Anesthesia in Experimental Animals

General anesthesia, a condition in which the animal is unconscious and completely insensible to pain, is required for surgery and other painful procedures. In addition, because surgical and other invasive procedures are expected to cause pain or discomfort beyond the duration of general anesthesia, post-procedural analgesia (pain relief) must be provided until no longer needed by the animal. General anesthesia may also be used as a method of restraint for nonsurgical procedures that require immobilization, cause distress for the animal or expose human handlers to potential hazards. Inhalation (gas) anesthesia using isoflurane is recommended for rodent surgeries. Survival rates under gas anesthesia are generally higher and recovery faster compared to injectable anesthesia.

Analgesia in Experimental Rodents

Investigators are responsible for the assessment and management of all types of pain in their research animals and must include a pain management plan in their research protocols. Please contact an ARP veterinarian if you have questions or need additional information than what is provided in this website.

Controlled Drugs

A number of drugs used for anesthesia, analgesia, and euthanasia are regulated by the DEA (Drug Enforcement Administration) because of the potential for abuse. Investigators using these drugs are responsible for complying with applicable regulations including storage in a substantial locked cabinet or safe and maintenance of written records accounting for quantities received and dispensed. Investigators who will be using controlled drugs must apply to the DEA for the appropriate license.

Anesthetic Monitoring

Verify that the animal is not able to feel pain by firmly pinching the rear paw at least every 5 minutes. If the animal moves it is too lightly anesthetized for painful procedures. Consider using an alarm set to beep at 5-minute intervals to remind you to check the animal. In addition, monitor respiration, heart rate and mucous membrane color frequently throughout anesthesia. Shallow, infrequent breathing; pale or blue mucous membranes; and decreasing heart rate can be indicators that the animal is too deeply anesthetized. Note: anesthetized animals that appear to be gasping with an open mouth are often dying. A pulse oximeter may be used to monitor the heart rate and blood oxygen levels. The SpO2 should be at least 95%. Values less than 95% indicate mild hypoxia. SpO2 values of 90% or less indicate significant hypoxia and require immediate O2 supplementation and reduction in isoflurane flow.

Body Temperature

Body temperature drops significantly within a few minutes after induction of anesthesia in small animals, especially rodents. Below normal body temperature (hypothermia) significantly affects physiological function, prolongs anesthetic recovery and is a common cause of death in anesthetized mice. Rectal probes are available for monitoring body temperature in anesthetized animals. During surgery, circulating water blankets or far-infrared heating pads specifically made for small animals are commonly used. Post-operatively, recovery cages may be placed onto these or other devices that will provide gentle heat. With any device, care must be taken to prevent overheating. Electric heating pads and heat lamps are not acceptable as they can burn the skin and extremities. Anesthetized animals must be provided with supplemental heat at least until they are recovered from anesthesia. In some cases, after prolonged or invasive surgical procedures, supplemental heat should be provided to the cage for up to several days.

Prevention of Fluid Loss

Animals may experience extensive fluid loss during and after surgery due to evaporation from exposed body tissues and cavities, bleeding and/or lack of fluid intake perioperatively. To reduce intra-operative fluid loss the surgeon should control blood loss by clamping and/or ligating bleeding vessels and may gently irrigate exposed tissues with warmed sterile saline. In addition, it is recommended that warm, sterile fluids (e.g., 0.9% NaCl or LRS) be provided to animals undergoing surgery. In rabbits, fluids may be given intraveneously during and after surgery. In rodents, fluids are usually injected subcutaneously (Mouse: 40 ml/kg total, not > 1.0 ml per site; Rat: 25 ml/kg total, not > 5.0 ml per site), divided between 2-3 sites prior to anesthetic recovery. An animal’s water and food intake and body weight should be monitored as part of post-operative care and additional fluids administered as needed. Please consult with an ARP veterinarian for further information.

Inhalant Anesthesia

Inhalant anesthetics allow precise control of the depth and duration of anesthesia in research animals and can be used in almost any species. Isoflurane is the most commonly used inhalant anesthetic at PSU and provides rapid induction and recovery from anesthesia. The depth of anesthesia can be easily and quickly altered. Virtually no metabolism occurs in the body because isoflurane is primarily eliminated in expired air. Liver microsomal enzymes are minimally affected which results in little interference with drug metabolism or toxicology studies. The use of isoflurane requires an anesthetic machine fitted with a precision vaporizer to deliver controlled amounts of anesthetic and oxygen. Anesthetic can be delivered to the animal via an anesthetic chamber, facemask or endotracheal tube. Typically, anesthesia is induced with the animal in an anesthetic chamber. After consciousness is lost, the animal is moved to a nosecone for maintenance of anesthesia during the procedure. Isoflurane anesthetic systems are available in the University Park campus animal facilities. Please contact the ARP for information on training and use of this equipment.

