Handling and restraint for minor procedures without anaesthesia

Techniques for handling and restraint of laboratory animals should be safe for both the handler and the animal. The methods used should also cause the minimum amount of distress to the animal. Stress and pain are known to alter body responses to a wide range of stimuli, hence in addition to important welfare considerations, efforts to reduce pain or discomfort to a minimum should also be considered good scientific practice.

The immediate response to handling is often an acute stress response, with increased heart rate and respiratory rate. Rodents will often urinate and defecate as a response to the anxiety caused by handling. In addition to these responses which can easily be observed, blood pressure will increase, blood glucose concentrations increase, and a range of other physiological responses (the “flight or fight” response) are triggered (Balcombe et al, 2004¹). Some of these changes may have longer lasting effects, for example, changes in gastrointestinal activity in rabbits and guinea pigs.

These changes will vary on each occasion that the animal is handled, and if the handling is undertaken in order to administer substances or obtain samples as part of a study, then they can result in unwanted variability in the study outcomes. To reduce this effect, animals can be trained to accept handling and restraint, for example by use of positive reinforcement each time the animal is handled. In most species, a relatively brief period of training (5-7 repetitions at daily intervals) will greatly reduce these responses, and may also enable minor procedures to be conducted with minimal restraint.

Modifying the handling technique can also have a major effect on the stress responses in some species. For example, it has been shown that picking up mice by the tail induces aversion and high levels of anxiety. Using tunnels or cupping the mice in open hands leads to more rapid acceptance of physical restraint. Habituation to this type of initial restraint persists even when mice are subsequently restrained more securely, for example by the scruff to allow injections to be carried out (Gouveia and Hurst, 2017², Hurst and West, 2010³) (National Centre for the Replacement, Refinement and Reduction of Animals in Research – www.nc3rs.org.uk/mouse-handling-tutorial ).

Adapting animals to handling and restraint by positive reinforcement, and by use of different methods of restraint need not add to the duration of a study. The adaptation can be conducted during the period of acclimatisation after animals arrive in the facility, prior to study, or are transferred from a breeding area and allocated to experimental groups.

This period of acclimatisation is important in order to reduce variability in the data that will be obtained from the study and ensure data is being obtained from animals that are physiologically normal. It will also minimise the impact of procedures on the welfare of the animals.

Factors that cause stress to animals (“stressors”) have additive effects. Allowing animals to recover from the stress caused by transportation or movement to a new environment and enabling them to adapt to their new environment and social group, and to the diet, bedding, etc, will often make them better able to adapt to the additional stress caused by experimental procedures. Some of the results of these stressors can be monitored easily – the initial stress associated with transport frequently results in body weight loss, which is regained rapidly over 1-2 days. Since changes in body weight are often used to monitor the effects of experimental procedures, it is clearly advisable to allow these non-specific effects to subside, and the animal to resume its normal growth rate or regain a stable body weight. It is generally recommended that between 7-14 days are allowed for this acclimatisation period, with the duration varying depending upon the species, and the type of study to be undertaken (Obernier and Baldwin, 2006⁴, Arts et al, 2014⁵).

In addition to adapting animals to handling and restraint to enable minor procedures to be conducted, it is also important to review the procedures to ensure they include appropriate refinements. For example, oral dosing of compounds can sometimes be avoided by training animals to ingest material voluntarily (Zhang, 2011⁶, Corbett et al, 2012⁷, Küster et al 2012⁸). Repeated blood sampling or intravenous injections may be avoided by placement of an indwelling catheter. The greater stress associated with catheter implantation may be offset by the reduced stress of sampling and the greater consistency in sample volume and quality. Repeated injections of material can be avoided by use of slow release pellets or osmotic minipumps. As with indwelling catheters, an assessment needs to be made of the relative impacts of, for example, daily intraperitoneal injections, compared to a minor surgical procedure to implant a minipump. When making this judgement, the greater reliability and consistency of minipump delivery of material should also be considered.

Adopting the measures described above not only increases the quality of data that is obtained, but also reduces the distress caused to animals. Implementing such refinements is a requirement of EU and UK legislation, so it is important to remain up-to-date with developments in this area. Websites such as the UK National Centre for the 3Rs (www.nc3rs.org.uk ) are valuable sources of current information.

When carrying out procedures, make sure you are wearing appropriate protective clothing, both to protect yourself from hazards such as allergens, and to protect the animal from an inadvertent transfer of potential infectious agents.

1. Balcombe, J.P., Barnard, N.D. and Sandusky, C., 2004. Laboratory routines cause animal stress. Journal of the American Association for Laboratory Animal Science, 43(6), pp.42-51.
2. Gouveia, K. and Hurst, J.L., 2017. Optimising reliability of mouse performance in behavioural testing: the major role of non-aversive handling. Scientific Reports, 7, p.44999.
3. Hurst, J.L. and West, R.S., 2010. Taming anxiety in laboratory mice. Nature methods, 7(10), p.825.
4. Obernier, J.A. and Baldwin, R.L., 2006. Establishing an appropriate period of acclimatization following transportation of laboratory animals. ILAR journal, 47(4), pp.364-369.
5. Arts, J.W., Kramer, K., Arndt, S.S. and Ohl, F., 2012. The impact of transportation on physiological and behavioral parameters in Wistar rats: implications for acclimatization periods. ILAR journal, 53(1), pp.E82-E98.
6. Zhang, L., 2011. Voluntary oral administration of drugs in mice. Protocol Exchange, 10.
7. Corbett A, McGowin A, Sieber S, Flannery T, Sibbitt B. A method for reliable voluntary oral administration of a fixed dosage (mg/kg) of chronic daily medication to rats. Laboratory animals. 2012 Oct;46(4):318-24.
8. Küster, T., Zumkehr, B., Hermann, C., Theurillat, R., Thormann, W., Gottstein, B. and Hemphill, A., 2012. Voluntary ingestion of antiparasitic drugs emulsified in honey represents an alternative to gavage in mice. Journal of the American Association for Laboratory Animal Science, 51(2), pp.219-223.