Animal Care Services

These guidelines have been developed to introduce investigative staff to procedures recommended for general injection techniques when working with small laboratory animals. This document is intended to supplement hands-on instruction by an experienced member of your laboratory or a member of the Animal Care Services (ACS) staff. ACS clinical staff is available to provide hands-on instruction. You can contact the ACS Training Coordinator at 392-9536 to register for the next available session.

There are a variety of other techniques in addition to those described in this document that are suitable alternatives. You can contact the ACS to discuss their suitability and describe them in your animal care and use protocol.

ACS offers free training for the administration of injections in a variety of laboratory species. In addition, ACS can provide experimental or therapeutic injections for your animals on a fee-for-services basis to investigators for a nominal fee. Contact the ACS Training Coordinator at 392-9536 for more details on blood collection training, or 846-0984 to arrange fee-for-service procedures to be performed by ACS personnel.

Parenteral Injections
The ability to administer materials by injection is essential for most experimental studies employing laboratory animals. Anesthetics and analgesics, therapeutic agents, and test compounds must frequently be administered to animal subjects by injection. There are five commonly used routes of parenteral administration: subcutaneous (SC/SQ), intraperitoneal (IP), intravenous (IV), intradermal (ID), and intramuscular (IM). Not all techniques are appropriate for each species. For example; IM injections are avoided in rodents because the amount of material that can be injected into the rodent's limited muscle mass is so small that the technique is not practical; IP injections are not administered to rabbits as other techniques are more suitable.
It is essential that the appropriate parenteral site be selected. Systemic absorption and distribution differ considerably between sites. Dosage and volume of material administered must be carefully considered relative to the type of agent, site of injection, and species used. The size of syringe and needle must also be considered. In order to assure the delivery of an accurate volume of injected material, the volume of the syringe should, in general, not exceed the volume of material to be administered by 10 fold. The length of the selected needle should be long enough that sufficient tissue penetration is achieved but not be so long that it becomes unmanageable or is likely to be inserted too far. You do not need to advance the needle to the hub; simply as far as is necessary to get the tip of the needle to the desired delivery site. The volume and viscosity to the material to be injected directly impacts the selection of a particular delivery system. The needle's size should be as small (highest gauge) as possible to limit tissue trauma but should also be large enough that the injection can be made relatively rapidly, without applying excessive pressure to the syringe plunger. Syringe and needles should be of the locking type in order to prevent accidental dislodgement, which may result in back spray or the need for a second injection. Proper disposal of used needles and syringes is essential. Needles should never be recapped, as the risk of accidental injection is highest during recapping, and they should always be disposed of in a designated sharps container.
Injection volumes provided in this document are general recommendations. Under some circumstances it may be inappropriate to inject the recommended volume. For example, volumes should be reduced when the agent is irritating, hypotonic, or hypertonic. Volumes may be increased when giving isotonic fluids for rehydration and fluid maintenance. For example, rabbits, cats, and dog can receive 10 – 20 mL/kg (LRS or 0.9%NaCl) subcutaneously for anorexia or surgery.

“Good practice” administration routes and volumes for mice, rats, and rabbits are indicated in Table 1 .

Recommended needle sizes and injection sites for various species are indicated in Table 2.

PROCEDURES

Mice
Subcutaneous injection
SC injections can be administered easily to mice. The needle is inserted between the folds of skin into the base of the triangle that is formed when traction is applied to the skin overlying the animal's scruff. The syringe's plunger should be retracted to verify that a vacuum is created and no blood is aspirated. Subsequently, the plunger is depressed to administer the material. Several sites over the animal's back should be used if larger volumes must be administered. In general, needles should be the smallest diameter possible for the solution to be administered.
Intraperitoneal injection
The administration of material into the peritoneal cavity is frequently performed in mice. The aim of this technique is to administer material into the space surrounding the abdominal organs, avoiding injection directly into any organ. Restrain mice with their abdomen exposed (uppermost) and their head pointed downward, this causes the freely moveable abdominal organs to move towards the animal's diaphragm making accidental puncture of organs less likely. The needle is inserted into the abdominal cavity in the animal’s lower right quadrant to avoid the cecum and urinary bladder. The needle should be directed towards the animal's head at an angle of 15 - 20 degrees and inserted approximately 5 mm. Aspiration should be attempted to ensure that an abdominal organ (such as the bladder or intestines) has not been penetrated. If bladder content or intestinal material is aspirated, the syringe must be removed and discarded. Never inject GI tract contents or urine into the peritoneal cavity, as a bacterial or chemical peritonitis will likely result.
Intravenous injection
The veins on the lateral aspects of the mouse's tail are excellent sites for IV administration. The principal function of these veins is for thermoregulation. They will dilate when the mouse's body temperature rises in order to dissipate heat. Application of heat to the whole animal or locally to the tail can be used to cause vasodilatation making vascular access easier. Dilate the tail vessels by placing the tail in warm water (37oC), never exceeding 40 - 44oC range, or under a heat lamp (25-30 cm away using a 60W bulb). The animal’s body temperature should never exceed 104oF (40oC) for over 5 minutes. Animals must be constantly monitored for signs of distress for these heat exposure techniques. The mouse should be restrained so that its tail is accessible. A 25-28G needle is used. The vein is located, the needle inserted by directing the needle into the vein with its bevel facing upward at an angle of approximately 20 degrees. The needle is inserted slowly. Visualize the needle as it enters the vein. Once the vein's wall has been penetrated, decrease the needle’s angle and the needle should be directed cranially approximately 2 mm. Blood should be aspirated into the needle's hub before making an injection. During material administration the vein should blanch and no material or swelling should be detectable at the injection site. Material should be administered slowly to avoid vascular overload or rupture of the vein from excess pressure. Pressure should be applied over the injection site by gently holding a piece of gauze over the injection site for approximately 30 seconds to prevent hematoma formation. Preferably the needle should be inserted into the vein midway down the tail, permitting additional attempts for venipuncture proximally if the initial attempt is unsuccessful.

