Login processing...

Trial ends in Request Full Access Tell Your Colleague About Jove
JoVE Science Education Library
Lab Animal Research
 

Anesthesia Induction and Maintenance

Article

Transcript

Anesthesia induction and maintenance forms an integral component of veterinary care of laboratory animals undergoing any form of surgical procedure. The goal of anesthesia is to adequately immobilize the animal and alleviate all pain sensations. In addition to induction, precise and constant monitoring is required to safely maintain the correct anesthetic depth throughout the procedure.

In this video, we will first briefly discuss the levels of rodent anesthesia and what stage one should aim to achieve. Next, we will review the different induction and maintenance methods, various ways to ensure that the animal is always in the desired anesthetic stage, and finally a few real-world experiments involving use of different anesthetics for varied purposes.

Let's start by discussing the levels. There are four stages of anesthesia and four planes within the stage three or the surgical stage.

During stage one, the animal becomes disoriented. Stage two is marked by an irregular respiratory rate and loss of the righting reflex. In plane one of stage three, the palpebral and swallowing reflexes are absent. During the plane two, the laryngeal and corneal reflexes are lost.Up until this point, the anesthetic has not induced amnesia or analgesia.

It is in plane three that amnesia and analgesia progresses from partial to complete, and the animal is fully anesthetized for a surgical procedure. Plane three is also signified by paralysis of the intercostal muscles, which results in diaphragmatic respiration that is shallow breathing. In plane four, the animal has been overdosed and can proceed quickly into stage four, where there is complete paralysis of both intercostal muscles and diaphragm, which can cause respiratory arrest, and ultimately lead to death.

Anesthetics are available as an inhalant or injectable, and a veterinarian must decide what to use for the procedure to be performed. This choice is based on numerous aspects including: the extent and duration of the procedure, the species and strain, the age, and the physiological status of the animal.

The class of commonly used inhalant anesthetics includes compounds like Isoflurane, Sevoflurane, and Desflurane. These compounds allow for easy control of the anesthesia depth. There are a few options in equipment that one can choose from to administer inhalant anesthetics.

One of the choices is a bell jar, which should be used under the hood -- and not on the bench -- to avoid personnel exposure to the anesthetic gases. Assemble the jar with a ceramic or plastic perforated platform creating a space between the bottom of the jar and the platform. Next, while wearing impervious gloves, saturate a cotton ball with anesthetic and place it under the platform so that it rests at the bottom of the jar. Then immediately secure the lid to prevent escape of the anesthetic vapor. To place the animal, slide the lid to one side, introduce the animal and secure it immediately. Following that, observe the activity and respirations to determine the depth of anesthesia, and expose the animal to the inhalant to effect. Note that the platform serves as a barrier and prevents the animal from coming into direct contact with the liquid anesthetic.

An alternative to the bell jar is an induction chamber used in conjunction with a precision vaporizer machine connected to an oxygen tank. The first step is to ensure that the vaporizer is filled with appropriate amount of the liquid anesthetic. Next, check the waste gas scavenging system. If it is the commonly used passive system, then weigh the canister to determine if it is still effective. Generally an increase of fifty grams above the starting weight is the point at which the canister is spent. The next step is to assemble the induction chamber. Ensure that the input is from the vaporizer and output is to the waste gas scavenging system.

To start, place the animal into the induction chamber and secure the lid. Once the animal is in the chamber, first start the oxygen flow at the rate of 1 liter per min, and then adjust the precision vaporizer setting to an induction level of 3-4 % for isoflurane. Like bell jar, expose the animal to the anesthetic to effect. Once the animal is fully anesthetized, flush the chamber with oxygen by turning the isoflurane off before gently removing the animal. This is to prevent personnel exposure to anesthetic gases.

Another method for anesthesia induction is via nose cone or facemask also connected to the precision vaporizer. However, because anesthetic gases have an unpleasant smell, animals may object to being masked for induction. In addition, there is also a risk of causing asphyxiation because of grasping too firmly. Therefore, the preferred method is to use the induction box or the bell jar to induce anesthesia followed by maintenance with the nose cone. More often than not, the assembly is such that the cone and the induction chamber both are connected to the same vaporizer with a toggle in between to switch the anesthetic vapor delivery from the induction chamber to the nose cone and vice versa. After anesthetizing the animal in the chamber, secure its face in the cone, and switch the toggle on the tubing to redirect the gas flow to the nose cone. Monitor the respiration and after confirming that the animal is relaxed, reduce the anesthetic to a maintenance level of 0.5 - 1.5 %. Also, apply ophthalmic ointment to the eyes to prevent corneal drying.

