This site focuses particularly on the manual skills needed to carry out the different procedures humanely and efficiently, and on the need to handle animals carefully to reduce any distress caused by the procedure.

Administering substances to animals, for whatever reason, can have a significant impact on their welfare. If carried out incorrectly, not only can animal welfare be compromised, but the scientific goals of the study can be affected. If the administration is for a therapeutic purpose, then incorrect administration can lead to a failure of the treatment.

The selection of a particular route of administration must balance a number of factors – for example the volume and physicochemical properties of the substance, the required speed of onset, and other factors such as the degree of tissue irritation that could be caused. These topics are discussed in more detail in a number of different guidelines.

If a study requires repeated injection of substances the use of Osmotic minipumps may represent a significant refinement.

If you have not carried out an injection procedure before, you should first ensure you are familiar with the relevant anatomy of the species (by examining an animal euthanased for another purpose), and also develop basic skills in handling a hypodermic syringe correctly. A separate tutorial will be available soon to demonstrate correct use of a syringe. It is usually helpful to first practice injection on an inanimate object (e.g. an orange or a specially designed simulator). Before carrying out an injection technique yourself, you should observe and assist more experienced colleagues, then practice the technique on a dead animal (euthanased for another purpose). Only then should you carry out the procedure on a live animal.

Administration of some substances could result in adverse affects on the animal so it is important you know how to recognise these. Further guidance can be found at www.ahwla.org.uk and www.nc3rs.org.uk.

Remember that in the UK both your personal licence, and the relevant project licence, must authorise the procedure you are carrying out, and that it is not permissible to practice procedures on living animals.

All of the animals filmed or photographed were required to undergo the particular procedure as a necessary part of various research projects being undertaken in the UK.

This website and the resources it contains were developed with the support of the IAT and the NC3Rs.

Handling and restraint of the animal

Almost all laboratory animals can be restrained safely and humanely provided they are handled correctly. All animals benefit from being first accustomed to being handled, and this can be done during the period of acclimatization needed before they are used on a research procedure.

It has been shown that picking up mice by the tail induces aversion and high levels of anxiety. (Hurst & West 2010*) 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.

If injections are to be repeated on a regular basis, then familiarization with restraint assumes even greater importance, and training of the animal to co-operate with the procedure may also be helpful.

Restraint will be stressful, even in animals that have become accustomed to handling, so the duration of restraint should be minimized.

If you are relatively inexperienced in the technique that you plan to use, you should ensure you have an experienced assistant who can restrain the animal safely and humanely, and assist and supervise the procedure.

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 inadvertent transfer of potential infectious agents.

*J Hurst & R West (2010) Taming anxiety in laboratory mice. Nature Methods 7 (10), 825-842

Material can be administered orally using a number of different techniques, although gavage using a stomach tube is the most widely used. To minimise the risk of adverse effects associated with this procedure, it is important that the operator is skilled both in the technique and the restraint method needed.

Although gavage can be undertaken using rigid dosing cannulae, flexible catheters or tubes are preferred, and these are less likely to cause oesophageal trauma. Inadvertent dosing into the lung may occur, and this usually results in the animal showing immediate signs of respiratory distress. If this is observed, then the animal should be humanely killed.

As an alternative to gavage, some materials may be consumed voluntarily in palatable mixtures (e.g. flavoured gelatine) (Zhang, 2011*) (Küster, Zumkehr, Hermann, Theurillat, Thormann, Gottstein, Hemphill, 2012**). Material can also be dosed using a small flexible catheter introduced only into the animal’s mouth.

When possible, these approaches are to be preferred to dosing using a stomach tube, since they are not associated with complications, such as tracheal dosing or oesophageal rupture.

