pellets, water-rich fruits, or vegetables such as apples, or use commercially available “solid water” gel.

Autoclave filled water bottles and drinking tubes but stay mindful: autoclaving doesn’t prevent contamination by the animal, whose person may carry contaminants to the tubes or who may push faeces and bedding into the drinking tube.

Control bacterial contamination by acidification (pH 2-3) or chlorination (15 – 20 parts per million active chloride). Remove trace elements, heavy metals, and organic chemicals by reverse osmosis, deionisation, or microfiltration. Offer water in a bottle attached to the cage and change it at least once a week.

Automatic watering systems may appear to save time but there’s more to it than you may imagine. Monitor the water pressure continuously to ensure that the flow is adequate and that the valve isn’t stuck. In this way you avoid both flooding and dehydration. Furthermore you must clean the central reservoir regularly and check for contamination with bacteria and fungi.

The term euthanasia is derived from the Greek euthanatos: eu meaning easy and thanatos meaning death9. Whether chemical or physical, however, no endof- life option is perfectly pain free. CO2 delivered at high concentration brings quick but painful death. Cervical separation requires highly experienced handlers. A guillotine must be kept razor edged and run by highly expert operators. Microwave is a specialised euthanasia technique that brings about a very quick death but may cause stress, as you use a restrainer in order to position the animal. Anaesthetising the animal before killing it may practically eliminate consciousness and so the experience of stress and pain.

Practice all physical methods on dead animals and get the OK from a senior, experienced technician or investigator before attempting to euthanise an animal. All physical methods should be followed by exsanguinations in order to confirm death. Demonstrate expert knowledge about every chemical method available to you—and have your knowledge base confirmed by an expert—before taking it upon yourself to euthanise an animal.

We have reserved discussion of environmental enrichment for the next article, “Expression of Species-Specific Behaviour”, which will be featured in the May/June issue of ALN World.

For discussion of laboratory procedures, see The Laboratory Mouse (Chapter 21) in Robert Hubrecht and James Kirkwood (eds). The UFAW Handbook on the Care and Management of Laboratory and Other Research Animals, 8th Edition, Wiley-Blackwell: April 2010.


  1. Bauman, Vera. “Science-based assessment of animal welfare: laboratory animals”. Rev. sci. tech. Off. int. Epiz. 24.2 (2005): 503-14.
  2. Kilkenny, C, WJ Browne, IC Cuthill, M Emerson, and DG Altman. “Improving Bioscience Research Reporting: The ARRIVE Guidelines for Reporting Animal Research”. PLoS Biology. 8.6 (2010): e1000412.doi:10.1371/ journal.pbio.2000412
  3. Vessel, E.S. “Induction of drug-metabolizing enzymes in liver microsomes of mice and rats by softwood bedding”. Science. 157.3792 (1967): 1057-58.
  4. van Loo, PLP, CLJJ Kruitwagen, van Zutphen, LFM et al. “Modulation of aggression in male mice: influence of cage cleaning regime and scent marks”. Animal Welfare. 9.3 (2000): 281-95
  5. Baumans Vera, Schlingmann Freek, Vonck Marlice and Van Lith Hein A. “Individually Ventilated Cages: Beneficial for Mice and Men?” Contemporary Topics, Jan 2002
  6. van Loo, Pascalle, Nynke Kuin, LP, et al. “Impact of 'living apart together' on postoperative recovery of mice compared with social and individual housing”. Laboratory Animals. 41. (2007): 441-455.
  7. Garner, JP, BI Dufour, Gregg, LE, et al. “Social and husbandry factors affecting the prevalence and severity of barbering (whisker trimming') by laboratory mice”. Applied Animal Behaviour Science. 89. (2004): 263-82
  8. Kasanen, IH, and KJ Inhila. “The diet board: welfare impacts of a novel method of dairy restriction in laboratory rats”. Laboratory Animals. 43.3 (2009): 215-23
  9. Merriam Webster Online Dictionary


