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Mouse and Rat Anesthesia and Analgesia
Judith A. Davis1
1

APPENDIX 4B

NIAAA/NIH, Rockville, Maryland

ABSTRACT Many animal models used in neuroscienc

e 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 stateof-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. C 2008 by John Wiley & Sons, Inc. Keywords: mouse r rat r anesthesia r analgesia

INTRODUCTION
Management of rodent anesthesia and analgesia presents a formidable challenge due to the animals’ small size and variability in sensitivity to anesthetics and analgesics. A common mistake is to focus on the surgical procedure while giving little attention to selection of an appropriate anesthetic or the animal’s physiological status during the procedure and post-operative period. Preparation is critical with small animals, especially since most investigators perform multiple surgeries in a single session with no assistance. Investigators should familiarize themselves with the available types of anesthetic agents, their physiologic effects, and the risks involved in their use. Some anesthetics have a wide margin of safety—i.e., a wide range of doses produce the desired effect (anesthesia) with minimal undesired effects (e.g., death)—and can be used by inexperienced personnel, whereas others have a narrow margin of safety and require experienced personnel. Proper preparation prior to surgery, coupled with appropriate monitoring of the animal both during and after the procedure, will contribute to successful creation of the surgical model. This unit provides protocols for injectable anesthesia for the mouse and rat (see Basic Protocol 1), inhalant anesthesia for the mouse and rat (see Basic Protocol 2), and analgesia for the mouse and rat (see Basic Protocol 3). The Commentary describes advantages and disadvantages associated with each anesthetic method and provides tips for successful monitoring of the animal. The Commentary further describes aspects of rodent physiology (see Table A.4B.1) that must be considered to prevent common problems associated with anesthesia in rodents. The advantages for using analgesia in rodents are also discussed, along with a few strategies for studies in which commonly used analgesic agents are contraindicated. NOTE: An external heat source should be provided to all anesthetized animals to maintain body temperature. This can be accomplished by using recirculating warm water blankets, heat lamps, forced warm air units, or pocket warmers. NOTE: All protocols using live animals must ?rst be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow of?cially approved procedures for the care and use of laboratory animals.

Animal Techniques
Current Protocols in Neuroscience A.4B.1-A.4B.21, January 2008 Published online January 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/0471142301.nsa04bs42 Copyright C 2008 John Wiley & Sons, Inc.

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Table A.4B.1 Normal Physiological Parametersa of Mice and Rats

Mouse Temperature Heart rate Respiratory rate Tidal volumeb Minute ventilationb Body surface area Blood volume 36.5? -38.0? C 300-650 beats/min 150-220 breaths/min 0.09-0.23 ml 24 ml 10.5 × (wt in g)2/3 75 ml/kg

Rat 37.0? -38.0? C 250-370 beats/min 70-115 breaths/min 0.6-2.0 ml 200 ml 10.5 × (wt in g)2/3 58 ml/kg

a Published parameters vary, which probably re?ects differences in strain, age, and gender of the species. b Tidal volume is the volume of gas drawn into the respiratory tract with each breath. Minute ventilation is the volume of

gas breathed in 1 min (tidal volume × respiratory rate). Rodents maintain minute ventilation by high respiratory rate and low tidal volume. During anesthesia, both parameters fall, causing a decrease in minute ventilation

BASIC PROTOCOL 1

INJECTABLE ANESTHESIA FOR MOUSE AND RAT
Because mice and rats have high metabolic rates, they require higher anesthetic dosages than larger animals with low metabolic rates to achieve an effective level of anesthesia. The duration of anesthesia is typically shorter (20 to 30 min), and mice and rats are less likely to survive respiratory arrest from overdosage. The high metabolic rate also increases the risk of hypothermia and dehydration due to exposed membranes and small body size. Usually a group of mice or rats are anesthetized simultaneously or serially to allow short surgical procedures on a group of animals. This format is amenable to injectable anesthetics, because relatively inexperienced personnel can rapidly administer the agent. These drugs are convenient to use since many are readily available as noncontrolled substances and require no special equipment. Also, minimal cost is incurred in parenteral anesthetic delivery. Regardless of which injectable anesthetic is used, each animal must be weighed and dosed accordingly. Similarly, once the animal loses sternal recumbency an external heat source should be used to help maintain body temperature. Rodents do not require fasting prior to administration of anesthetic as they cannot vomit. CAUTION: Once a chemical anesthetic is given, it is dif?cult to adjust the dose. In some animals the standard dose may be inef?cient to achieve a stable plane of anesthesia while in other animals the same dose may be excessive and may lead to mortality. Repeated doses of some drugs may lead to delayed toxicity or more acutely an overdose and death.

Materials Laboratory mice or rats Anesthetic of choice (see Table A.4B.2) Petroleum-based arti?cial 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)

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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)

1. Prepare the operating area so it is clean, uncluttered, and provides adequate room for instruments, equipment, recirculating warm water blanket or lamp, and, if required, a stereotaxic apparatus. Ensure that the area is well lit. 2. Set up a recirculating warm water blanket or lamp and turn it on. If using a recirculating warm water blanket, cover it with a sterile drape. 3. Retrieve the animal from its cage, place it in a tared container, and determine its weight. 4. Return the animal to its cage. 5. Calculate the appropriate dosage of the chosen anesthetic using Table A.4B.2. Fill either a 1- or 3-ml syringe with selected anesthetic.
For example, according to Table A.4B.2, for a 20-g mouse, the amount of pentobarbital to inject is 0.020 kg × 40 to 70 mg/kg = 0.8 to 1.4 mg.

6. Manually restrain the animal and inject the selected dose intraperitoneally (APPENDIX 4F) using a 22-G needle for mice or 20-G needle for rats.
Proper intraperitoneal injection requires that the animal be held with its head tilted downward so that the needle is inserted into the lower left abdominal quadrant. Do not inject on the midline due to the risk of puncturing the bladder.
Table A.4B.2 Guidelines for Injectable Anesthetics in Rodents

Agent Pentobarbitalc Ketamine/xylazine
d

Species Mouse Rat Mouse Rat

Dosage (mg/kg) and route of administrationa 40-70, i.p. 30-50, i.p. 60-100 ketamine + 5-7.5 xylazine, i.p. 50-90 ketamine + 5-10 xylazine, i.p. 25-30, i.p. 30-40, i.p. NA 75 ketamine + 0.5 medetomidine, i.p. 125-160, i.p. 300, i.p. 0.02-0.05 mg/kg, s.c., i.p. 0.04 mg/kg, s.c., i.p.

Duration of surgical anesthesia (min)b 20-40 15-60 20-25 60-80 20-35 20-45 NA 15-60 15-30 15-20 30-40 15-20
continued Animal Techniques

Ketamine/xylazine/ acepromazinee Ketamine/ medetomidinef

Mouse Rat Mouse Rat

Tribromoethanolg Atropine
h

Mouse Rat Mouse Rat

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Table A.4B.2 Guidelines for Injectable Anesthetics in Rodents, continued

Agent Bupivicaine (Marcaine)i

Species Mouse

Dosage (mg/kg) and route of administrationa 0.5% solution. Local in?ltration or drop-wise administration to incision site.