Injectable Anesthesia

Injectable anesthetics may not reliably produce a surgical plane of anesthesia, particularly when injected intraperitoneally in rodents. The duration of injectable general anesthesia is often variable and recovery slow. Small animals, especially mice, breathing room air after administration of injectable anesthetic drugs become hypoxic (i.e., decreased oxygen levels in the blood) within a few minutes after losing consciousness. Hypoxia may lead to complications including delayed anesthetic recovery and/or death. It is recommended that 100% oxygen be supplied via nosecone to all rodents anesthetized with injectable drugs. Drugs are available that can shorten ('reverse') anesthesia time for certain injectable anesthetics, but these should not be given until at least 30 minutes after anesthetic administration. Please consult with an ARP veterinarian for additional information regarded injectable anesthetic use.

The amount of drug administered to each animal is based on its body weight. Each animal should be weighed on the day of anesthesia. Many other factors (e.g., strain, sex, age, and stress levels) also influence how an anesthetic drug and dosage affects an individual animal. Sterile technique must be used in the preparation and administration of injectable drugs. All needles, syringes and containers used to deliver or store drugs must be sterile. The use of nonsterile equipment can result in animal infections and/or illness.

Scroll down for dosage tables for several anesthetic drugs in a variety of species. The following link provides dilution instructions for common anesthetic drugs used in research rodents:

Anesthetic drug mixtures for rodents

Preparation and Use of Injectable Anesthetic Drugs

Most injectable anesthetic drugs are purchased as sterile, ready-to-use products from veterinary distributors. More than one drug may be combined to provide better anesthetic results (see below). The amount of drug administered to an animal is based on its body weight. Each animal needs to be weighed on the day of anesthesia. Purchased drugs are typically supplied at a concentration that requires dilution with sterile water for ease of use in rodents. Sterile technique must be used in the preparation and administration of all injectable drugs. All needles, syringes and containers used to prepare, deliver or store drugs must be sterile.

Injectable Anesthetic Dosages

Drug(s)	Mouse	Rat	Rabbit	Hamster
Ketamine (K): Xylazine (X)	80-120 mg/kg (K):10-16 mg/kg (X) IP (20-40 min. of anesthesia)	80-100 mg/kg (K): 5-10mg/kg (X) IP (20-50 min. of anesthesia)	22-50 mg/kg (K):2.5-10 mg/kg (X) IM (25-40 minutes of anesthesia)	50-200 mg/kg (K) + 5-10 mg/kg (X) IP
Ketamine(K): Dexmedetomidine (D)	75 mg/kg (K): 0.5 mg/kg (D) IP	75 mg/kg (K): 0.5 mg/kg (D) IP		100 mg/kg (K) + 0.125 mg/kg (D) IP
Ketamine (K): Xylazine (X): Acepromazine (A)		50 mg/kg (K): 5 mg/kg (X): 1 mg/kg (A) IP (30-45 minutes of anesthesia)	35 mg/kg (K): 5 mg/kg (X): 0.75 mg/kg (A) IM (45-75 min. of anesthesia)
Ketamine (K): Diazepam (D) - short duration; not surgical anesthesia			5 mg/kg (K): 0.25 mg/kg (D) IV. Mix equal volumes of ketamine (100 mg/ml) and diazepam ( 5 mg/ml). Dosage rate: 1 ml per 20 lbs body weight.	70 mg/kg (K) + 2 mg/kg (D) IP (Immobilization; not surgical anesthesia)
Inactin (thiobutabarbital, EMTU)		80-100 mg/kg IP (60-240 min. of anesthesia)
Pentobarbital -variable anesthetic depth; poor analgesia	30-50 mg/kg IP (20-40 min. of anesthesia)	40-50 mg/kg IP (20-60 min. of anesthesia)		Not recommended for hamsters due to high mortality
Tribromoethanol (Avertin)	240 mg/kg IP (15-45 min. of anesthesia)
Atipamezole (for reversal of dexmedetomidine & xylazine)	1.0 mg/kg IP or SQ	1.0 mg/kg IP or SQ	0.1-1.0 mg/kg IM, IP, SQ or IV (dose required depends on dose of xylazine administered)	0.1-1 mg/kg SQ or IP
Glycopyrrolate (preanesthetic)			0.1 mg/kg IM, SQ (decreases respiratory secretions, prevents bradycardia)

For information on other anesthetic drugs and dosages please consult an ARP veterinarian.

The information and drug dosages presented in this website are intended as a resource for Pennsylvania State University research investigators. No guarantee of drug efficacy or safety is made nor must information obtained from this site be substituted for professional veterinary advice.