Rats
Subcutaneous
SC injections are performed in rats using the same technique as was described for mice.
Intravenous
Tail vein IV injection technique for the rat is similar to the mouse. However, the vessels are more difficult to visualize, especially in adult rats. The skin overlying the vessels in adults becomes quite thick, making vascular access more difficult. For this reason the preferred site for vascular access is near the tail base. The sublingual or the penile veins are also acceptable routes. Material should be administered slowly to avoid vascular overload.
Intraperitoneal
The technique for IP injections in rats is virtually identical to mice. Rats should be restrained with their abdomen exposed and their head held downward. The injection site and technique are as described for mice.

Rabbits
Subcutaneous injections
SC injections are easily performed in rabbits because of the laxity of their skin and the large area into which material can be administered. The technique is the same as described for mice, however injections should not be administered over the neck as this is the site from which the animal is picked up. Volumes should not exceed 5 mL per site unless isotonic fluids are being administered.
Intramuscular injections
The recommended sites and technique for IM injection in the rabbit are the cranial thigh muscles, or the lumbar muscles to the side of the spine and just cranial to the pelvis. Caudal thigh injections are more likely to damage the sciatic nerve and should be avoided. Rabbits large muscle mass requires a relatively long needle (1/2” - 1”) to adequately introduce the material deep within the muscle tissue, a distance of approximately 7 - 10 mm in an adult rabbit.
Intravenous injections
IV injections are straightforward in rabbits because of the ease of vascular access. The marginal ear veins located on the lateral aspect of the rabbit's ears are readily visible. The veins can be made more prominent by occluding the vessel at the base of the ear by gently holding off with your fingers. The vein is swabbed with an alcohol soaked cotton pad. The needle is inserted as described for injecting rodent tail veins. Remember to aspirate to verify placement of the needle within the vein. Remove your occluding fingers prior to injection. A butterfly needle or a catheter may be inserted if larger volumes are to be administered or repetitive injections will be given. Pressure should be applied over the injection site by gently holding a piece of gauze over the injection site for approximately a minute to prevent hematoma formation.
Intradermal injections
ID injections may be used to immunize rabbits. The technique is as described for guinea pigs with the following changes. The neck and anterior thoracic region should be avoided for injection as rabbits are handled by grasping this region.

Hamsters
Subcutaneous
SC injections are performed in hamsters using the same technique as was described for mice. The hamster has considerably more loose skin overlying the injecting site permitting a proportionately larger volume of material to be administered, up to 3 mL.
Intravenous
IV injections are difficult to perform in the hamster because of the lack of easily accessible veins. The ACS staff should be contacted for additional information.
Intraperitoneal
The technique for IP injections in hamsters is virtually identical to those described for mice and rats. Hamsters should be restrained with their abdomen exposed and their head held downward. The injection site, method and needle size is as described for mice. Because of their larger size up to 3.0 mL of material can be administered to an adult hamster.