For injectable anesthetics, a mixture of Ketamine and other sedatives or muscle relaxers including Xylazine and/or Acepromazine. Different combinations can be prepared using these compounds. See text below for commonly used ratios. Note that Ketamine is a controlled substance and therefore the amount used must be noted on the Controlled Drug Log and the mixtures must have their individual Controlled Substance Logs. Depending on the species, age and health status of the animal, the mixture and the dose of the anesthetic are selected and the solution may be injected intraperitoneally or intramuscularly. Usually, injection and inhalation anesthetics are used in combination to achieve surgical anesthesia.

Now that you know how to induce anesthesia, let's learn about anesthetic depth assessment, which is important to monitor every 10-30 minutes to ensure that the animal is not harmed during the procedure. There are several methods for doing so in rodents.

A commonly used method is the toe pinch. Extend the animal's leg and isolate the webbing between the toes. Then firmly pinch the area using either the fingernails or atraumatic forceps. A positive reflex is indicated by the retraction of the leg or withdrawing of the foot. Another method is the tail pinch performed at the tip of the tail. A positive reaction is demonstrated by twitching or tail movement. You can also pinch the tip of the pinna, and if there is shaking of the head or the movement of the whiskers forward, then the animal is not in the surgical plane of anesthesia.

To check the anesthesia depth, one can also touch the medial canthus or the inner corner of the eye to elicit the palpebral reflex -- indicated by a blink in response to touching of the eyelids. Even if there is movement of the eyelids, whiskers, or marked increase in respirations the animal is not in the surgical plane of anesthesia.

Lastly, one can check the corneal reflex by touching the cornea with gloved finger or a cotton-tipped applicator. A positive response is indicated by a blink.

It is important to alternate between sites to assess the anesthetic depth. Using the same toe or ear for repeated pinches will desensitize the area and the response will be repressed and not give an accurate assessment of anesthetic depth.

In addition to these physical stimuli methods of assessment, one should also monitor the physiological indicators including the heart rate, respiratory rate, blood pressure, mucous membrane color, and capillary refill time. While general observations can be useful to detect changes in the respiratory rate, to utilize the heart rate for depth assessment, specialized equipment like electrocardiograph may be used. For measuring the blood pressure, there are a variety of devices that can be fitted over the tail or even over the entire body. The color of mucous membranes, eyes, ears, mouth, nose, anus, paws, and tail can also indicate anesthetic depth. These areas should be pink, suggesting adequate respiration and heart rate.

To check the capillary refill time, press on the pinna of the anesthetized animals, and count the number of seconds that it takes for the blanched area to return to a pink color. This should not be more than 1 to 2 seconds. An extended refill time suggests a reduction in heart rate or strength of cardiac contraction, indicating the animal may be too deeply anesthetized and near death. After removing the animal from anesthesia, they should not be returned to the housing facility until recovered from anesthesia, unless they are continuously monitored in the housing area.

Now that we've learned the principles and procedures of rodent anesthesia induction and maintenance, lets look at some of the frequent applications of anesthetics in biomedical research today.

Probably the most common use for rodent anesthesia is prior to and during surgery. For example, here researchers wanted to develop a model of stroke caused by clot formation in brain. In order to achieve that, they induced anesthesia in mice and then drilled the cranium to create a thin window. And while the animal was still sedated, these scientists injected a photosensitive dye into the circulation. Next, they induced photoactivation with the help of a laser through the drilled cranium to cause formation of a clot in the cranial vasculature.

Another instance in which rodent anesthesia is required is for performing physiological analysis. For example, scientists often use ECG electrodes on anesthetized animals to monitor heart activity. Or they use ultrasound probes to determine the rate of diaphragm movement to more accurately quantify the respiratory rate.

Lastly, use of anesthesia is mandatory when preforming survival in utero experiments. For example, in utero electroporation -- a method in which a pregnant female is anesthetized, an incision is made to expose the developing embryos, and electrodes are used to induce embryonic cellular uptake of the injected genetic material.

You have just watched JoVE's video on anesthesia administration and maintenance. Since rodent anesthesia facilitates the execution of such a wide range of biological experiments, it is imperative that every scientist possesses the skill of inducing and maintaining the correct anesthetic depth throughout an experiment. As always, thanks for watching!

Read Article

Get cutting-edge science videos from JoVE sent straight to your inbox every month.

Waiting X
simple hit counter