*Zhang, Lei, Voluntary oral administration of drugs in mice, Protocol Exchange, 2011. doi:10.1038/protex.2011.236. Published online 11 May 2011

**Voluntary Ingestion of Antiparasitic Drugs Emulsified in Honey Represents an Alternative to Gavage in Mice, Authors: Küster, Tatiana; Zumkehr, Beatrice; Hermann, Corina; Theurillat, Regula; Thormann, Wolfgang; Gottstein, Bruno; Hemphill, Andrew, Source: Journal of the American Association for Laboratory Animal Science, Volume 51, Number 2, March 2012 , pp. 219-223(5) Publisher: American Association for Laboratory Animal Science

Subcutaneous administration of material often causes minimal pain or discomfort, provided the material is non-irritant, has a near-neutral pH, and is not excessively hypertonic or hypotonic.

The most usual site for injection is over the shoulders, into the loose skin over the neck, but other sites with loose folds of skin can also be used, for example over the flank. When repeated doses of material are needed, varying the site of injection can help reduce the likelihood of local skin reactions.

It is not usually necessary to try to sterilize the skin with antiseptics – their use is almost always ineffective and they simply prolong the duration of restraint needed and may cause additional disturbance to the animal.

As with all injection sites, using a new needle for each animal, and injecting fluid that is at body temperature will reduce any discomfort caused by the procedure.

As with other routes, if repeated injections of material are needed, consider alternatives such as use of minipumps.

Intravenous administration of material can be technically difficult, and use of a restraining device is often required. These should be selected carefully to be an appropriate size for the animal to be injected.

Too small a device can result in injuries to the animal, and can interfere with respiratory movements. Too large a restrainer can also result in injury, caused by movements during restraint.

After use, restraining devices should be cleaned thoroughly, to avoid pheromonally-induced stress or cross-infection.

Since small movements of the animal can dislodge a hypodermic needle from the vein, alternative delivery systems may be found useful, especially in the rat. These will be described in more details on a future tutorial on catheter and needle use.

Although rapid buffering by the blood and rapid dilution can allow a wider range of materials to be administered, injection into a peripheral vein can result in irritation, and in some circumstances flushing with normal saline should be considered.

It is common practice to stimulate dilation of the tail veins in rats either by placing them in a warmer environment (e.g. at 28-30 ºC) for up to 30 minutes, or by placing the tail in warm (30-35 ºC) water. If a warming box or incubator is used, its temperature should be monitored carefully. It is good practice to place an electronic thermometer adjacent to the animal’s cage in the incubator, as the temperature registered by the device thermostat may not be accurate.

In the rat and other small rodents, the very small muscle mass makes intramuscular administration both technically difficult and painful for the animal because of the distension of the muscle.

If intramuscular injections are necessary, they can be made into the front or back of the thigh in all small rodents. In the rat and guinea pig the muscle mass is usually sufficient for accurate administration of small volumes of material, ideally 0.05ml or less.

It is not usually necessary to try to sterilize the skin with antiseptics – their use is almost always ineffective and they prolong the duration of restraint needed and may cause additional disturbance to the animal.

As with all injection sites, using a new needle for each animal, and injecting fluid that is at body temperature will reduce any discomfort caused by the procedure.

Material that is irritant or with a high or low pH can cause pain both during and following injection.

Material can be administered orally using a number of different techniques, although gavage using a stomach tube is the most widely used. To minimise the risk of adverse effects associated with this procedure, it is important that the operator is skilled both in the technique and the restraint method needed.

Although gavage can be undertaken using rigid dosing cannulae, flexible catheters or tubes are preferred, as these are less likely to cause oesophageal trauma. Inadvertent dosing into the lung can occur, which usually results in the animal showing immediate signs of respiratory distress. If such signs are observed, then the animal should be immediately humanely killed.

As an alternative to gavage, some materials may be consumed voluntarily in palatable mixtures (e.g. flavoured syrups, Leach et al, 2010*, Corbett et al, 2012**). Rats can also be trained to drink voluntarily from a syringe (see video on right) and this approach has been shown to be both effective and to result in accurate dosing (Atcha et al, 2010***). Material can also be dosed using a small flexible catheter introduced only into the animal’s mouth. When possible, these approaches are to be preferred to dosing using a stomach tube, since they are not associated with complications, such as tracheal dosing or oesophageal rupture.