Duration of surgical anesthesia (min)b NA

Rat
intraperitoneally; s.c., subcutaneously.

NA

a Dosages are from a variety of sources. The dose may need to be adjusted for individual situations. Abbreviations: i.p., b Duration 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.
c Dilute 1:10 (v/v) with sterile saline. This is a controlled substance. Barbiturate classi?cation (long, short, ultrashort) is

misleading, as species differences in barbiturate pharmacokinetics are responsible for signi?cant 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. d Ketamine 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, 2005). e Mix 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. f Ketamine/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 re?ex. The loss of righting re?ex is also gender speci?c, occurring more rapidly in males than in females. Males have loss of re?exes 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 ?uids. 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, P?zer). CAUTION: This drug combination does not produce an adequate plane of anesthesia for surgery! It does provide adequate chemical restraint (Cruz et al., 1998). g Make 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; ?lter the solution using a 0.2-?m ?lter. 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, 1993). h Atropine 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. i Bupivicaine 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.

7. Return the animal to an empty cage and monitor for the depth of anesthesia (e.g., loss of righting re?ex, loss of palpebral blink re?ex, rate and depth of respiration). Do not put the animal in a cage containing other animals.
A common problem at this stage results from impatience. If suf?cient time is not allowed for the anesthetic to take effect, the animal will move or struggle when the procedure begins. Inexperienced personnel may make the mistake of redosing, because it seems that the animal is not suf?ciently anesthetized. A reasonable amount of time to wait before administering a second dose of injectable anesthetic is 10 min. However, this parameter is very dif?cult to judge; many variables in?uence anesthesia, such as strain, gender, body weight, age, and choice of injectable anesthetic. Overdosage of injectable anesthetics is a frequent cause of death.

8. Apply a petroleum-based arti?cial tear ointment to the eyes to prevent corneal desiccation.
Mouse and Rat Anesthesia and Analgesia

9. Optional: Immediately prior to making the surgical incision, inject 60 ml/kg prewarmed (35? C) lactated Ringer’s solution under the skin (subcutaneously between

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the shoulder blades; APPENDIX 4F) to provide maintenance hydration. Use a 23-G needle for mice, or a 21-G needle for rats.
It is important that ?uids be warmed before administering to avoid hypothermia. This can be achieved by running hot water over the bag containing the lactated Ringer’s solution for 3 min, using a microwave oven (low setting), or placing the bag in an incubator prior to use. The ?uid should be tested before use; it should feel warm, not hot, to the touch (35? to 36? C).

INHALANT ANESTHESIA USING ISOFLURANE FOR MOUSE AND RAT
The most consistent and reliable anesthetic protocols for rodents involve inhalant anesthesia. Mastering delivery and maintenance of inhalant anesthesia is bene?cial in many ways. Inhalational anesthetic agents allow greater control over depth and duration of anesthesia, thus providing a greater survival rate. Also, the animal recovers quickly (saving time), and physiologic responses of the surgical model are more reliable and consistent. Another advantage is the ability to choose an anesthetic agent that requires minimal metabolism, biotransformation, or excretion. These advantages translate into minimized variability within the experimental model. Inhalant anesthetic concentrations can be adjusted rapidly to maintain appropriate anesthesia; most inhalants (iso?urane) are minimally absorbed, and the recovery period is short. For example, iso?urane is a relatively safe anesthetic for both the surgeon and the animal and is commonly used in rodent surgery. Gas anesthesia is simple to use, but does require special equipment. Delivery of volatile anesthetic agents to rodents is usually done using a face mask, since rodents’ small size makes endotracheal intubation more dif?cult than in larger species like a dog or cat. Use of a face mask is easy and unskilled persons can readily master placement of a face mask. Face masks may have to be specially designed if they are going to be used with other devices (e.g., bite bars for stereotaxic surgery). Commercial face masks can also be purchased. An endotracheal tube (ETT) allows control of the animal’s airway in the event of cardiac or respiratory arrest, as the animal can be arti?cially ventilated. However, a high level of technical competence is needed to consistently insert an ETT, and the equipment for scavenging anesthetic gas to ensure operator safety may not be available. Attempts to intubate rodents, even by experienced personnel, often traumatize the oropharynx, larynx, trachea, and at times, the esophagus. CAUTION: Use of a face mask instead of endotracheal intubation makes positive pressure ventilation less effective in the case of an emergency.

BASIC PROTOCOL 2

Materials Iso?urane Laboratory mice or rats Petroleum-based arti?cial 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)
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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 Scienti?c or SurgiVet) Warm water recirculating heating pad or heat lamp, 35? ± 1? C Sterile drape (optional) Precision vaporizer (Kent Scienti?c or SurgiVet) Oxygen tank with ?owmeter (Kent Scienti?c 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)
CAUTION: All procedures with iso?urane 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 Critical Parameters).

Mouse and Rat Anesthesia and Analgesia

Figure A.4B.1 Assembly of a simple face mask for use in rodent anesthesia. (A) Syringe with attached rubber glove ?nger. (B) Hole cut in glove ?nger tip to ?t over animal’s muzzle.

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Prepare work area 1. Prepare the operating area so it is clean, uncluttered, and provides adequate room for instruments, equipment, recirculating warm water blanket or lamp, and, if required, a stereotaxic apparatus. Ensure that the area is well lit. Make face mask (optional): two types Type 1: For procedures that do not require head stabilization (i.e., stereotaxic apparatus): 2. Remove the plunger from a 6-ml syringe (for mice) or a 20- or 30-ml syringe (for rats). Cut the syringe barrel off at approximately the 4- to 5-ml mark.
3. Cut a ?nger from a latex glove and stretch it over the cut end of the syringe (Fig. A.4B.1A). With scissors, cut a small hole (1- to 3-mm diameter) in the tip of the ?nger (Fig. A.4B.1B).
This simple face mask allows expired gas to escape around the animal’s mouth and muzzle; therefore, the animal should be placed on a down-draft table or in a fume hood, or an external gas scavenger should be placed near the head of the animal for the surgeon’s safety.

Type 2: For procedures that require head stabilization (i.e., stereotaxic apparatus): 4. To make an inhalation mask for a rat stereotaxic frame, ?rst:
a. Cut the top 6 cm off a 60-ml syringe case (see Fig. A.4B.2 for inhalation mask components). b. Cut a 3-cm hole in the center of the 60-ml syringe case cap and then cut a “D-shaped” piece out of the side of the cap to allow better visibility of the rat’s nose/mouth through the syringe case.

Figure A.4B.2 Components, made from readily available equipment, needed to make a rat face mask to use with a stereotaxic apparatus. A similar mouse face mask can be made using smaller syringe case components.

