Mouse and Rat Anesthesia and Analgesia

Judith A. Davis1

1 NIAAA/NIH, Rockville, Maryland
Publication Name:  Current Protocols in Neuroscience
Unit Number:  Appendix 4B
DOI:  10.1002/0471142301.nsa04bs42
Online Posting Date:  January, 2008
GO TO THE FULL TEXT: PDF or HTML at Wiley Online Library


Many animal models used in neuroscience research must be surgically created and/or anesthetized for imaging studies. The purpose of this unit is to review the advantages and disadvantages of various anesthetic and analgesic agents in rodents; to discuss state‐of‐the‐art methods for monitoring anesthesia; and to provide tips for troubleshooting problems with anesthesia. Curr. Protoc. Neurosci. 42:A.4B.1‐A.4B.21. © 2008 by John Wiley & Sons, Inc.

Keywords: mouse; rat; anesthesia; analgesia

PDF or HTML at Wiley Online Library

Table of Contents

  • Introduction
  • Basic Protocol 1: Injectable Anesthesia for Mouse and Rat
  • Basic Protocol 2: Inhalant Anesthesia Using Isoflurane for Mouse and Rat
  • Basic Protocol 3: Analgesia for Mice and Rats
  • Commentary
  • Literature Cited
  • Figures
  • Tables
PDF or HTML at Wiley Online Library


Basic Protocol 1: Injectable Anesthesia for Mouse and Rat

  • Laboratory mice or rats
  • Anesthetic of choice (see Table 4.0.2)
  • Petroleum‐based artificial tear ointment
  • Lactated Ringer's solution, warmed to 35° to 36°C (optional; e.g., A.J. Buck or J.A. Webster)
  • For the surgeon:
    • Clean laboratory coat
    • Face mask
    • Head cover (optional)
    • Sterile gloves
  • Warm water recirculating heating pad or heat lamp, 38° ± 2°C
  • Sterile drape (optional)
  • Cages for mice or rats
  • Laboratory scale or balance (capacity 800 g; accuracy 0.1 g)
  • 1‐ or 3‐ml syringe
  • 20‐ or 22‐G needle
  • 21‐ or 23‐G needle (optional)
  • Additional reagents and equipment for intraperitoneal and subcutaneous injection ( appendix 4F)
    Table 0.b.2   MaterialsGuidelines for Injectable Anesthetics in Rodents

    Agent Species Dosage (mg/kg) and route of administration c Duration of surgical anesthesia (min) d
    Pentobarbital e Mouse 40‐70, i.p. 20‐40
    Rat 30‐50, i.p. 15‐60
    Ketamine/xylazine f Mouse 60‐100 ketamine + 5‐7.5 xylazine, i.p. 20‐25
    Rat 50‐90 ketamine + 5‐10 xylazine, i.p. 60‐80
    Ketamine/xylazine/acepromazine g Mouse 25‐30, i.p. 20‐35
    Rat 30‐40, i.p. 20‐45
    Ketamine/medetomidine h Mouse NA NA
    Rat 75 ketamine + 0.5 medetomidine, i.p. 15‐60
    Tribromoethanol j Mouse 125‐160, i.p. 15‐30
    Rat 300, i.p. 15‐20
    Atropine k Mouse 0.02‐0.05 mg/kg, s.c., i.p. 30‐40
    Rat 0.04 mg/kg, s.c., i.p. 15‐20
    Bupivicaine (Marcaine) l Mouse 0.5% solution. Local infiltration or drop‐wise administration to incision site. NA
    Rat NA

