Source: Kay Stewart, RVT, RLATG, CMAR; Valerie A. Schroeder, RVT, RLATG. University of Notre Dame, IN
The Guide for the Care and Use of Laboratory Animals ("The Guide") states that pain assessment and alleviation are integral components of the veterinary care of laboratory animals.1 The definition of anesthesia is the loss of feeling or sensation. It is a dynamic event involving changes in anesthetic depth with respect to an animal's metabolism, surgical stimulation, or variations in the external environment.
Precise and constant monitoring of anesthesia is required to safely maintain the depth needed for a procedure. Parameters to be monitored include heart rate, respiratory rate, body temperature, and blood oxygen levels. For mice and rats, none of these parameters are easily monitored due to these animals' small body sizes. Because the heart rate in rodents is so rapid, the stethoscope normally used for auscultation is inadequate for capturing an accurate heart rate. The stethoscope can only be used to detect the presence or absence of a heartbeat. The normal heart rate for a mouse is 328-780 beats per minute, while the regular rate for a rat is 250-600 beats per minute. Respiratory rates in rodents are also elevated above what can be accurately counted using visual methods or during auscultation. The normal respiratory rate for a mouse is 90-220 breaths per minute, and for the rat this value is 66-144 breathes per minute. To accurately ascertain a heart rate and respiratory rate, specialized electronic monitoring equipment is required. Sensors are either surgically implanted into the animal, or placed externally and interact with the monitoring platform onto which the animal is placed.3,4
The most common cause of anesthesia-related deaths in rodents is due to hypothermia. Rodents have a high surface area to body mass ratio. Additionally, an anesthetized animal loses the ability to shiver to maintain body temperature. Thus, body temperature monitoring and supplemented heat, such as a heating pad, are essential during survival surgical procedures. The normal body temperature for a mouse is 96.6-99.7°F (35.8-37.4°C)5 and for a rat is 96.6-99.5°F (35.9-37.5°C).5 Most thermometers were designed for larger animals and modeled after those used for humans. Mercury thermometers have been largely replaced with digital and electronic versions. Although the digital and electronic thermometers have been documented as accurate when used rectally, orally, and in the ear, their size is inappropriate for small rodents. Rectal probes designed specifically for mice and rats are commercially available, and their use is encouraged.
Blood oxygenation levels are used to evaluate adequate oxygen uptake from the lungs resulting in the appropriate concentration of oxygen in a rodent's arterial blood. Monitoring oxygen uptake also indirectly monitors respiration and ventilation, as it reveals if there is adequate inspiration of oxygen and expiration of waste gases. The heart rate is also implicated in the oxygenation of the blood, as a decrease in the heart rate will result in a reduction in oxygen levels, which could cause inadequate perfusion of blood.6
The goal of the anesthetist is to adequately immobilize and alleviate all pain sensations for an animal with the lowest dose or concentration of anesthesia. Properly assessing the depth of anesthesia is required to achieve this goal. There are four stages of anesthesia and four planes within the surgical stage of anesthesia. During stage one, the animal becomes disoriented. In stage two, there is an excitatory phase with an irregular respiration rate, including breath holding in some mouse and rat strains. The righting reflex-that is the ability to roll back over when placed in a dorsal position-is also lost.
Stage three is the surgical stage of anesthesia. In Plane I, the palpebral and swallowing reflexes are absent. Laryngeal and corneal reflexes are lost in Plane II. With Planes I and II, there are no amnesia or analgesic effects; thus, the animal must reach Plane III prior to the beginning of a surgical procedure. Plane III creates paralysis of the intercostal muscles that results in diaphragmatic respiration. Although initially in Plane III there is only partial analgesia, it progresses to complete amnesia and analgesia as the anesthesia level deepens. It is at this level that the animal is fully anesthetized for a surgical procedure. At Plane IV, the animal has been overdosed and can segue quickly into Stage IV.
As the anesthesia level further deepens, there are complications that can result in the death of the animal. In Stage IV there is complete paralysis of both the intercostal muscles and the diaphragm, which causes severe apnea. This results in respiratory arrest, medullary paralysis, vasomotor collapse, and finally death. The pupils dilate, remaining fixed in dilation while the muscles relax.
The proper choice of anesthetics for surgery and other potentially painful procedures must be determined by a veterinarian. This 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.
Anesthetics are available as inhalants or injectables. Surgical anesthesia can be accomplished using a combination of injectable and inhalant anesthetics.2
1. Inhalant anesthesia induction
Inhalant anesthesia includes isoflurane, sevoflurane, and desflurane, with isoflurane being used most commonly. These anesthetics are used more often because, with them, it is easier to control the depth of anesthesia. Induction of anesthesia using inhalation anesthetics can be accomplished with a bell jar or an induction chamber that is fitted to a precision vaporizer.