*Leach et al (2010) Influence of preferred foodstuffs on the antinociceptive effects of orally administered buprenorphine. Laboratory Animals 44, 54-58
**Corbett, A, et al (2012) A method for reliable voluntary oral administration of a fixed dosage (mg/kg) of chronic daily medication to rats. Lab Animals 46, 318—324
***Atcha et al, (2010) Alternative method of oral dosing for rats. JAALAS, 49, 335-343

Although widely used as a means of administering substances, particularly injectable anaesthetics, this is an inherently unreliable technique, since inadvertent injection of some material into the gut, abdominal fat and subcutaneous tissues is a relatively frequent occurrence (Steward et al, 1968, Gaines Das and North, 2007)*. For this reason, it may be preferable to use other routes such as subcutaneous or oral administration.

It is not usually necessary to try to sterilize the skin with antiseptics – their use is almost always ineffective and they simply prolong the duration of restraint needed and may cause additional disturbance to the animal.

As with all injection sites, using a new needle for each animal, and injecting fluid that is at body temperature will reduce any discomfort caused by the procedure.

Material that is irritant or with a high or low pH can cause pain both during and following injection.

As with other routes, if repeated injections of material are needed, consider alternatives such as use of minipumps.

*Gaines Das, R and North, D (2007) Laboratory Animals, 41, 312-320
Steward et al. (1968) Applied Microbiology, 16 (9): 1418-1419.

Subcutaneous administration of material often causes minimal pain or discomfort, provided the material is non-irritant, has a near-neutral pH, and is not excessively hypertonic or hypotonic.

The most usual site for injection is over the shoulders, into the loose skin over the neck, but other sites with loose folds of skin can also be used, for example over the flank. When repeated doses of material are needed, varying the site of injection can help reduce the likelihood of local skin reactions.

It is not usually necessary to try to sterilize the skin with antiseptics – their use is almost always ineffective and they simply prolong the duration of restraint needed and may cause additional disturbance to the animal.

As with all injection sites, a new needle should be used for each animal, since this will reduce discomfort caused by the procedure and also reduce this risk of any injection-site infection. Any discomfort can be further reduced by injecting fluid that is at body temperature.

As with other routes, if repeated injections of material are needed, consider alternatives such as use of minipumps.

Intravenous administration of material can be technically difficult, and use of a restraining device is often required. These should be selected carefully to be an appropriate size for the animal to be injected. Too small a device can result in injuries to the animal, and can interfere with respiratory movements. Too large a restrainer can also result in injury, caused by movements during restraint.

After use, restraining devices should be cleaned thoroughly, to avoid pheromonally-induced stress or cross-infection.

Although rapid buffering by the blood and rapid dilution can allow a wider range of materials to be administered, injection into a peripheral vein can result in irritation, and in some circumstances flushing with normal saline should be considered.

It is common practice to stimulate dilation of the tail veins in mice either by placing them in a warmer environment (e.g. at 28-30 ºC) for up to 30 minutes, or by placing the tail in warm (30-35 ºC) water. If a warming box or incubator is used, its temperature should be monitored carefully. It is good practice to place an electronic thermometer adjacent to the animal’s cage in the incubator, as the temperature registered by the device thermostat may not be accurate.

Although widely used as a means of administering substances, particularly injectable anaesthetics, this is an inherently unreliable technique, since inadvertent injection of some material into the gut, abdominal fat and subcutaneous tissues is a relatively frequent occurrence. For this reason, it may be preferable to use other routes such as subcutaneous or oral administration.

It is not usually necessary to try to sterilize the skin with antiseptics – their use is almost always ineffective and they simply prolong the duration of restraint needed and may cause additional disturbance to the animal.

As with all injection sites, a new needle should be used for each animal, since this will reduce discomfort caused by the procedure and also reduce this risk of any injection-site infection. Any discomfort can be further reduced by injecting fluid that is at body temperature.

Material that is irritant or with a high or low pH can cause pain both during and following injection.

As with other routes, if repeated injections of material are needed, consider alternatives such as use of minipumps.