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c. Cut a 3.5 cm-diameter circle (approximate) from the palm or sleeve of a latex glove to use as a face diaphragm just under the syringe case cap. d. Place a 3/8-in. brass sleeve insert through the syringe case just behind the front cap for anesthesia supply gas and another 1/4 in. brass sleeve insert just in front of the rear cap for the exhaust gas to exit the mask. Use 3/8 in. (i.d.) vinyl tubing to connect the brass nipples to the precision gas vaporizer and exhaust gas scavenging system. e. The rear cap that ?ts around the stereotaxic mouth bar is made from a 20-ml syringe case cap to form the rear of the face mask. Cut an inverted “T-shaped” hole in the center of the rear cap to match the size and shape of the upright portion of the stereotaxic mouth bar. f. Place the rear cap over the mouth bar and nose clamp and insert the 60-ml syringe case into the rear cap. Now, place the latex diaphragm over the large end of the 60-ml syringe case and hold it in place by sliding the 60-ml syringe case cap over the diaphragm and syringe case. Ensure the “D-shaped” cut out is positioned on top so you can see the rat’s nose and nose clamp more clearly inside the syringe case. If necessary, lengthen the nose clamp adjustment screw by using a longer screw, or cut the head off a similar sized screw (e.g., #6-32, 2-in. machine screw) and hold the two screws together using a piece of tight ?tting tubing. g. After you stretch the latex diaphragm across the cut end of syringe case, cut a small hole (1- to 3-mm diameter) in the center of the diaphragm for the rat’s (or mouse) nose/mouth. (Fig. A.4B.3)

Mouse and Rat Anesthesia and Analgesia

Figure A.4B.3 The face mask assembled and attached to the stereotaxic apparatus. The anesthetized animal’s nose can now be placed through the diaphragm, positioning the animal’s upper incisors through the incisor hole of the mouth bar.

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5. To use the inhalation face mask, clip and prep the surgical site in an area away from the stereotaxic frame using a separate anesthesia circuit. Adjust the nose clamp up enough to easily accept the animal’s nose between the nose clamp and the mouth bar. Place the animal’s nose through the latex diaphragm and position the animal’s upper incisors through the incisor hole of the mouth bar. Insert and adjust the ear bars, then tighten the nose clamp and level the nose with the mouth bar height adjustment.
If performing mouse stereotaxic surgery, follow the above instructions but use sizes 50% smaller for the different parts of equipment; e.g., 30-ml syringe case.

Prepare induction chamber 6. Plug in and turn on a recirculating warm water blanket or lamp. If using a recirculating warm water blanket, cover it with a sterile drape.
7. Pour iso?urane into the precision vaporizer to the ?ll line. 8. Connect the vaporizer and oxygen tank to an induction chamber. 9. Turn the oxygen valve on to provide high ?ow (100% oxygen) to precharge the chamber.

Deliver anesthesia 10. Retrieve the animal from its cage and place it in the induction chamber. Expose the animal to 100% oxygen for 2 to 3 min, and then adjust the precision vaporizer (dial) to provide 5% iso?urane.
The chamber should be rapidly ?lled for fast induction. If the volume of the chamber is known, the required ?ll time can be estimated as follows: chamber volume/?ow rate = time to ?ll. For example, if the ?ow rate is 5 liters/min and the chamber volume is 10 liters, the ?ll time is 2 min.

11. Monitor the animal until it becomes ataxic, recumbent, and immobile, which signi?es the onset of anesthesia. 12. Remove the animal from the induction chamber and place the face mask over the muzzle of the animal to maintain anesthetic delivery (Fig. A.4B.4).
If necessary, use Stomahesive Paste (ConvaTec) to seal the face mask around the animal’s muzzle for a tight ?t.

13. Connect the tubing from the induction chamber to the face mask, adjust dial on the vaporizer to deliver 1.0% to 1.5% iso?urane, and adjust the ?owmeter to deliver 1 liter of oxygen per minute for maintenance anesthesia.
Oxygen ?ow rate should be at least three times the minute ventilation rate (10 ml/kg or ?0.2 ml/mouse) to lower CO2 .

14. Apply a petroleum-based arti?cial tear ointment to the eyes to prevent corneal desiccation.
Keratitis sicca (drying of the cornea) frequently occurs secondary to high-?ow delivery of anesthetic via the face mask (Kufoy et al., 1989). Use of the petroleum-based ointment protects the eyes.

15. Optional: Immediately prior to making the surgical incision, inject 60 ml/kg prewarmed (35? to 36? C) lactated Ringer’s solution under the skin (subcutaneously between the shoulder blades; APPENDIX 4F) to provide maintenance hydration. Use a 23-G needle for mice, or a 21-G needle for rats.
It is important that ?uids be warmed before administering to prevent hypothermia. This can be achieved by running hot water over the bag containing lactated Ringer’s solution for 3 min, using a microwave oven (low setting), or placing the bag in an incubator prior

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Figure A.4B.4

Animal with face mask.

to use. The ?uid should be tested before use; it should feel warm (35? to 36? C), not hot, to the touch.

16. At the end of the procedure, turn the vaporizer off and deliver 100% oxygen to the animal for ?5 min.
This will speed recovery and prevent the development of hypercapnia (an increase in arterial carbon dioxide content). BASIC PROTOCOL 3

ANALGESIA FOR MICE AND RATS
During general anesthesia, the cerebral cortex, but not the rest of the nervous system, is depressed. A scalpel incision stimulates primary sensory afferents, transmitting nociceptive impulses to the spinal cord. Such activity sensitizes the nociceptor system to further input from injured tissue, and such sensitization outlasts the duration of the original stimulus. Once the animal is in pain, there is an increase in sympathetic activity; epinephrine increases the sensitivity of injured neurons to stimulation. A re?ex increase in muscle tension surrounding the area of damaged tissue also commonly occurs, which adds to the pain. Analgesia, in the strictest sense, is the absence of pain. Practically, this means trying to reduce the intensity of the pain perceived. The goal is to diminish the pain as much as possible without undue depression of the animal. Intraoperative as well as post-operative analgesics should be provided during experimental manipulations. Many commonly used injectable anesthetics, such as pentobarbital or ketamine, have poor analgesic potency. Immobility does not mean that pain or distress is not present. Many injectable anesthetic combinations (e.g., ketamine/xylazine) include an analgesic drug (xylazine). A common mistake is to assume that the analgesic component remains after the procedure is completed and the animal has recovered. In fact, most analgesics have very short half-lives in rodents. An additional analgesic dose should be given at the end of the procedure.
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Selection of an analgesic regimen depends upon the type of procedure performed, the nature of the pain (severity and duration), and the potential in?uence on research objectives. Many options are available (topical, regional, central, and peripherally acting), along with other means of protecting injured tissue (e.g., soft bedding, Elizabethan collars). It is dif?cult to make general recommendations concerning analgesia choices, since species and strain of animal, choice of anesthetic, and invasiveness of the procedure all warrant consideration. For example, if using iso?urane, buprenorphine should be given before surgery; if using an injectable anesthetic, it should be given at the end of surgery. If the procedure is simple, like a jugular catheterization, administration of a nonsteroidal anti-in?ammatory drug (NSAID) or application of a topical anesthetic, such as lidocaine, on the incision line may be bene?cial. When major surgery is completed, administration of buprenorphine (for 24 to 36 hr) followed by an NSAID alone for 24 hr is helpful. Good post-operative care is essential both for the animal’s welfare and because it is good scienti?c practice.

