Anesthesia and Analgesia
This module will provide a brief introduction to analgesia and anesthesia in the mouse.
Your veterinarian should always be consulted for advice on selection and administration of analgesia or anesthesia.
The use of analgesics and/or anesthetics must be described in detail in your approved Animal Study Proposal.
The most commonly used analgesics for mice at NHGRI are shown below.
For the management of pain in association with surgery or other procedures causing discomfort, injectable analgesics may be used.
Refer to the table shown below and your veterinarian for appropriate dosages and frequency of administration.
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Click the image below to see a TABLE OF COMMONLY USED ANESTHETICS AND ANALGESICS FOR MICE, including:
tribromoethanol (Avertin), bupivacaine (Marcaine), buprenorphine (Buprenex), butorphanol (Torbutrol), ethyl chloride, halothane (Fluothane), isoflurane (Aerrane), ketamine/xylazine (Rompun and Ketaset), ketoprofen (Ketofen), pentobarbital (Nembutal), proparacaine (Alcaine), ropivacaine (Naropin) and tetracaine.
It is important to weigh the mice prior to dosing with injectable anesthetics to avoid over or under dosing the animals.
As an adjunct to, or in lieu of injectable analgesics, topical anesthetics may also be used.
These long-acting agents are painted or dropped into the surgical wound before the skin is closed.
To facilitate retro-orbital sinus blood collection, an ophthalmic anesthetic is used as a topical
A single drop of the solution is placed on the eye.
Be careful not to touch the tip of
the applicator to any part of the mouse. This will cause contamination of the anesthetic.
After approximately 5 - 10 seconds, gently blot away the excess anesthetic with clean gauze, being
careful not to scratch the cornea.
Proceed with blood collection.
Pay special attention to the
storage requirements for the ophthalmic anesthetics, as some require refrigeration.
For mice undergoing tail snips for genotyping, a topical hypothermic agent is sprayed on the
This topical analgesic is required for mice greater than 10 days of age.
The tail is sprayed continuously for 3 - 7 seconds from a distance of 3 - 9 inches.
The tail will blanch showing a proper amount of the agent has been applied.
The distal 5 mm or less of the tail is then excised, as described in the genotyping section of this program.
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At NHGRI, the most commonly used injectable anesthetics for mice are tribromoethanol, pentobarbital and a ketamine/xylazine mixture.
Since tribromoethanol, known by its European brand name Avertin, is not commercially available in the U.S., it is prepared in the laboratory.
This requires strict aseptic technique and special storage considerations.
The solution should be filtered through a 0.22 micron filter.
This will remove debris, most bacteria, and some viruses.
The concentrated stock solution should be stored at -20° Centigrade or colder.
The working solution should be stored at 4° Centigrade in a sterile amber or aluminum foil-wrapped bottle.
When diluting the stock solution of Avertin to the working solution, it is important to use a buffered diluent.
Decomposition of the anesthetic can result from improper storage.
The pH should be greater than 5.
Avertin is administered as an IP injection.
The most common dose is 0.24 to 0.4 micrograms per gram body weight.
Refer to NHGRI Guideline 03.2 “The Use of Tribromoethanol in Mice” for additional information on storage and handling requirements.
Pentobarbital, buprenorphine and butorphanol are controlled substances and can only be purchased through your Controlled Substances Officer.
Ketamine is also considered a controlled substance and must be purchased through your Controlled Substances Officer.
Detailed records are required for the use of controlled substances.
Failure to maintain proper records can result in the loss of your privilege to use these agents.
Please note that dosages may vary according to the strain, age, sex, etc. of the animals.
At NHGRI, the most commonly used inhalant anesthesia is isoflurane.
There are several types of gas anesthesia machines available for use.
Isoflurane is administered in 100% 02.
Induction concentrations of isoflurane are 3 – 4%.
Maintenance concentrations are 1.25 – 1.75%.
Inhalant anesthetics must be used with scavenging devices.
One acceptable scavenging method is the use of a downdraft table.
Chemical fume hoods, charcoal canisters or Type IIB2 biosafety cabinets, which are vented to the outside, can also be used.
Note that charcoal canisters must be weighed before, and after, each use.
Most must be replaced after an increase in the recommended weight.
Depending on the size of the canister and the vendor, the canister should also be weighed during especially long procedures to assure
its continued effectiveness.
Contact your veterinarian for further training in the appropriate use of the anesthesia machines
available within your facility.
Anesthetized animals must be closely monitored during the procedure to assure that they are
maintained in the proper anesthetic plane.
If the anesthetic plane is too light, the animals may start to move or struggle.
If the anesthetic plane is too deep, the animals may die.
Once the anesthesia has been administered and enough time has elapsed for it to take effect, the anesthetic
plane can be assessed by pinching the toe, tail or ear of the animal.
Any reaction from the animal indicates that the anesthesia is too light and that additional anesthesia should be given.
The respiration and color of the mucous membranes and exposed tissue of the animal should
also be closely monitored. The respiration rate should be even.
An increased respiration rate is a sign that the anesthesia is too light.
A deep, shallow, decreased or irregular respiration rate is indicative of anesthesia that is too deep.
The color of the mucous membranes and exposed tissues should be bright pink to red. Dusky gray or blue color is indicative of an anesthetic
plane that is too deep.
Core body temperature can also be monitored in rodents.
The most common anesthetic complication is hypothermia.
Measures must be taken to control the body temperature before,
during and after anesthesia.
There are several choices of warming devices.
Your veterinarian should be consulted to determine the appropriate equipment to meet your research needs.
During recovery, the animal should be placed on clean, dry gauze or paper toweling to avoid
contact with the bedding, which may be inadvertently inhaled and result in asphyxiation.
Recovery from anesthesia can also be aided by the administration of warmed fluids given
subcutaneously or intraperitoneally.
Your veterinarian should be consulted for appropriate volumes and routes of administration.
Once the animal has reached sternal recumbency and appears to be making a normal recovery,
it may be returned to the animal holding area.
Animals should continue to be monitored closely
for several days following the procedure.
It is important to identify the cages with “Special Watch” cards or other similar methods so that particular attention will be afforded these
animals during the daily health checks.
A surgical record of some form must also be kept when
surgery is performed on rodents.
These records must be maintained for at least three years
following closure of the protocol.
Depending on the nature of the procedure, it may be beneficial to monitor weight and hydration for several days.
A softer, more palatable diet and/or fluids may be necessary.
- Consult your veterinarian for advice on the selection and administration of anesthesia
- The use of anesthetics and or analgesics must be described in your Animal Study
- It is important to weigh animals prior to dosing.
- Use a topical hypothermic prior to performing tail snips.
- Use a topical analgesic prior to performing retro-orbital sinus blood collection.
- Be certain to follow NHGRI Guideline 03.2 when preparing or handling Avertin.
Source: US National Institutes of Health