Guinea Pigs
Subcutaneous
SC injections are performed in guinea pigs using the same technique as was described for mice with the following differences. The volume of material administered can be increased to approximately 5 mL per site in an adult guinea pig.
Intravenous
IV injections are very difficult to perform in guinea pigs because of the lack of easily accessible veins. The veins at the base of the tongue or penile vessels (males) may be used. Consult the ACS staff for additional information.
Intraperitoneal
The technique for IP injections in guinea pigs is virtually identical to mice. Guinea pigs should be restrained with their abdomen exposed and their head held downward. The injection site, method and needle size is as described for mice. Because of their larger size up to 5.0 mL of material can be administered to an adult guinea pig.
Intradermal injections
ID injections may be used to immunize guinea pigs. In contrast to SC injections where material is deposited into the space between the skin and body wall, ID injections deposit material within the layers of the skin. Therefore, the volume of material that can be administered is very small (0.1 mL per site; 0.05 mL recommended). The fur should be clipped so that the injection site can be clearly observed. A 0.5 inch 25 or higher gauge needle and a 1 cc syringe are recommended. The area to be injected is swabbed with an alcohol soaked cotton pad. The needle is inserted bevel up into the skin at approximately a 15 - 20 degrees angle. The needle is advanced approximately 1 mm. The material is injected slowly creating a small bleb that typically takes several minutes to resolve. Immediate dissolution of the bleb indicates that the material has been injected subcutaneously rather than intradermally. ID injections should be made over the dorsal thoracic and lumbar region. Multiple sites (up to 10) can be used.

Table 1. Administration volumes considered good practice (and possible maximal dose volumes)a Journal of Applied Toxicology

Species route and volumes (mL kg-1)
                  Oral           s.c.          i.p.          _ i.m.       __ i.v. (bolus)  i.v. (slow)
Mouse        10 (50)       10 (40)     20 (80)      0.05b (0.1)b     5           (25)
Rat            10 (40)         5 (10)     10 (20)      0.1b (0.2)b       5           (20)
Rabbit        10 (15)         1 (2)        5 (20)      0.25 (0.5)         2           (10)

a For non-aqueous injectables, consideration must be given to time of absorption before re-dosing. No more than two intramuscular sites should be used per day. Subcutaneous sites should be limited to two or three sites per day. The subcutaneous site does not include Freund’s adjuvant administration.
b Values in milliliters per site.

Table 2. Needle Sizes and Recommended Injection Sites; Adapted from: Formulary for Laboratory Animals, 3rd Ed., Hawk, Leary, Morris, 2005

Species	Injection Site
	SC	IM	IP	IV
Feline	Scruff, back, 21 – 23G	Quadriceps, 23G	21 – 23G	Cephalic vein, 21 – 25G
Canine	Scruff, back, 21 – 23G	Quadriceps, 21 - 23G	21 – 23G	Cephalic vein, 21 – 25G
Ferret	Scruff, 21 – 23G	Quadriceps, 23 - 25G	21 – 23G	Cephalic vein, 21 – 25G
Guinea pig	Scruff, back, 23 – 25	Quadriceps, 25G	23 – 25G	Ear vein, Saphenous vein, 25 – 27G
Hamster	Scruff, 25G	Quadriceps, 25G	23 – 25G	Femoral or jugular vein, 25 – 27G
Mouse	Scruff, back, 25G	Quadriceps, 27G	25 – 27G	Lateral tail vein, 26 – 28G
Primate (small)	Scruff, back, 23 – 25G	Quadriceps, 23 - 25G	21 – 25G	Lateral tail vein, 21 – 25G
Primate (large)	Scruff, 21 – 25G	Quadriceps, triceps, 23 - 25G	21 – 23G	Cephalic vein, recurrent tarsal vein, jugular vein 21 – 25G
Rabbit	Scruff, flank, 21 – 25G	Quadriceps, lumbar muscles, 25G	21 – 23G	Marginal ear vein, 23 – 25G
Rat	Scruff, 25G	Quadriceps, 25G	23 – 25G	Lateral tail vein, 21 – 23G
Bird	Pectoral, interscapular or inguinal fold 1 – 3% BW bid or tid, <21 G	Pectoral/per site 0.2 ml/<100g BW 0.2-0.5ml/100-500g BW 0.5-1.0ml/>500g BW <25G	Not applicable	Cutaneous ulnar vein, <25G, short bevel

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Credits
American Association for Laboratory Animal Science

Harkness JE and Wagner JE, The Biology and Medicine of Rabbits & Rodents.

A Good Practice Guide to the Administration of Substances and Removal of Blood,
Including Routes and Volumes, Diehl, et.al., Journal of Applied Toxicology, 21, 15–23 (2001)

Formulary for Laboratory Animals, 3rd Ed., Hawk, Leary, Morris, 2005, Blackwell Publishing

Ferrets, Rabbits, and Rodents Clinical Medicine and Surgery, 2nd Edition, K. E. Quesenberry and I. W. Carpenter, Saunders, (Elsevier), St. Louis, Missouri

University of Washington Training Series

Hawk, C. and Leary, S. (1995), Formulary for Laboratory Animals, Iowa State University Press, Ames.