Preoperative Medications Selection of preoperative medications will help manage the animal by eliminating unnecessary anxiety and distress. Comments are provided here for preoperative medications, along with points to consider for analgesic selection. Table A.4B.3 lists guidelines for injectable analgesic agents in rodents.
Preoperative stress and pain can be relieved by administration of tranquilizers or analgesics, respectively. Benzodiazepines (diazepam, midazolam), classi?ed as minor tranquilizers, are commonly used as co-induction agents in combination with injectable
Table A.4B.3 Guidelines for Injectable Analgesics in Rodents

Agent Buprenorphineb Butorphanol
c

Species Mouse Rat Mouse Rat Mouse Rat Mouse Rat Mouse Rat Mouse Rat
d

Dosage (mg/kg) and route of administrationa 0.05-0.1 mg/kg, i.p., s.c. 0.01-0.05 mg/kg, i.p., s.c. 1.0-5.0 mg/kg, s.c. 2.0-2.5 mg/kg, s.c. 2.5 mg/kg, s.c. 1.1 mg/kg, s.c. 5 mg/kg, s.c. 5 mg/kg, s.c. 10 mg/kg, s.c. 2.5-5.0 mg/kg, s.c. 300 mg/kg, p.o. 100-300 mg/kg, p.o.

Frequency of dosage Every 6-12 hr Every 8-12 hr Every 4 hr Every 4 hr Every 12 hr Every 12 hr Every 12-24 hr Every 12-24 hr Every 12-24 hr Every 12-24 hr Every 4 hr Every 4 hr

Flunixin (Banamine) Ketoprofen Carprofen Acetaminophen

a Dosages are from a variety of sources. The dose may need to be adjusted for individual situations. Published dosages for

oral administration are generally much larger and, in some cases, are not effective (Flecknell et al., 1999). For mice, use a 1-ml syringe with a 22-G needle; for rats, use a 6-ml syringe with a 22-G needle. Abbreviations: i.p., intraperitoneally; p.o., perorally; s.c., subcutaneously. b Buprenorphine is a partial agonist that is dif?cult to reverse if ventilation becomes compromised. It may be prudent to begin with a low published dose and redose if necessary. This is a controlled substance. This drug increases locomotor activity and in rats, may cause pica (indiscriminate appetite, such as eating bedding). c This is a controlled substance with mild analgesic activity in rodents. d Published data for clinical trials is not available for mice; however, experimental tests using analgesiometry tests indicate that ketoprofen is an effective analgesic in mice (Flecknell, 1998).

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anesthetics. They do not provide analgesia, but have minimal effects on the cardiopulmonary, renal, and hepatic systems. However, benzodiazepines potentiate the respiratory depression induced by opioids, so attention should be given to the selected injectable anesthetic combinations. α-Adrenoceptor agonists (xylazine, detomidine, medetomidine) generally provide sedation and analgesia with some side effects, such as bradycardia, hypothermia, and respiratory depression. In mice and rats, sedation outlasts analgesia. Because mice and rats frequently hold their breath and have stress-induced catecholamine release, they are particularly susceptible to respiratory acidosis and hypoxemia. Dopram is a CNS respiratory stimulant that can be incorporated into the preoperative regimen by placing one full-strength drop on the animal’s tongue. Atropine (0.05 mg/kg intraperitoneally or subcutaneously) can be given ?10 min before anesthetic induction to prevent bradycardia; it acts in concert with Dopram as a respiratory stimulant. As a word of caution, the use of anticholinergics (atropine) in rodents is controversial. Atropine protects the heart from vagal inhibition or opioid-induced bradycardia, but also increases the viscosity of airway secretions (Flecknell, 1996). In rodents, increased viscosity of secretions may inadvertently obstruct small airways or the trachea. The respiratory pattern should be monitored closely when an anticholinergic is used, and the animal should be ventilated vigorously when an inhalant is used. Similarly, the heart rate should be monitored if anticholinergics are not used.

Analgesic Drugs Analgesics are best provided pre-emptively because sensitization of primary afferents increases the transmission of nociceptive impulses towards the CNS. If given before surgery, the analgesic diminishes the hyperexcitability of neurons that occurs with nociceptive stimulation, thus reducing the amount of post-operative analgesics needed. Some analgesics also potentiate the anesthetic effect, decreasing the amount (often by 40% to 60%) of inhalant anesthesia needed for surgical anesthesia. For example, many opioids have been shown to reduce the dosage of anesthetic required for surgical anesthesia (Hecker et al., 1983; Flecknell et al., 1990).
Narcotic (opioid) analgesia: Opioids act as agonists of presynaptic and post-synaptic receptors in the CNS; the af?nity of the opioid for the receptor correlates with analgesic ef?cacy (Basbaum and Levine, 1991). The primary advantage of opioids is profound analgesia, but they also induce CNS depression characterized by hypothermia, bradycardia, and respiratory depression. If clinically effective dosages are given, however, opioids have minimal effect on the rodent cardiovascular system. Bradycardia can be managed by administering atropine, using 0.04 mg/kg for mice or 0.05 mg/kg for rats (Flecknell, 1996). Opioid agonists/antagonists: Butorphanol has mild analgesic activity at κ receptors with marked antagonist activity at the ? receptor (Flecknell, 1996). It has a relatively short (3 to 4 hr) duration of effect. Buprenorphine hydrochloride is a partial ? agonist with prolonged duration (8 to 12 hr) of activity. Use of buprenorphine has an “anestheticsparing” effect, in that less anesthesia is required for the animal. Using a narcotic agent prior to surgery will also reduce the analgesic dosage required in the post-operative stages to keep the animal comfortable. One side effect, however, is that the animal will be sedated and, therefore, less likely to eat and drink while receiving these types of analgesics. This is an important point if weight and activity are to be monitored in the post-operative period as a means to assess pain and/or discomfort. Several studies suggest that buprenorphine should be used in rats no longer than 48 hr post-operatively (Liles et al., 1988; Liles and Flecknell, 1993).

