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Restraining techniques used in laboratory animals

In biomedical research, rodents play a central role as animal models for genetic disorders and diseases. Animals perform well if they are accustomed to handling. However, they are not easily tractable, so researchers need hands-on training and specialized equipment for handling and restraining the laboratory animals. Improper management could result in stress, discomfort, and injury to the animal as well as the experimenter (Scobie-Trumper, 1987). Proper handling procedures should not only be considered within the remit of experimental work but also a part of routine husbandry procedures to make the animals comfortable (Thomas & Tilla, 2004).

Restraining is the procedure of immobilizing the animal to avoid movement. Restraining is done by safely holding the body or body parts of the animal manually or with the help of a physical device. Animals are restrained for brief periods, usually minutes, to administer drugs under study or to collect blood samples for investigations. Conscious animals require physical restraint during the experiment. You can use anesthesia and sedation besides manual control. (Thomas & Tilla, 2004). Correct handling and restraining procedures are touchstones for a successful experiment. This document provides a detailed insight into techniques used to restrain laboratory animals.

Manual Restraint

Handling and restraining procedures vary depending on animal’s species, size, age, strain and temperament (Machholz, Mulder, Ruiz, Corning, & Pritchett-Corning., 2012). However, all the procedures have a common underlying principle that they should not cause pain, stress, or discomfort to the animal.

One-handed manual restraining technique for mice consists of following steps:

One-handed restraint

Noise or disturbance in the laboratory room could disturb the animal behavior which could alter experimental findings and associated behavior. The animal is safely taken out of its cage and put on wire-bars or a rough surface for grasping. Gently approach the head from the back and firmly grasp the skin between the thumb and index finger. Grip the loose skin behind the neck to avoid turning of the head, while keeping the tail of the animal under little finger of the hand holding neck of the animal.

Note: Approach the animal with gentle, but firm hands. Too soft of a hand could set the animal free while too harsh of a hand could result in injury or stress.

In addition to one-handed mouse restraint, biomedical research uses two-handed restraint technique for mice.

Two-handed restraint

Lift the mouse by the tail and place it on a cage or a rough surface, then gently pull the mouse from the back so that it can grasp the wire bars with fore-paws. And quickly grab the subject by the scruff of its neck with the other hand. Gather the loose skin at the back of the neck to limit the animal. Now you can observe or inject the subject as per experimental requirement.

Note: Two-handed manual restraint is preferred for an inexperienced or a new experimenter to instill confidence in handling and restraining laboratory animals (Suckow, Danneman, & Brayton., 2000).

Rats are relatively larger and less cooperative than mice. They can also be handled manually by scruffing, gripping shoulders, or by holding from under the shoulders (Sharp & Villano., 2012).


As rats are less receptive to scruffing than mice, so this procedure is limited to smaller rats. First, hold the rat form its tail and gently pull it back on a rough surface. Second, clutch the rear end firmly and approach the animal from the scruff. Apply gentle pressure on the back and grasp the scruff from the skull base between palm and fingers of the hand.

Note: Rat’s head should be controlled with caution as its bite can cause serious injury. The abovementioned holding may vocalize the animal.

Gripping over the shoulders

With your dominant hand, remove the rat from its cage and place it on a rough surface by grasping it from the tail. Approach the head from the back with the help of non-dominant hand. Hold the rat around the thorax with thumb, pinkie and ring finger by keeping its head between middle and index finger. In this position, the animal could be injected or observed.

Note: Thorax should not be compressed firmly.

Gripping under the shoulders

Place the rat on a rough surface and grasp its tail. Gently place the non-dominant hand on its back approaching from the caudal side. Hold the rat’s thorax under the shoulder blades and gently push its forearms with the thumb and index finger. Observe or inject the rat in this position.

Note: Fore-arms should cross rat’s chin to avoid biting and turning of the head. Do not compress the thorax.

Ferret Restraint

Ferrets are usually aggressive and non-cooperative. They can be restrained by grasping from the neck and the shoulders. Hold the ferret with one hand under the shoulders by placing the thumb under the jaw and support the caudal part with another hand. You can now inject the animal or collect its blood for scientific investigations (Assessing the Health and Welfare of Laboratory Animals).

