Mouse Anesthesia: The Art and Science

Abstract

There is an art and science to performing mouse anesthesia, which is a significant component to animal research. Frequently, anesthesia is one vital step of many over the course of a research project spanning weeks, months, or beyond. It is critical to perform anesthesia according to the approved research protocol using appropriately handled and administered pharmaceutical-grade compounds whenever possible. Sufficient documentation of the anesthetic event and procedure should also be performed to meet the legal, ethical, and research reproducibility obligations. However, this regulatory and documentation process may lead to the use of a few possibly oversimplified anesthetic protocols used for mouse procedures and anesthesia. Although a frequently used anesthetic protocol may work perfectly for each mouse anesthetized, sometimes unexpected complications will arise, and quick adjustments to the anesthetic depth and support provided will be required. As an old saying goes, anesthesia is 99% boredom and 1% sheer terror. The purpose of this review article is to discuss the science of mouse anesthesia together with the art of applying these anesthetic techniques to provide readers with the knowledge needed for successful anesthetic procedures. The authors include experiences in mouse inhalant and injectable anesthesia, peri-anesthetic monitoring, specific procedures, and treating common complications. This article utilizes key points for easy access of important messages and authors' recommendation based on the authors' clinical experiences.

Keywords: Mus musculus; anesthesia; animal research; animal welfare; refinement.

Figure 1

The pain pathway. A pain pathway graphic summary, including its major components. A nociceptive stimulus (injuries or surgeries) activates nociceptors (transduction). Stimulus information (ie, pain) travels through the nerve fibers such as A-δ and C fibers (transmission) to the spinal cord (modulation). Sufficient pain signals up-regulation, causing stimulus information to travel up the nervous system to the brain where pain perception occurs in conscious animals (perception). Commonly used analgesics are identified at their pain pathway action sites. *, Indicates the analgesic has both a standard and long-acting formulation available. §, Indicates the drug suppresses the perception of pain (due to a surgical anesthesia plane) but does not provide analgesia. Figure created by Janis Atuk-Jones.

Figure 2

Skin tenting. Example of skin tenting. Skin tenting may be performed on an awake or anesthetized mouse. Under normal hydration, the skin should rapidly return back to the normal position. Skin tenting may indicate approximate 8–10% dehydration.

Figure 3

Body condition scoring. Body condition scoring (ie, BCS) uses physical attributes to indirectly assess the health status of an animal. A BCS is based on a scale of 1–5. Different strains/sexes/disease models may inherently have a lower or higher BCS (eg, strains used to study obesity). (Reprinted with permission from AALAS. Ullman-Cullere, MH and CJ Foltz. Comp Med. 49:319–323).

Figure 4

Water circulating heating pad. During anesthesia, thermal support such as with a warm water circulating pad should be provided.

Figure 5

Rodent surgical record. Anesthetic monitoring should be continuously monitored and the parameters recorded at least every 15 minutes.

Figure 6

Translucent sterile surgical drape. (A) Press’nSeal as a cost-effective translucent sterile drape material. (B) Tegaderm as a translucent sterile drape material. The boundaries of the drape are illustrated by the dotted black outline. Use of a translucent surgical drape will facilitate monitoring of mucous membrane color and respiratory function during surgical procedures. Drapes can also provide additional insulation to minimize heat loss.

Figure 7

Rectal thermometer and pulse oximetry. Pulse oximetry and temperature measurements can be tracked using instruments such as the Kent Scientific PhysioSuite unit. Probe for pulse oximeter (black arrow), infrared warming pad (red arrow), and rectal thermometer (green arrow). The temperature probe relays body temperature information to the monitoring system, and the warming pad will adjust the temperature accordingly to a maintain physiologic temperature (blue arrow).

Figure 8

Indirect blood pressure measurement. Blood pressure can be monitored indirectly with equipment, such as with the Kent Scientific CODA Monitor System, which uses a tail-cuff to monitor the indirect blood pressure.

Figure 9

Recovery cage for mice. Example of a recovery set-up for mice after any anesthetic procedure. The cage should be clean with no bedding and transparent to allow for observation of RR, coloration, return of righting reflex, etc. Half of the cage should be placed over a warming device, such as a circulating water blanket to provide heat support during the recovery process. Recovering animals should not be overcrowded or recovered with animals that have regained ambulatory function.

Figure 10

Nutritional supplements for mice. Shown here is an example of 1 brand that provides gelled water and nutrition for mice. These gels provide easy to access nutrition for mice needing supportive care. Additional nutritional supplements, such as EnerCal, though advertised for larger animal species, have been used anecdotally as a nutritional supplement in mice by some researchers.

Figure 11

Mouse CT imaging. (Left) Individual mouse arrangement for CT imaging. The mouse can be secured with surgical tape. Care must be taken to ensure the chest is not too tightly secured, limiting respiration. A heat supply has been arranged under the mouse to prevent hypothermia. (Middle) Individual mouse prepared to enter a MRI scanner. The animal receives inhalant anesthesia via a nose cone at the end of the bed and is monitored via respiration (blue tube) and rectal temperature (black cable). The body temperature is maintained with warm air. (Right) Anesthesia can be delivered by modifying commonplace equipment. Here, slip-tip 60-mL syringes are connected to anesthetic tubing to deliver inhalant anesthesia to mice. Images courtesy of Dr Laura Pisani of Stanford Center for Innovation in in Vivo Imaging and Dr Tim Doyle and Wu Tsai Neuroscience Imaging at Stanford University.

Figure 12

Monitoring during mouse CT imaging. (Left) An example set-up of pulse oximetry equipment for use during mouse anesthesia. (Middle) Arrangement of pulse oximeter sensor attached to mouse. (Right) Arrangement of pulse oximetry and temperature probe to monitor vitals while the mouse is undergoing imaging in a CT machine. Image courtesy of SA Instruments and Dr G. Ronald Morris.

Figure 13

Lidocaine gel for topical use. A small amount of topical lidocaine gel can be used for stereotaxic bar placement to mitigate pain in mice. The gel should be wiped off at the end of the procedure.

Figure 14

Hypothermia anesthesia performed in neonatal mice. Neonate is placed on top of a latex glove in crushed ice with water.

Figure 15

Inhalant anesthesia performed on a neonatal mouse. Snout of the neonate is placed and sealed in a nose cone covered with a latex glove. A circular hole is cut into the latex glove to fit around the snout of the neonate.