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α2 agonist analgesia: α2 adrenergic agonists (e.g., xylazine, medetomidine) are generally regarded as sedative hypnotics with a wide range of effects, including sedation, visceral and somatic analgesia, mild to moderate muscle relaxation, peripheral vasoconstriction, bradycardia, and hypothermia. Due to the inhibitory effect on sympathetic out?ow, gastrointestinal and endocrine functions are also depressed. Species sensitivity and response to these drugs varies greatly. Xylazine, the most commonly used α2 agonist in mice and rats, has a rapid (3 to 5 min) onset of action and a short duration. Duration of sedation and analgesia is dosage dependent. Mice and rats are sedated for 1 to 2 hr, but analgesia is brief (15 min). Xylazine should be supplemented with other analgesic agents if prolonged (>30 min) analgesia is needed (Thurmon et al., 1996). Medetomidine, the most potent α2 agonist, has fewer side effects because it is more selective for α2 receptors. Although α2 agonist drugs are excellent additions to anesthetic injectable cocktails due to their prolonged sedative effects, their role as sole analgesic agents remains to be established in mice and rats (Abbott and Bonder, 1997). The detrimental side effects of these drugs can be reversed with the α2 adrenergic antagonist atipamezole (1.0 mg/kg subcutaneously, intraperitoneally). Before using a reversal agent, it is important to carefully consider the advantages and disadvantages of doing so. Once an α2 adrenergic antagonist is given, duration of anesthesia is markedly reduced and recovery of re?exes occurs within 5 min of administration. If given too soon, rapid loss of anesthesia with concomitant loss of muscle relaxation may occur prior to the end of the procedure. Another major disadvantage is the loss of analgesia for animals undergoing a painful procedure. In this situation, the analgesic can be augmented with another opioid or NSAID (see below) before the α2 agonist is reversed. Advantages of using a reversal agent, other than faster recovery time, are reversal of bradycardia and respiratory depression, thus reducing the occurrence of complications associated with prolonged sedation. Local anesthesia: Local anesthetics block Na+ channels in cell membranes, preventing depolarization and subsequent activation of peripheral nociceptors. Properties of the selected anesthetics, size of dose, and accuracy of injection placement in?uence duration of response. To reduce pain from local irritants, such as the ear bars of the stereotaxic apparatus, 2% lidocaine gel can be placed into the ear canal and on the ear bars. Topical anesthetics can also enhance analgesia during surgery. In?ltrating 0.1 to 0.2 ml of 2% mepivicaine or lidocaine into the skin along the edges of the incision will enhance analgesia and reduce the needed anesthetic dosage. If use of systemic analgesics is contraindicated by the experimental study, topical anesthesia provides pain relief at the incision site. Working with a veterinarian, pain medications can be formulated into topical gels and creams. Organogel, a lecithin-based matrix, is used to compound a 40% lidocaine cream (38% stronger than 2% commercial preparations) for studies confounded by narcotics and nonsteroidal drugs. A word of caution: the potential for toxicity (arrhythmia) does exist if the animal ingests the topical analgesic cream or ointment. Nonsteroidal anti-in?ammatory drugs (NSAIDs): NSAIDs act peripherally to suppress in?ammation and decrease production of kinins and prostaglandins. Use of NSAIDs allows faster anesthetic recovery and less post-recovery sedation while providing analgesia. Ketoprofen may be given twice daily, beginning with the initial anesthesia. Administering ketoprofen in warm lactated Ringer’s solution is an excellent way to provide both ?uids and analgesic simultaneously. Fluids provide needed hydration and protect against renal damage. NSAIDs are a good choice for craniotomies requiring a stereotaxic apparatus. Generally, post-operative pain is from the mechanical pressure (trauma) of the ear bars, not the craniotomy; NSAIDs are ideal for this type of pain. If NSAIDs (ketoprofen, carprofen) are to be used, one should consider including an H2 -blocking agent, such as Zantac (0.5 mg/g intramuscularly) or Pepcid (2.5 mg/kg subcutaneously) to protect

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against gastrointestinal damage. Since animals are more sensitive to the acute renal effects of NSAIDs after surgery, administration of warmed ?uids with an NSAID is advised to help protect the kidneys. Oral analgesia: Concerns about causing additional stress through handling animals following surgery and giving them injections have led many to try oral preparations of analgesic agents. These preparations also offer convenience, especially if the agent is placed in the drinking water. Several factors may in?uence the decision to use the oral route. First, high dosages are generally needed, due to metabolism of the drug following oral administration; high dosages of buprenorphine have been associated with consumption of bedding and gastric distension (Clark et al., 1997). Second, palatability is a concern. Water consumption following addition of acetaminophen reduced overall intake of water by rats in one study (Cooper et al., 1997) or failure to completely eat an analgesic-treated Jell-O cube (Flecknell et al., 1999). Third, surgery may depress food and water intake. An additional consideration is the method of housing; if more than one animal is housed per cage, one cannot ensure that each animal consumes his or her required dosage of drug.

COMMENTARY Background Information
Investigators struggle with the issue of anesthesia for a variety of reasons, including lack of training, unfamiliarity with the equipment required for sophisticated anesthesia protocols, and a perception that choice of anesthetic agent really does not matter. Complicating the issue is the size of the mouse or rat, which presents technically challenging problems that have led to acceptance of a mortality rate of 30% to 40%, which would be considered alarming in larger animals. The considerably higher success rate in human anesthesia highlights the inherent safety of properly administered anesthesia and the value of intense monitoring during recovery that occurs with humans. Comparisons to human anesthesia and monitoring suggest that investigators are often guilty of poor selection of anesthetic agents, as well as poor surgical techniques and insuf?cient physiological monitoring of rodents, both intra- and post-operatively. Hesitancy to anesthetize rodents can be overcome by becoming familiar with the selected anesthetic agent and its effects on the animal model. This information is especially important when determining dosages of injectable anesthetic agents. Adverse effects of chemical anesthesia are dif?cult to reverse, while insuf?cient anesthesia is dif?cult to safely augment. Published dosage regimens vary widely because many of these protocols come from laboratories with wide differences in complexity of surgical procedures, skills required, and, in some situations, acceptance of high mortality. Any published anesthetic dosage regimen must be tailored to the individual animal and experimental procedure.