Guinea Pig Restraint

Guinea pigs are docile and easy to handle relatively. However, they get startled and disturbed easily. So they should be approached gently and softly to avoid frightening (Species-Specific Information: Techniques for Handling, Sexing, Injection, and Blood Collection). Do not move unnecessarily or make noise. Place the thumb under the jaw of the guinea pig. Support the caudal quarter with another hand. Another person should collect blood or inject the animal.

Rabbit Restraint

Like guinea pigs and ferrets, rabbits also need adequate handling and restraining procedures for experimental manipulations. As if not controlled, their hind legs can cause serious spine injury to them. Also, the weight of the rabbit is relatively higher than that of rats and mice (Rabbit – Sexing, Handling, and Restraint, 2017).

Check the condition of the rabbit before the experiment. Approach the animal with confidence as rabbits get stressed easily. Firmly grasp the rabbit from the nape of its neck. With the help of the second experimenter, firmly and gently place the hand on rabbit’s back. If a single person is handling, then a towel or a cat bag with zipping can be used to restrain the rabbit firmly.

Manual restraint is recommended for experienced and trained individuals, for brief periods usually for minutes. However, if experiments involve complicated and prolonged manipulations then, plastic or acrylic restrainers are suggested.

Mechanical restraint

Restraining devices for rodents including rats, mice, guinea pigs, rabbits, etc. are being continuously developed to facilitate scientific investigations. These devices are usually made up of acrylic, high-quality plastic, glassware, macrolon or metal. A variety of designs are available depending upon the type, strain, temperament, and size of the animal. Size of the restrainer should be a prime consideration; the larger device can set the animal free while smaller can cause suffocation and breathing problems. Duration of restraint should also be considered, as most restrainers do not allow full body heat dissipation and consequently, raise the body temperature. One of the significant advantages of using device restrainers is that they enable the researcher to use both their hands.

Different types of restrainers include Guinea pig restrainer, Bowman restrainer, Ferret restrainer, Soft restrainer, Rabbit acrylic restrainer, Flat bottom restrainer, Broome style restrainer, etc. However, underlying protocol is usually similar (Donovan, Brown, Reeves, & Reeves, 2003). Procedure for restraining the animal with the help of a physical device is described below:

Plastic/glass restrainer

Put the restrainer on a flat surface and clean its parts. And lift the animal up by gripping its tail. Place the head of the animal near restrainer’s entry groove. After the introduction of the animal in the device, gently push the animal from its back and then pull it back to motivate it to enter the device. After releasing the thorax, maintain tension on the tail to move the animal further. In the end, expose the rear leg or tail of the animal after closing the restrainer for blood collection or drug administration.

Note: While handling rabbits, keep in mind that rabbits get stressed if placed in a closed box or restrainer for extended periods.

Restraining wall

The restraining wall is another example of restraining devices for the laboratory animals. The animal is grasped from the tail and lifted up for entry into the wall. Position the subject in front of the wall by keeping its tail outside the slit. Gently pull the animal back to motivate it for further escaping beyond the fence. Close the slit while holding the tailpiece outside. Now the animal is ready for experimentation like blood collection or injections.


Decapicone is made up of a thin flexible plastic designed like a small cone with a small hole at one end. To restrain the animal in a decapicone, firstly, it is put out of the cage and lifted up, then placed in the decapicone of appropriate size. The animal is pushed forward to enter the plastic sheet by pulling the tail while keeping the tailpiece out of the groove for experimental procedures or injections.

Decapicone is useful as thin plastic allows the injections through it. However, its significant disadvantage is that animals may overheat because of suffocation and poor dissipation of heat.

Chemical restraint

To avoid pain and distress, some researchers use anesthetic agents to restrain the laboratory animals in research labs. These agents can be used with adjuvants to get desired effects.

Dosage, intensity, and duration of these agents depend upon the animal’s size, age, nutritional health, sex, genetic makeup and environmental conditions. Anesthesia is usually given to the animal through intraperitoneal (IP) injection because it is regarded as a safe route for uniform absorption (Wixon & Smiler., 1997). The solution should be diluted 5-10 folds before the infusion depending on the size of the animal.

Injectable anesthesia

The protocol followed for anesthetizing the laboratory animal through intraperitoneal injection consist of following steps:

  • Calculate the weight of the animal.
  • According to the calculated weight, prepare the appropriate dose of anesthetic injection for the animal (for 15g mouse, 100 mg ketamine/20 mg xylazine).
  • Restrain the animal manually or with the help of a physical device.
  • Inject the mixture with the help of an injection intraperitoneally.
  • Return the animal to its cage safely and observe the effects of the anesthetic agent.

