Bloodcollectionandadministrationoffluidsanddrugs(mouse)2

Intraperitoneal:

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.

Subcutaneous:

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.

Intramuscular:

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.

Blood Collection:

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.

Site Preparation:

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.

Tail vein venipuncture in mice:

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.

Toe clipping or tail clipping to obtain blood samples: Clipping toes is an unacceptable method of blood collection. Tail clipping is not a preferable method for blood collection.

Cardiac puncture:

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.

Orbital sinus venipuncture in mice:

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.

Axillary bleed:

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.