Critical Parameters
Selection of an anesthetic regimen depends on a variety of factors, as discussed below. Whatever anesthetic regimen is chosen, it is important to remember that the purpose of anesthesia is to prevent pain and provide humane restraint. Selecting the method of anesthesia that is least likely to interfere with the aims of the research is dif?cult, and the physiological effects of the anesthetic agent should be researched. For example, it may be important to maintain blood pressure within a normal range. Many anesthetic agents (e.g., ether) maintain blood pressure by stimulating the sympathetic nervous system, so animals have elevated catecholamines. Is this important to the research project? Simply adopting a method of anesthesia described in publications for the model of interest is no guarantee that the proper regimen is used. Finally, although an appropriate anesthetic regimen should be selected for each individual research project, attention must also be given to management of the anesthetized animal throughout and following the procedure, so that the animal does not become hypothermic, hypoxic, or hypercapnic. Species variation Risk is inherent with anesthesia, regardless of choice of agent or animal species. Wide variation occurs among animal species in their responses to drugs, and extrapolation from

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one species to another should be avoided. The small size of mice and rats, coupled with the technical dif?culties encountered when trying to monitor physiological parameters, increases the risk of anesthesia in these two species. Variability of response to ketamine is particularly relevant. Small dosages of ketamine can immobilize nonhuman primates, but ketamine alone rarely immobilizes ruminants. In rodents, only doses that produce profound respiratory depression are effective and, at these dosages, mortality can exceed 50%. For this reason, ketamine should always be combined with a benzodiazepine or a α2 adrenergic agonist (xylazine). Because of the small size of rodents, dosages must be carefully calculated for each animal. Dosing based on an average weight will result in uneven anesthesia and possibly death due to variance of individual animal weights from the average. Most injectable agents must be diluted 5- to 10-fold for accurate dosing. Rodent metabolic rate The metabolic balance of small animals is precarious. Mice and rats frequently develop metabolic acidosis, hypoglycemia, and electrolyte disturbances when anesthetized. These physiologic responses underscore the vulnerability of small animals and accentuate the detrimental effects of catabolic responses to additional stress triggered by surgery. The high metabolic rate of rodents entails two risk factors. First, higher dosages of an anesthetic agent are required to achieve a stable plane of anesthesia. Second, the drugs are metabolized faster, shortening the duration of anesthesia. If the dose is increased in an effort to prolong anesthesia, the risk of killing the animal also increases. Aggressive monitoring is critical to avoid complications. It is important to overcome the tendency to focus on the procedure, neglecting the animal’s physiologic status, regardless of anesthetic regimen chosen. Environmental and drug interactions Because health status, age, strain, nutritional status, environment, and route of administration must all be considered, it is dif?cult to provide precise dosages of recommended injectable agents. A recommended drug dosage that provides adequate depth of anesthesia (loss of re?exes) in one animal is often insuf?cient in another animal due to animal variability of response to the drug (Lovell, 1986a,b,c; Ahmed et al., 1989; Flecknell, 1996). Impatience to begin the procedure can be extremely

detrimental, since death from overdosage is often a result of redosing after the animal struggles or vocalizes following the initial dose of anesthetic. Some re?exes persist, even when periods of recumbency are long, prompting questions about the adequacy of anesthesia. Published dosages, such as those in Table A.4B.2, should be used only as a guide. To minimize risks, the anesthetic and its proper dosage should be carefully considered. Since administration of a predetermined amount of injectable anesthetic produces responses varying from inadequate anesthesia to death (Smith, 1993), a single animal should be anesthetized and the response assessed before larger numbers of animals are used. Dosing should begin at the low end of the suggested dose range, and the animal should be monitored. It may be necessary to try various drug dosages in the context of the experiment and the desired effect. Remember, a signi?cant problem with chemical anesthesia is the inability to adjust the animal’s dose, once given. Another signi?cant problem with injectable anesthetics is prolonged recovery times. Many physiological factors (e.g., cardiac output) that are often adversely affected by the drug itself in?uence the pharmacokinetics of chemical anesthesia. For example, several published regimens provide 30 to 45 min of anesthesia, but sleep time may extend up to 180 min. During this time, the animal is susceptible to hypothermia, respiratory failure, or injury from cage mates if recovering in a group. To avoid this problem, consider an anesthetic regimen that allows use of a reversal agent to counteract the anesthetic effect. General considerations common to rodents are the impact of the home cage environment on choice of anesthetic, and the in?uence of concurrent medications on the selection of anesthetic and/or analgesic agents. If barbiturate anesthesia is used, softwood bedding (pine, cedar) or soiled bedding (ammonia) can alter microsomal hepatic enzymes, affecting sleep duration. Other drugs given as part of the experimental study should also be considered. For example, if ketamine is used as part of a cocktail to anesthetize rats for capsaicin injection, adverse reactions, including death, can occur. Route of administration The route of administration of injectable agents largely depends on formulation of the agent and recommendations contained in the package insert. If multiple choices for the route of administration exist, the choice

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is largely based on the species and desired duration of effect. The most common route of administration in rodents is intraperitoneal injection. This is generally easy to do and safe if performed correctly. Correct technique is essential to achieve uniform absorption of the anesthetic agent; inadvertent injection into the liver, spleen, or mesenteric fat results in uneven absorption and uneven anesthesia. If properly administered, predictable periods of surgical anesthesia are achieved; however, tissue redistribution of drugs in overweight animals often makes proper anesthesia dif?cult. Intramuscular injections in rodents are discouraged. Rodents lack suf?cient muscle mass for accurate injection, making the procedure technically demanding. Necrosis and pain are not infrequent outcomes, due to the small muscle mass and acidic pH of many agents, such as ketamine. If this route must be used, the location of the sciatic nerve must be determined precisely, and the volume delivered should be limited to 0.05 to 0.1 ml/site (mice) or 0.1 to 0.3 ml/site (rats) by dividing the full dosage and performing multiple injections bilaterally in the caudal thigh muscles. The paralumbar muscles provide an additional or alternative site for intramuscular delivery. Subcutaneous injection of anesthesia is attractive due to ease of administration and the large space between the skin and subcutaneous tissues. The major drawback is uneven, slow absorption, leading to unpredictable levels of anesthesia. For optimal absorption, drugs should be hypoosmolar (<300 milliosmoles) with a neutral pH. Similar to agents injected intramuscularly, necrosis at the site of entry is a major side effect with highly acidic or basic substances. Inhalant anesthetics The most consistent and reliable anesthetic regimens for rodents involve inhalant anesthesia. Various inhalant anesthetics currently available include halothane, iso?urane, and ether. Methoxy?urane, while popular, is no longer marketed. Ether is strongly discouraged because it is both ?ammable and explosive. Ether is also highly irritating to the respiratory tract, and excessive respiratory secretions can cause potential problems for the animal. Chemicophysical properties of inhalants determine the type of apparatus necessary for their safe administration, the rates of induction and recovery, and the dose needed to maintain surgical anesthesia. Iso?urane and halothane require special equipment that has impeded their popularity. However, inhalation anesthesia is

