Injection of Laboratory Animals - Thiel College

Thiel College Institutional Animal Care and Utilization Committee

Injection of Laboratory Animals

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 inTable 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

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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

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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.

Table 1. Administration volumes considered good practice (and possible maximal dose volumes) a Journal of Applied Toxicology; Species route and volumes (mL kg-1)

Mouse Rat Rabbit

Oral 10 (50) 10 (40) 10 (15)

s.c. 10 (40) 5 (10) 1 (2)

i.p. 20 (80) 10 (20) 5 (20)

i.m. 0.5b (0.1)b 0.1b (0.2)b 0.25 (0.5)

i.v. (bolus) 5 5 2

i.v. (slow) (25) (20) (10)

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

Species

Feline

Canine

Ferret

Guinea pig

Hamster

Mouse

Primate (small) Primate (large)

Injection Site SC Scruff, back, 21 ? 23G Scruff, back, 21 ? 23G Scruff, 21 ? 23G Scruff, back, 23 ? 25 Scruff, 25G Scruff, back, 25G Scruff, back, 23 ? 25G Scruff, 21 ? 25G

Rabbit Rat

Scruff, flank, 21 ? 25G Scruff, 25G

IM Quadriceps, 23G

IP 21 ? 23G

Quadriceps, 21 - 23G Quadriceps, 23 - 25G Quadriceps, 25G

21 ? 23G 21 ? 23G 23 ? 25G

Quadriceps, 25G

23 ? 25G

Quadriceps, 27G

25 ? 27G

Quadriceps, 23 - 25G Quadriceps, triceps, 23 - 25G

21 ? 25G 21 ? 23G

Quadriceps, lumbar muscles, 25G Quadriceps, 25G

21 ? 23G 23 ? 25G

IV Cephalic vein, 21 ? 25G Cephalic vein, 21 ? 25G Cephalic vein, 21 ? 25G Ear vein, Saphenous vein, 25 ? 27G Femoral or jugular vein, 25 ? 27G Lateral tail vein, 26 ? 28G Lateral tail vein, 21 ? 25G Cephalic vein, recurrent tarsal vein, jugular vein 21 ? 25G Marginal ear vein, 23 ? 25G Lateral tail vein, 21 ? 23G

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Bird

Pectoral, interscapular Pectoral/per site 0.2 Not

Cutaneous ulnar vein,

or inguinal fold 1 ? 3% ml/ ................
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