Additional Reading

  • Bishop, Melanie J., and PFD Chevins. “Urine odours and marking patterns in territorial laboratory mice (mus musculus)”. Behavioural Processes. 15.2-3 (1987): 233-48.
  • Castelhano-Carlos, MJ, and V Baumans. “Noise and Light in the Vivarium”. ALN World. January/February 2010:12-19.
  • Clough, G, J Wallace, MR Gamble, ER Merryweather, and E Bailey. “A positive, individually ventilated caging system: a local barrier system to protect both animals and personnel”. Laboratory Animals. 29. (1995): 139-51.
  • Conlee, KM, ML Stephens, AN Rowan, and LA King. “Carbon dioxide for euthanasia: concerns regarding pain and distress, with special reference to mice and rats”. Laboratory Animals. 39. (2005): 137-61.
  • Feoktistova, Natalia Yu, Svetlana V Naidenko, and et al. “The Influence of Predator Odours and Overcrowded Mouse Odours on Regulation of Oestrous Cycles in House Mice (Mus Musculus)”. Wildlife Damage Management, Internet Center for USDA National Wildlife Research Center -Staff Publications. (2003): 173- 175.
  • Hawkins, Penny. “Recognising and Assessing Pain, Suffering and Distress in Laboratory Animals: A Survey of Current Practices in the UK with Recommendations”. Laboratory Animals. 36.4 (2002): 378-95
  • Hubrecht, Robert, and James Kirkwood. The UFAW Handbook on the Care and Management of Laboratory and Other Research Animals. 8th ed. Wiley-Blackwell, 2010.
  • Krohn, Thomas C. Pros and Cons of IVC systems (PPT) 20and%20Cons%20of%20IVC%20systems.pdf
  • Krohn, Thomas C, and Axel Kornerup Hansen. “Carbon dioxide concentrations in unventilated IVC cages”. Laboratory Animals. 36. (2002): 209-12.
  • Krohn, Thomas C., Axel Kornerup Hansen, and Nils Dragsted. “The impact of cage ventilation on rats housed in IVC systems”. Laboratory Animals. 37. (2003): 85-93.
  • “Mice Cages Can Alter Rodent Brains.” ALN. July 28, 2010.
  • Nevison, CM, SR Armstrong, J Beynon, RE Humphries, and JL Hurst. “The ownership signature in mouse scent marks is involatile”. Proc. Biol. Sci. 270. (2003): 1957-63.
  • Pham, TM, B Hagman, et al. “Housing environment influences the need for pain relief during post-operative recovery in mice”. Physiology & Behavior. 99. (2010): 663-68.
  • Rice, AS, D Cimino-Brown, and et al. “Animal models and the prediction of efficacy in clinical trials of analgesic drugs: a critical appraisal and call for uniform reporting standards”. Pain. 139.2 (2008): 243-47.
  • Stoke, EL, PA Flecknell, and CA Richardson. “Reported analgesic and anaesthetic administration to rodents undergoing experimental surgical procedures”. Laboratory Animals. 43. (2009): 149-54.
  • Webster, J. “The Assessment and Implementation of Animal Welfare: Theory into Practice”. Rev. sci. tech. Off. int. Epiz. 24.2 (2005): 723-34.
  • Wurbel, Hanno. “Ideal homes? Housing effects on rodent brain and behaviour”. Trends in Neuroscience. 24.2 (2001): 207-11.

Vera Baumans is a veterinarian who holds a PhD in Veterinary Anatomy. She was Chair, Laboratory Animal Science at Karolinska Institutet in Stockholm, Sweden for six years and retired as Animal Welfare Officer at Utrecht University, The Netherlands. Vera organises and teaches Laboratory Animal Science courses in many countries. Her research field is Environment, behaviour and well-being of laboratory animals. She is a founding member of the Veterinary European College of Laboratory Animal Medicine.

Helen Kelly is a freelance journalist covering management practice, management science and biomedical sciences internationally. She divides her time between Boston, Massachusetts USA and London UK.