state-of-the-art, poses fewer risks for the animal, and, for the investigator willing to learn to use the equipment, provides easily controlled anesthesia with remarkably fast induction and recovery times. Induction chambers and anesthetic delivery Induction chambers can be purchased or made (Gwynne and Wallace, 1992) from acrylic or other clear materials. Standard induction chambers have a fresh gas inlet and an outlet for exhaled and waste gases. They are clear for easy visualization of the animal. The rodent is placed inside the chamber for induction, then removed from the chamber with anesthesia maintained by delivery through a face mask. The chamber should be an appropriate size for the animal (25-cm long × 12-cm high × 10-cm wide for a mouse, or 30 × 23 × 12–cm for rats). Smaller chambers allow faster induction times. Both induction chambers and face masks require a calibrated vaporizer for precise control of the concentration of anesthetic gas delivered to the animal. Because oxygen is needed to volatilize the liquid anesthetic, oxygen is delivered to the animal and helps maintain blood oxygen saturation. Some vaporizers are designed to deliver one speci?c agent (e.g., methoxy?urane) and should not be used to deliver agents with greatly differing vapor pressures (e.g., iso?urane), since the concentration delivered may vary substantially. For example, the vapor pressure of methoxy?urane is 23 mmHg at 20? C, compared to 240 mmHg for iso?urane at 20? C. In contrast, the vapor pressure of iso?urane and halothane is similar; therefore, iso?urane can be accurately delivered from a vaporizer calibrated for halothane. It should be noted that vaporizers can be converted for a reasonable cost by some companies (Anesthetic Vaporizer Services). As a general rule, a vaporizer setting of 1 × minimal alveolar concentration (1 × MAC) will produce light anesthesia; 1.5 × MAC will produce a surgical depth of anesthesia; and 2 × MAC will produce deep anesthesia. Inhalant anesthesia potency is similar across species, so once an inhalant anesthetic is chosen, the vaporizer settings remain the same. When using a face mask, dead space, that part of the circuit that remains ?lled with expired gas at the end of expiration and is then reinhaled by the animal, should be minimized by the use of short, straight tubing. The resistance of the anesthetic circuit also in?uences anesthesia of the animal. The effort that the animal must make to move gas in and out of the

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circuit can be minimized by avoiding narrow or sharply angled components of the system. An understanding of the animal’s physiology will also enable smooth, stable anesthesia. Tidal volume is the volume of gas drawn into the respiratory tract with each breath. The minute ventilation is the volume of gas drawn into the respiratory tract in 1 min, so tidal volume × respiratory rate = minute ventilation. Importantly, the minute ventilation does not equal the gas ?ow rate needed to keep the animal anesthetized. The rate of fresh gas ?ow in a system using a face mask should be three times the minute ventilation. If the gas ?ow rate is too low, the animal will inhale room air from around the edges of the mask, diluting the anesthetic concentration. Gas scavenging Because a high ?ow of fresh oxygen is required for either chamber or mask delivery of inhalant anesthetic, adequate suction of waste anesthetic gases is necessary to avoid exposure to personnel. Scavenging systems are absolutely necessary if a downdraft table or fume hood is not available in the laboratory. There are numerous ways to scavenge anesthetic waste gas, including charcoal scavenging canisters that are commercially available from almost any veterinary source, or an active gas evacuation system, such as the Anesco Evacuation System (AES) available from SurgiVet. Monitoring depth of anesthesia Monitoring the animal’s level of anesthesia is critical to successfully estimate the effectiveness of the dosage and route of administration. The rate and depth of respiration are frequently used to assess cardiopulmonary function. Remember that rodents normally maintain minute ventilation by high respiratory rates and low tidal volumes. During anesthesia, both tidal volume and minute ventilation fall; therefore, the animal should be occasionally ventilated to avoid the risk of respiratory acidosis with or without hypoxemia. Noxious stimuli, such as the toe pinch, will elicit a re?ex motor response and quickening of respiration if the level of anesthesia is inadequate. Rodent surgery is usually done in groups, with several animals operated sequentially. Operator fatigue during these prolonged procedures may lead to error, so use of electronic equipment to provide continuous monitoring of physiological variables can be invaluable.

pH Disturbances in systemic pH are common and can be minimized by shortening anesthesia time during which the animal is unconscious and unresponsive to painful stimuli. Reliable pH monitoring is currently available for rats, but not for mice. Use of pulse oximeters Pulse oximeters are available for use in small animals, although the low volume of tissue available for monitoring limits their use in small animals (Alexander et al., 1989; Vender et al., 1995). Several models can be used in animals weighing >200 g (e.g., rats), but are not accurate for mice. Pulse oximeters are valuable for monitoring arterial oxygenation and pulse rate. Oxygen saturation should always exceed 95%. Detection of arterial hypoxemia provides a “common pathway” to detect problems of different origin (hypoventilation, airway obstruction, equipment-related problems). However, pulse oximeters provide no information about CO2 concentration and therefore are of little use in warning of respiratory failure due to CO2 retention. Heart rate Monitoring the heart rate allows early detection of changes associated with cardiovascular system depression or tachycardia during surgical manipulations, indicating an inadequate depth of anesthesia. Use of a pediatric incubator To facilitate speed of recovery, a pediatric incubator (35? C) is helpful, or if an incubator is not available, the rodent’s cage can be placed on a slide warmer or recirculating warm water blanket. In any case, the animal should be maintained in a warm cage environment, checked every 15 min, and turned to assist circulation and to stimulate the animal. Care should be taken to extend the neck to help with breathing. If the animal is placed on bedding, a piece of gauze or cloth should be placed under the animal’s mouth to prevent accidental inhalation of bedding. Physiological interactions Rodents are subject to hypothermia due to their high metabolic rate and high body surface area to body weight ratio. In addition, most anesthetic agents depress the thermoregulatory centers of the brain. The longer the animal is anesthetized, the greater the risk of lowering core body temperature. Stainless steel

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tables, open body cavities, and use of cold ?uids, drugs, solutions, and so on all contribute to rapid loss of body heat. Hypothermia, the most common cause of delayed recovery from anesthesia, delays anesthetic metabolism and urinary excretion of the anesthetic agent. The metabolic rate falls, and the residual injectable anesthesia effectively re-anesthetizes the animal. Unfortunately, mice and rats lack the physiological ability to overcome these events, and the outcome is frequently death. Warm water recirculating blankets, warm ?uid lavage intraoperatively, pocket warmers, and plastic wrap for insulation all provide heat sources necessary for safe maintenance and recovery of the animal. Recirculating warm (35? to 36? C) water blankets are safer than electrical heating pads, because there is no risk of burning the animal. A rectal probe with a thermistor works well for monitoring the core body temperature of rodents. Monitoring body temperature while the animal is anesthetized is critical to avoid hypothermia and secondary complications. It is critical to keep the animal warm until it is fully ambulatory. A common, often fatal, mistake (due to hypothermia) is mistaking sternal recumbency for full recovery and prematurely returning the animal to the animal holding room, which is probably cooler than the surgical room. If individually ventilated cages (IVCs) are used, the forced ventilation at cage level further cools the animal. Surgical procedures, even if minor, result in ?uid loss due to exposure of mucous membranes, open body cavities, and possibly hemorrhage. Fluid therapy should be considered for every surgical procedure. Warmed ?uids can be given subcutaneously and can also contain an analgesic that will aid in post-operative pain management. Investigators should consult with their veterinarian to determine the dosage and frequency of ?uid administration. Post-operative nutrition Mice and rats have high energy needs. Nutritional support is critical upon recovery to avoid hypoglycemia. Placing a drop of undiluted 50% dextrose on the tongue or subcutaneously injecting 3 to 15 ml (depending on the size of the animal) of warmed 5% dextrose solution is bene?cial. Volume de?cits can be corrected by subcutaneous or intraperitoneal injection of warmed lactated Ringer’s solution. Animals recovering from surgery bene?t from additional nutritional supplements such as fruit, Jell-O, and bacon softies (http://www.Bio-Serv.com).

