Blood collection and administration of fluids and drugs (mouse)

When drugs, vaccines, injectable anesthetics or other agents are to be administered, one of several different routes may be selected. The route is governed by the nature of the agent being administered, the animal, and the purpose of the administration, among other factors.

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picture ). Flexible or plastic tubes may be bitten or chewed and are not recommended for rodents. Feeding needles are inserted through the mouth into the stomach or lower esophagus. Placing the needle next to the mouse so that the end of the needle is adjacent to the last rib and marking the position on the needle that is adjacent to the tip of the nose will determine the distance the feeding needle should be advanced into the oral cavity to insure administration of the compound into the stomach (picture ). Care must be taken that the tube or needle does not enter the trachea or puncture the esophagus or stomach, therefore knowledge of the oropharyngeal anatomy is necessary. In most cases, introduction of the tube toward the rear of the mouth will induce swallowing and the tube will readily enter the esophagus. A violent reaction (coughing, gasping) usually follows accidental introduction of the tube into the larynx or trachea. With the mouse restrained in one hand the feeding needle is introduced in the space between the incisors and the beginning of the molars (diastema)(picture ). If introduced from the left diastema the needle should be directed caudally toward the right ramus of the mandible. As the needle approaches the pharynx the mouse will usually swallow allowing introduction into the esophagus. Using the feeding needle to gently extend the neck facilitates introduction into the stomach (future video). With the stomach tube fitted to a syringe or aspirator, materials may be administered or withdrawn as required.

1. Intravenous (iv) - - directly into the vascular system through a vein
2. Intra-arterial (ia) - - directly into the vascular system through an artery
3. Intraperitoneal (ip) - - into the abdominal cavity
4. Subcutaneous (sc) - - under the skin
5. Intramuscular (im) - - into a muscle
6. Intradermal (id) - - between layers of skin

General Considerations: Parenteral routes of administration involve injections into various compartments of the body. Sites used for collection of blood from veins may also be used for intravenous administration. Intraperitoneal administration is one of the most frequently-used parenteral routes in rodents. Other locations are the musculature and subcutis. Materials given intramuscularly must be in very small volumes and is generally not recommended unless necessary. Absorption by this route is more rapid than from subcutaneous administration. Regardless of the route used, it is essential that the subject be securely restrained to prevent unnecessary struggling by the animal and to avoid injury to personnel by dislodged needles.

The investigator should know the physiological and chemical properties of the substance that he/she plans to inject. Considerable tissue damage and discomfort can be caused by irritating vehicles, drugs or solutions when injected into animals. The use of the foot pad as an injection site for antigens with or without adjuvant is discouraged since it is a needless and painful procedure. More suitable sites for antigen injections are subcutaneously in the axilla or lateral thoracic wall, deep in large muscle masses, or into the popliteal lymph node.

Demonstration/instruction sessions can be arranged with the ACU.

Equipment: 27-30 g needle, 1 ml tuberculin syringe, mouse holder, warming lamp.

Volume: The volume injected IV into an adult mouse should not exceed 0.2 ml.

The lateral veins of the tail are the most frequently-used veins. Best results are obtained if the tail is immersed in warm water for 5 to 10 seconds or the mouse warmed for 5 to 15 minutes in the cage with a warming lamp with a 40 to 100 watt bulb. The veins can be seen when the tip of the tail is lifted and rotated slightly in either direction. The tip of the needle can be followed visually as it penetrates the vein. Trial injection verifies proper needle placement. Also, accurate placement can be confirmed when the vessel is visually flushed when the compound is administered. The formation of a bleb at the site indicates improper placement of the needle. A second attempt can be performed by removing the needle and trying a site on the same vessel in a more proximal location on the tail. Practice is essential.

Equipment: Syringe and 23 to 27 g, 1/2 to 1-inch needle, preferably with a short bevel.

Volume: The volume injected IP into an adult mouse should not exceed 2 ml.

The mouse is grasped as previously described , and held in dorsal recumbency in a head-down position. The injection is made in the lateral aspect of the lower left quadrant (picture ). The use of a short bevel needle inserted through the skin and musculature and immediately lifted against the abdominal wall, aids in avoiding puncture of the abdominal viscera. Immobilizing the left leg is also essential in reducing this risk. Rapid injection, especially with a large syringe, may cause discomfort and tissue damage and should be avoided.