     cDosages are from a variety of sources. The dose may need to be adjusted for individual situations. Abbreviations: i.p., intraperitoneally; s.c., subcutaneously.
     dDuration of surgical anesthesia (unconscious, nonresponsive to painful stimuli) is not the same as “sleep time” (quiet, not moving, but responsive to stimuli), which is generally much longer.
     eDilute 1:10 (v/v) with sterile saline. This is a controlled substance. Barbiturate classification (long, short, ultrashort) is misleading, as species differences in barbiturate pharmacokinetics are responsible for significant variation in duration of action. As barbiturates do not provide analgesia, they are often combined with sedatives or tranquilizers to provide deep anesthesia and smooth recovery.
     fKetamine and xylazine can be safely mixed and given as a single injection. After mixing, dilute 1:10 (v/v) with sterile saline. Ketamine is a controlled drug. Caution: Some strains of rats are at increased risk for developing post‐anesthetic corneal lesions with this drug combination (Turner and Albassam, ).
     gMix 1.5 ml (100 mg/ml) ketamine, 1.5 ml (20 mg/ml) xylazine, and 0.5 ml (2 mg/ml) acepromazine together; stable at room temperature (shelf life of the ingredients). Do not use in preweanling animals. Ketamine is a controlled drug.
     hKetamine/medetomidine provides marked differences in surgical anesthesia between male and female mice and rats. Male mice require less ketamine whereas female mice require a higher dose of ketamine to effect loss of righting reflex. The loss of righting reflex is also gender specific, occurring more rapidly in males than in females. Males have loss of reflexes for 25‐60 min; females, 140‐150 min. Sleep time is also marked; males 135‐160 min, females 240‐300 min. Heavy urination occurs, which may lead to dehydration, indicating the need for parenteral fluids. Wetting of fur may also lead to hypothermia. Ketamine is a controlled drug. NA, data not available. The effects of this drug combination can be reversed with atipamezole (Antisedan, Pfizer). CAUTION: This drug combination does not produce an adequate plane of anesthesia for surgery! It does provide adequate chemical restraint (Cruz et al., ).
     JMake a 100% (w/v) stock solution by dissolving 5 g of 2,2,2‐tribromoethanol in 5 ml 2‐methyl‐2‐butanol (tert‐amyl alcohol). Gentle heating (50°C) provides better solubility. The anesthetic solution should be freshly prepared from the stock solution by adding 1.25 ml of stock solution to 48.75 ml of sterile saline. Both stock and anesthetic solutions are stable 2 to 4 months at 4°C in a dark bottle. Anesthetic solutions should be made fresh weekly; filter the solution using a 0.2‐µm filter. CAUTION: Stored solutions often deteriorate to irritant solutions that cause peritonitis and/or death following i.p. administration. Add 1 drop of Congo Red (0.1% w/v) to 5 ml of anesthetic solution. Purple color developing at pH < 5 indicates decomposition (Papaioannou and Fox, ).
     kAtropine is an anticholinergic, not an anesthetic. Atropine blocks acetylcholine at muscarinic receptors. Desirable effects include reduction in bronchial secretions and protection of the heart from vagal stimulation, which may occur during surgical procedures. If used, it should be given 5 to 10 min before the anesthetic agent.
     lBupivicaine applied to the incision site during or after closure of the incision augments analgesia by providing local anesthesia at the site of the incision. The local anesthetic effect may last for 10‐12 hr.

Basic Protocol 2: Inhalant Anesthesia Using Isoflurane for Mouse and Rat

  • Isoflurane
  • Laboratory mice or rats
  • Petroleum‐based artificial tear ointment
  • Lactated Ringer's solution, warmed to 35° to 36°C (optional)
  • For the surgeon:
    • Clean laboratory coat
    • Face mask
    • Head cover (optional)
    • Sterile gloves
  • 6‐, 20‐, or 30‐ml syringe (optional)
  • Latex glove (optional)
  • 30‐ or 60‐ml syringe case cover and cap
  • 3/8–in. brass tube sleeve insert (optional; Anderson Barrows PB96760PT, no. 25080)
  • 3/8‐in. (i.d.) vinyl or polyethylene tubing (optional)
  • #6–32, 2 in. machine screw (optional)
  • Commercial rodent face mask (optional; Kent Scientific or SurgiVet)
  • Warm water recirculating heating pad or heat lamp, 35° ± 1°C
  • Sterile drape (optional)
  • Precision vaporizer (Kent Scientific or SurgiVet)
  • Oxygen tank with flowmeter (Kent Scientific or SurgiVet)
  • Induction chamber (20‐cm length × 10‐cm height × 10‐cm width; 2 liters) with inlet and outlet ports
  • Additional reagents and equipment for subcutaneous injection ( appendix 4F)
All procedures with isoflurane require a scavenging device for expired anesthetic gases. Frequently, a chemical fume hood, down draft table, or safety cabinet is used for continuous exhaust of anesthetic gases away from personnel (see ).
PDF or HTML at Wiley Online Library



