Euthanasia of laboratory animals must be performed by trained personnel using appropriate techniques, equipment, and reagents in order to effect a death that is humane and satisfies research requirements. Acceptable methods of euthanasia are painless or minimize distress, and are quick and easy to perform, safe for those performing the procedure, and efficient and economic. They are aesthetically acceptable and are done in the absence of other animals. In addition, these methods do not result in gross histological or histochemical changes that would adversely affect research results. This unit offers protocols for euthanasia employing carbon dioxide asphyxiation (see Basic Protocol 1), pentobarbital overdose (see Basic Protocol 2), exsanguination, and cervical dislocation for the mouse, rat, hamster, and rabbit.

This unit describes the techniques for the following routes of injection for mice, rats, hamsters and rabbits: intramuscular, intradermal, subcutaneous, intravenous, intraperitoneal, footpad, and intrathymic. Guidelines are also given regarding injection volumes and temperatures, and the use of proper restraints.

This unit describes the techniques for the following routes of injection for mice, rats, hamsters and rabbits: intramuscular, intradermal, subcutaneous, intravenous, intraperitoneal, footpad, and intrathymic. Guidelines are also given regarding injection volumes and temperatures, and the use of proper restraints.

A continuous femoral nerve block (cFNB) involves the percutaneous insertion of a catheter adjacent to the femoral nerve, followed by a local anesthetic infusion, improving analgesia following total knee arthroplasty (TKA). Portable infusion pumps allow infusion continuation following hospital discharge, raising the possibility of decreasing hospitalization duration. We therefore used a multicenter, randomized, triple-masked, placebo-controlled study design to test the primary hypothesis that a four-day ambulatory cFNB decreases the time until each of three predefined readiness-for-discharge criteria (adequate analgesia, independence from intravenous opioids, and ambulation ≥ 30 meters) are met following TKA compared with an overnight inpatient-only cFNB. Preoperatively, all patients received a cFNB with perineural ropivacaine 0.2% from surgery until the following morning, at which time they were randomized to either continue perineural ropivacaine (n=39) or switch to normal saline (n=38). Patients were discharged with their cFNB and portable infusion pump as early as postoperative day three. Patients given four days of perineural ropivacaine attained all three criteria in a median (25 th -75 th percentiles) of 47 (29-69) hours, compared with 62 (45-79) hours for those of the control group (Estimated ratio=0.80, 95% confidence interval: 0.66-1.00; p=0.028). Compared with controls, patients randomized to ropivacaine met the discharge criterion for analgesia in 20 (0-38) vs. 38 (15-64) hours (p=0.009), and intravenous opioid independence in 21 (0-37) vs. 33 (11-50) hours (p=0.061). We conclude that a four-day ambulatory cFNB decreases the time to reach three important discharge criteria by an estimated 20% following TKA compared with an overnight cFNB, primarily by improving analgesia.

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Blood Collection Blood is most frequently sampled for evaluation of serum antibodies or analysis of surface markers on peripheral blood cells. This unit describes blood collection methodology for small rodents and rabbits. Blood collection is the most common interventional procedure conducted with laboratory animals and is an essential requirement for many studies. Selection of an appropriate method will depend on the species, amount of blood needed, frequency of sampling, and whether the animal's survival is required (see Commentary). The protocols offered in this unit describe collection of blood from the orbital sinus or plexus of the mouse, rat, or hamster. With appropriate techniques, small amounts of blood can be obtained with little ill effect on the animal. Collection from the auricular vein or artery of the rabbit is also relatively unstressful to the animal. Bleeding procedures that should be performed on the anesthetized animal include collection from the mouse axillary plexus, cardiac puncture of the mouse, rat, hamster, or rabbit, and collection from the hamster abdominal aorta or vena cava. All procedures benefit from training with an experienced animal technician. NOTE: Wear disposable gloves when handling animals in the following protocols. BASIC PROTOCOL 1 BLOOD COLLECTION FROM ORBITAL SINUS OR PLEXUS OF MOUSE AND RAT Materials Sterile saline or phosphate-buffered saline (PBS; APPENDIX 2) Microhematocrit tube Gauze sponge or swab Additional reagents and equipment for handling and restraint (UNIT 1.3) and anesthesia (UNIT 1.4) 1. Manually restrain and anesthetize the animal. 2. Introduce the end of the microhematocrit tube at the medial canthus of the orbit as shown in Figure 1.7.1. 3. Slowly, and with axial rotation, advance the tip of the microhematocrit tube gently towards the rear of the socket until blood flows into the tube.

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Anesthetic agents are used in laboratory animals to prevent pain or distress due to an experimental procedure or for restraint to facilitate a technically difficult procedure. This unit provides three basic protocols: injectable anesthesia for mouse, rat, and hamster; inhalant anesthesia using methoxyflurane for mouse, rat, and hamster; and injectable anesthesia using ketamine/xylazine for rabbit. An Alternate Protocol describes sedation using butorphanol/acetylpromazine in the rabbit. The Commentary further describes and compares these methods of anesthesia for various applications.