Equipment: Syringe and 25 to 27 g, 1/2 to 3/4-inch needle.

Volume: The volume injected SQ into an adult mouse should not exceed 2-3 ml.

This route is frequently used as an alternative to intramuscular injections in the mouse. The site usually chosen is the loose skin between the shoulder blades. Alternatively, the ventral abdomen is commonly used, employing one handed restraint (picture ). The needle is inserted through the skin and advanced 5 to 10 mm through the subcutaneous tissue to prevent leakage from the site.

Equipment : Syringe and 26 to 30 g, 1/2-inch needle.

Volume : The volume injected intramuscular in the adult mouse should not exceed 0.05 ml.

This route is usually not used and is not recommended because of the small muscle mass available and the danger of damaging vital structures. However, when it is used, the back and hind leg muscles are the usual sites selected.

The amount of blood needed and other factors will govern the method and sites of collection. Table I (future table) lists common blood withdrawal sites in laboratory animals and precautions and requirements for these procedures. Descriptions of the various techniques for venipuncture in different species is available in the Animal Care Unit (400 ML; 335-7985) in text and videotape format. Proper insertion of the needle into a vein or other part of the vascular system is normally the most difficult part of the procedure. Certain guidelines can be given, but only practice provides proficiency. Veins may be expected to roll, collapse, or shift, making entrance difficult. A precise, careful introduction of the needle is best and several attempts may be required. Starting at distal sites will allow repeat attempts more proximally. The needle is inserted parallel to the vein and the tip directed into the lumen along the longitudinal axis. When withdrawing blood from a vein, aspiration should be slow so the vessel does not collapse.

The area of injection or incision should be cleaned with alcohol. Some procedures will require sedation or anesthesia; others may be carried out without anesthesia provided suitable restraint is used. In order to better visualize veins dilation can be accomplished by immersing the tail in warm water for 5 to 10 seconds or by warming the animal with a low-wattage light bulb for 5 to 15 minutes prior to venipuncture. This also aids by providing additional light.

Equipment : scalpel blade, 25 to 30 gauge needle

The mouse is restrained (picture1 ; picture2 ) using a mechanical device. The veins may be seen laterally near the base of the tail but good illumination and dilation will normally be required. A small blood sample may be collected by capillary action using a microhematocrit tube inserted into the hub of a small needle previously placed into the tail vein (future picture). This technique normally recovers a few drops of blood, adequate for hemoglobin, microhematocrit and cell counts.

Larger blood samples can be obtained by making a small incision over the vessels 0.5 to 2 cm from the tail base using a scalpel blade. One half to one milliliter of blood can be withdrawn using this method. Anesthesia or sedation should be used.

Equipment: 0.90 to 0.50 mm needle

Cardiac puncture represents an accepted method of blood collection from mice when more than a few drops are required. However, this method also carries considerable risk to the animal and occasionally deaths occur. It is not recommended as a repetitive blood sampling procedure. Animals must be anesthetized and restrained in dorsal recumbency. The needle is inserted under the xyphoid cartilage slightly to the left of midline (picture ). The needle is advanced at a 20 to 30 degree angel from the horizontal axis to the sternum to enter the heart. Aspirate lightly while advancing. Blood should be withdrawn slowly, and the amount must be limited (up to 1 ml) unless euthanasia is planned.

Equipment: Capillary tubes

Blood collection from the orbital sinus of mice is frequently used. One quarter ml can be repeatedly collected from mice at weekly intervals from alternate sides. Bleeding requires that the tube be directed into the orbital sinus (picture , picture ) which surrounds the globe. In the mouse, the tube is inserted into the medial canthus of the eye and directed caudally and slightly dorsally. Knowledge of the location of the venous structures of the orbit of the mouse aids in establishing a successful peri-orbital bleeding technique. Pressure should be applied after blood collection to prevent hematomas. Anesthesia is required for all peri-orbital bleeding procedures.

Equipment: scalpel blade 3 to 5 cc syringe

Blood can be collected from the axillary region in a terminal exsanguination. Exsanguination of the mouse can be achieved by incising the right or left axillary region of an anesthetized mouse in dorsal recumbency. One to two ml of blood can be harvested in this manner.