Note: Respiration rate and depth are two commonly used indicators for assessing the effects of compounds on animals. Rapid or irregular breathing pattern indicate inadequate dosage while slow breathing suggests an appropriate dosage of anesthetic compound and adjuvant. Toe pinch can also initiate reflex extension of hind limb until adequate anesthesia is achieved. In rabbits, self-mutilation indicates positive anesthesia. In case of rabbits, anesthetics are used with extreme care as rabbits have caecum which can act as a reservoir for anesthetic agents (Species-Specific Information: Techniques for Handling, Sexing, Injection, and Blood Collection). Instead of atropine, glycopyrrolate can be used for rabbits.

Inhalant anesthesia

Anesthetize the laboratory animals through inhalation of methoxyflurane, ether, and isoflurane. However, for rodents, isoflurane and ether are not recommended as ether is flammable, explosive, and irritating. And isoflurane is lethal at even low concentrations. That’s why for rodents methoxyflurane is used. Methoxyflurane is flammable, so all procedures involving its use should be done in a safety cabinet or hood   (White & Field.). First, prepare a sponge gauze or cotton sponge soaked with methoxyflurane. Second, place the soaked sponge on a raised floor to vaporize it and put it in the one end of a container. Third, set the animal on another end of the tank and allow it to move further in the container.  Fourth, enable the animal to reach the sponge and breathe normally, then remove the animal from the tube. On the other hand, the induction chamber provides a more sophisticated inhalant anesthesia technique.

Restraining techniques and anesthetic agents are used on laboratory animals to allow blood collection or drug administration for scientific investigations and pharmacological studies. Researchers should know about the chemical and physical properties of the injectate. Consider concentration, pH, viscosity, pyrogenicity, and sterility before preparing injections. Needles and syringes should also be autoclaved or sterilized to avoid infections.  There are different administration routes for rodents. These routes are mentioned below:

  • Intramuscular
  • Intradermal
  • Intraperitoneal
  • Subcutaneous
  • Intravenous
  • Footpad injection
  • Oral route (Gavage)

The success of the research depends on proper handling, restraint and stress management of the animals. For that, the researcher should set the subject on ease. Also, they should be cognizant of the specific features of the animals like size, age, genetic background, and temperament.

The routes could affect bioavailability, suitability, and absorption of the administered compounds (Turner, Brabb, Pekow, & M.A.Vasbinder., 2011). Researchers should handle the animals with great care and patience so that experimental findings could reflect the actual behavior of the animals.


Assessing the Health and Welfare of Laboratory Animals. Retrieved from AHWLA:

Donovan, J., Brown, P., Reeves, J., & Reeves, P. (2003). Introduction to Basic Mouse Handling Techniques. Current protocols in human genetics, 225-226.

Machholz, E., Mulder, G., Ruiz, C., Corning, B. F., & Pritchett-Corning., K. R. (2012). Manual Restraint and Common Compound Administration Routes in Mice and. Journal of visualized experiments(67).

Rabbit – Sexing, Handling, and Restraint. (2017).

Scobie-Trumper, P. (1987). Animal handling and manipulations. In Laboratory Animals: An Introduction for New Experimenters, 153-170.

Sharp, P., & Villano., J. S. (2012). The Laboratory Rat. CRC Press.

Species-Specific Information: Techniques for Handling, Sexing, Injection and Blood Collection. (n.d.). In AN INTRODUCTION TO THE PHYSICS OF PARTICLE ACCELERATORS. World Scientific Publishing Co. Pvt. Ltd.

Suckow, M. A., Danneman, P., & Brayton., C. (2000). In The Laboratory Mouse. CRC Press.

Thomas, B., & Tilla, W. (2004). Handling and Restraint. Switzerland: Elsevier.

Turner, P., Brabb, T., Pekow, C., & M.A.Vasbinder. (2011). Administration of substances to laboratory animals: routes of administration and factors. JAALAS, 600-613.

White, W., & Field., K. (n.d.). Anesthesia and Surgery of Laboratory Animals. In The Veterinary Clinics of North America, Small Animal Practice, Exotic Pet Medicine (pp. 989-1017). Philadelphia.

Wixon, S., & Smiler., K. (1997). Anesthesia and analgesia in rodents. In Anesthesia and Analgesia in Laboratory Animals (pp. 156-200). San Diego. Calif: Academic Press.