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Evaluating analgesic ef?cacy Analgesic administration has its own set of concerns. Normal physiological behaviors, such as activity, eating and drinking patterns, and weight gain, are parameters frequently used to determine if an animal is experiencing pain and if the chosen analgesic is ef?cacious. Problems arise with this method of pain assessment if the researcher is not familiar with the normal behavior of mice and rats. It is important that investigators become familiar with the normal behavior of their animal model, while realizing that easily recognized signs of pain will rarely be present. Two problems impede species familiarity: (1) rodents are nocturnal (active at night), so observations made during the day may be inaccurate; and (2) rodents, unlike domesticated species, do not respond positively to human contact, often remaining immobile during observation. Food and water intake are frequently depressed when rodents are distressed or in pain, so these are popular parameters to monitor for evidence of pain. Decreased food intake triggers a cascade of events, including dehydration (reduced water intake), hypoglycemia (sluggishness), and hypothermia (insuf?cient caloric intake and minimal locomotor activity). Returning to normal food and water consumption upon administration of an analgesic is a good sign; however, behavioral changes may occur following analgesia alone. Similar to anesthetics, analgesics also evoke a wide variety of responses. It is important that the researcher recognize that some parameters should not be used as evidence of analgesic ef?cacy with certain analgesics. Buprenorphine in?uences normal locomotor behavior in rats (Liles and Flecknell, 1993); therefore, this parameter should not be used as evidence of buprenorphine’s ef?cacy. In contrast, NSAIDs do not in?uence locomotor activity in rats; therefore, this parameter could be used to assess the ef?cacy of an NSAID in pain management. An additional complication is that many analgesics have not been studied in commonly used laboratory animal species, especially mice and rats. Frequently, information from analgesiometric tests (e.g., hot-plate, tail ?inch; see UNIT 8.9) for determination of drug potency in man has been extrapolated to rodents. The assumption that an effective drug dosage in man is suf?cient to alleviate pain in rodents may or may not be accurate. Finally, in addition to considering the appropriate selection and use of both anesthetics and analgesics for an experimental procedure,

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Table A.4B.4 Troubleshooting Guide for Rodent Anesthesia/Analgesia

Problem

Possible cause

Solution Place a drop of atropine on the tongue, provide heat, make sure airway is clear (the mouth may need to be suctioned with the tongue pulled out), and assist in ventilation by gentle rapid compression of thorax between thumb and fore?nger. Give 0.1 mg epinephrine if no response to atropine. Make sure airway is clear, pull tongue out. Place animal in palm of hand and tip its head down and up. Movement of abdominal contents against the diaphragm will compress and expand the chest. Once the animal takes a breath, place a drop of Dopram on tongue and provide 100% oxygen. Stop. If using volatile agents, check the vaporizer. Is it full? Is there a full tank of oxygen? Is the animal completely connected to the system?

Bradycardia, cardiovascular Impending circulatory crisis, failure (blue mucous membranes, hypothermia, pain, pulmonary animal’s limbs are cool to touch) obstruction, blood loss. Pulse oximetry can be used for monitoring. Oxygen saturation is 95%-98%. A reduction of >5% indicates a potential problem; a reduction of ≥10% requires immediate action. Hypoventilation, respiratory arrest Respiratory rate below 40% of the preanesthetic rate signals impending respiratory failure. Check for pulmonary obstruction, depression of respiratory center by anesthetic overdosage.

Hyperventilation

May be due to lightening level of anesthesia

Hypoxia (blue membranes), <85% oxygen saturation

Oxygen tank is empty, animal is If using volatile agents, is oxygen being rebreathing expired gases, obstruction delivered? Turn vaporizer to zero and increase oxygen ?ow rate. If using injectable agents, consider an appropriate antagonist (reversal agent), such as yohimbine or atipamezole for α2 adrenergics or naloxone for opioids. Check if head and neck of animal are extended, open animal’s mouth and pull the tongue forward. Check for copious salivary secretions; if present, suction secretions from oral cavity. Wrap in plastic, place heat lamp over surgical area, use warmed ?uids Ensure an open airway. If the heart has stopped, hold the chest between thumb and ?nger, rapidly compress the area over the heart ?90 times/min. Once respiration and heartbeat have returned, consider a reversal agent if anesthetic overdosage is suspected.

Hypothermia (can happen Change in respiratory pattern quickly), peripheral limbs cool to touch Cardiopulmonary failure Manipulations of the viscera stimulate vagus nerve, resulting in bradycardia; accidental compression of the thorax (e.g., instruments, surgeon resting hand on chest)

it is important to recognize that the degree of pain and distress experienced by the animal will be markedly in?uenced by the expertise of the person performing the procedure(s), including simple handling and restraint. This highlights the ethical obligation of the scientist to ensure that all involved in the research project have received adequate training and have demonstrated competence in the procedures required.

Troubleshooting
Table A.4B.4 lists some problems that may be encountered with the use of anesthetics and analgesics in rodents. Also provided are explanations of the causes of the problems and suggested solutions for avoiding or overcoming them. The most common dif?culties occur from overdosage of anesthesia.
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Acknowledgement
Dr. Lee Chedester, NIAAA, NIH, contributed the idea and design for the homemade version of a face mask that works with a stereotaxic apparatus.

of laparotomy in the rat. Lab. Anim. 32:149161. 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. Ef?cacy of tribromoethanol anesthesia in mice. Lab. Anim. Sci. 43:189-192. Smith, W. 1993. Responses of laboratory animals to some injectable anesthetics. Lab. Anim. 27:3039. 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.

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. 68:368376. 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 ef?cacy 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. 17:1116. 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 modi?ed 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 in?uence of analgesic treatment. Br. Vet. J. 149:515-525. Mouse and Rat Anesthesia and Analgesia Liles, J.H., Flecknell, P.A., Roughan, J., and CruzMadorran, I. 1988. In?uence of oral buprenorphine, oral naltrexone or morphine on the effects

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
http://www.kentscienti?c.com This is an excellent site for ?nding information about equipment mentioned in this unit, as well as ?nding 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.

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http://www.surgivet.com 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. http://www.med-e-cell.com An excellent site for different sources of items such as catheters and infusion pumps. http://www.innovrsch.com/index.html 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. http://www.Bio-Serv.com An excellent resource for customized (and certi?ed) diets (all species). They also specialize in enrichment treats and medicated dosing systems.

Animal Techniques

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Current Protocols in Neuroscience Supplement 42


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