Literature Cited

   Abbott, F.V. and Bonder, M. 1997. Options for management of acute pain in the rat. Vet. Rec. 140:553‐557.
   Ahmed, F., Lundin, G.G., and Shire, J.G.M. 1989. Lysosomal mutations increase susceptibility to anesthetics. Experientia 45:1133‐1135.
   Alexander, C.M., Teller, L.E., and Gross, J.B. 1989. Principles of pulse oximetry: Theoretical and practical considerations. Anesth. Analg. 68:368‐376.
   Basbaum, A.I. and Levine, J.D. 1991. Opiate analgesia. N. Engl. J. Med. 325:1168‐1169.
   Clark, J.A., Myers, P.H., Goelz, M.F., Thigpen, J.E., and Forsythe, D.B. 1997. Pica behavior associated with buprenorphine administration in the rat. Lab. Anim. Sci. 47:300‐303.
   Cooper, D.M., DeLong, D., and Gillett, C.S. 1997. Analgesic efficacy of acetaminophen and buprenorphine administered in drinking water of rats. Contemp. Top. Lab. Anim. Sci. 36:58‐62.
   Cruz, J.I., Loste, J.M., and Burzaco, O.H. 1998. Observations on the use of medetomidine/ketamine and its reversal with atipamezole for chemical restraint in the mouse. Lab. Anim. 32:18‐22.
   Flecknell, P.A. 1996. Laboratory Animal Anesthesia, 2nd ed. Academic Press, London.
   Flecknell, P.A. 1998. Analgesia in small mammals. Semin. Avian Exotic Pet Med. 7:41‐47.
   Flecknell, P.A., Kirk, A.J.B., Fox, C.E., and Dark, J.H. 1990. Long‐term anesthesia with propofol and alfentanil in the dog and its partial reversal with nalbuphine. J. Assoc. Vet. Anesth. 17:11‐16.
   Flecknell, P.A., Roughan, J.V., and Stewart, R. 1999. Use of oral buprenorphine (“buprenorphine jello”) for postoperative analgesia in rats—a clinical trial. Lab. Anim. 33:169‐174.
   Gwynne, B.J. and Wallace, J. 1992. A modified anesthetic induction chamber for rats. Lab. Anim. 26:163‐166.
   Hecker, B.R., Lake, C.L., DiFazio, C.A., Moscicki, J.C., and Engle, J.S. 1983. The decrease in minimum alveolar concentration produced by sufentanil in rats. Anesth. Analg. 62:987‐990.
   Kufoy, E.A., Vytautas, A.P., Parks, C.D., Wells, A., Yang, C., and Fox, A. 1989. Keratoconjunctivitis sicca with associated secondary uveitis elicited in rats after systemic xylazine/ketamine anesthesia. Exp. Eye Res. 49:861‐871.
   Liles, J.H. and Flecknell, P.A. 1993. The effects of surgical stimulus on the rat and the influence of analgesic treatment. Br. Vet. J. 149:515‐525.
   Liles, J.H., Flecknell, P.A., Roughan, J., and Cruz‐Madorran, I. 1988. Influence of oral buprenorphine, oral naltrexone or morphine on the effects of laparotomy in the rat. Lab. Anim. 32:149‐161.
   Lovell, D.P. 1986a. Variation in pentobarbitone sleep time in mice. 1. Strain and sex differences. Lab. Anim. 20:85‐90.
   Lovell, D.P. 1986b. Variation in pentobarbitone sleep time in mice. 2. Variables affecting test results. Lab. Anim. 20:91‐96.
   Lovell, D.P. 1986c. Variation in barbiturate sleeping time in mice. 3. Strain X environment interactions. Lab. Anim. 20:307‐312.
   Papaioannou, V.E. and Fox, J.G. 1993. Efficacy of tribromoethanol anesthesia in mice. Lab. Anim. Sci. 43:189‐192.
   Smith, W. 1993. Responses of laboratory animals to some injectable anesthetics. Lab. Anim. 27:30‐39.
   Thurmon, J.C., Tranquilli, W.J., and Benson, G.J. 1996. Preanesthetics and anesthetic adjuncts. In Lumb and Jones' Veterinary Anesthesia, 3rd ed. (J.C. Thurmon, W.J. Tranquilli, and G.J. Benson, eds.) pp. 183‐209. Williams & Wilkins, Baltimore.
   Turner, P.V. and Albassam, M.A. 2005. Susceptibility of rats to corneal lesions after injectable anesthesia. Comp. Med. 55:175‐182.
   Vender, J.R., Hand, C.M., Sedor, D., Tabor, S.L., and Black, P. 1995. Oxygen saturation monitoring in experimental surgery: A comparison of pulse oximetry and arterial blood gas measurement. Lab. Anim. Sci. 45:211‐215.
Key References
   Bishop, Y.(ed). 1998. The Veterinary Formulary, 4th ed. Pharmaceutical Press, London.
  Provides dosages for a wide range of drugs, including antibiotics, for commonly used laboratory animals.
   Dorsch, J.A. and Dorsch, S.E. 1999. Understanding Anesthesia Equipment, 4th ed. Williams & Wilkins, Baltimore.
  This text is the most comprehensive source of information for equipment used in the United States.
   Flecknell, 1996. See above.
  This book offers practical advice for anesthesia and analgesia of laboratory animals. It is considered a “must have” for laboratory animal veterinarians.
   Kohn, D.F., Wixson, S.K., White, W.J., and Benson, G.J. (eds). 1997. Anesthesia and Analgesia in Laboratory Animals. American College of Laboratory Animal Medicine Series, Academic Press, New York.
  This is an overall excellent reference.
Internet Resources
  This is an excellent site for finding information about equipment mentioned in this unit, as well as finding other resources. Examples of major headings: Neuroscience, Cardiovascular, Respiratory Parameters, Data Acquisition, Recording Devices, Perfusion. The company's personnel are excellent at working with investigators to develop customized equipment.
  Another excellent site for all product information related to surgery. For example, several different types of scavenging systems for waste gas evacuation can be found at this site.‐e‐
  An excellent site for different sources of items such as catheters and infusion pumps.
  This site is provided for investigators wishing to explore alternative methods to deliver long‐term drugs, analgesics, and so on, in time‐release form to avoid repeated injections and maintain therapeutic concentrations.
  An excellent resource for customized (and certified) diets (all species). They also specialize in enrichment treats and medicated dosing systems.
PDF or HTML at Wiley Online Library