2. Induction of anesthesia using injectable anesthetics
Injectable anesthetics are primarily a mixture of ketamine and sedatives or muscle relaxers.
The common combinations are: 1) Rodent Cocktail, which consists of ketamine (100 mg/ml), xylazine (20 mg/ml), acepromazine (10 mg/ml), and sterile saline (0.9% NaCl); 2) ketamine/xylazine 2:1, which consists of ketamine (100 mg/ml), xylazine (20 mg/ml), and sterile saline (0.9% NaCl); and 3) ketamine/xylazine Mouse Mix, which consists of ketamine (100 mg/ml), xylazine (20 mg/ml), and sterile saline (0.9% NaCl). When using ketamine/xylazine combo, boosting should only be done with ketamine only, not xylazine, due to the half-lives of these drugs.
The combination of ketamine with sedatives and/or muscle relaxants needs to be prepared as a stock solution from which individual doses can be drawn. The agents must be precisely measured and diluted with sterile saline to ensure that proper doses are administered to the animals. Because ketamine is a controlled substance, the amount used from the bottles must be noted on a "Controlled Drug Log," and the mixtures must have individual "Controlled Substance Logs." When preparing mixtures, add the ketamine slowly to the bottle, as it tends to foam if injected with force. A sterile stoppered 20 ml bottle is used for the mixture. The bottles must be properly labeled with the name of the compounds, the date mixed, the expiration date, the ketamine lot number (as it is a controlled substance), and the suggested dosage. The expiration date may be determined by the date of the ingredient soonest to expire (depends on the rules/guidelines of the facility/state). For accurate recordkeeping of ketamine, both the empty bottle and the filled bottle must be weighed. Then, the weights must be recorded on the label of the mixture and on the individual Controlled Substance Log sheet that is prepared for each bottle. Store ketamine mixtures in a dark, temperature-controlled area to maintain potency.
3. Anesthesia Assessment
Anesthetic depth can be assessed by testing the response to various stimuli. Voluntary movement will result from physical stimuli of the body. See Table 1 for a list of physical methods utilized for anesthetic depth assessment.
| Method | Procedure | Response |
| Toe pinch | Extend the leg and isolate the webbing between the toes. This area is firmly pinched using either the fingernails or atraumatic forceps. | A positive reflex is indicated by the retraction of the leg or withdrawing of the foot. The animal is not at a surgical plane of anesthesia if there is leg or body movement, vocalization, or marked increase in respirations. |
| Tail pinch | The tail tip is pinched using either the fingers or atraumatic forceps. | A positive reaction is indicated by twitching or movement of the tail. The animal is not at a surgical plane of anesthesia if there is movement of the tail, vocalization, or marked increase in respirations. |
| Ear pinch | Using the fingers or atraumatic forceps, pinch the tip of the pinna. | A positive reaction is shaking the head or the movement of the whiskers forward. If there is movement of the head, whiskers, vocalization, or marked increase in respirations, the animal is not at a surgical plane of anesthesia. |
| Palpebral reflex | Using a fingertip, touch the medial canthus (inner corner) of the eye. | A positive reflex is indicated by a blink in response to touching the eyelids. If there is movement of the eyelids, whiskers, or marked increase in respirations, the animal is not at a surgical plane of anesthesia. |
| Corneal reflex | Using a cotton-tipped applicator, gently touch the cornea (eyeball). | A positive response is indicated by a blink. If there is movement of the eyelids, whiskers, or marked increase in respirations, the animal is not at a sufficiently deep plane of surgical anesthesia. |
Table 1. Physical stimuli methods for assessing anesthetic depth.2
Physiological indicators such as heart rate, respiratory rate, blood pressure, mucous membrane color, and capillary refill time should also be used. While general observations can be useful to detect changes in the respiratory rate of the animals, to utilize the heart rate, or blood pressure for depth assessment, specialized equipment is required. If an electrocardiograph is available, the rate and the strength of the heartbeats can be measured. For measuring the blood pressure, there are a variety of devices that are fitted over the tail or even over the entire body. Physical stimuli as described in Table 1 will cause an increase in all three of these parameters.
The color of the mucous membranes, eyes, ears, mouth, nose, anus, and-to a lesser extent-the paws and tail are observed for changes. The areas should be pink, indicating adequate respiration and cardiac function. When the animal moves to Stage IV anesthesia, the respirations cease, resulting in cyanosis-indicated by a blue or gray color-to the mucous membranes and surrounding skin.
Capillary refill time is defined as the amount of time taken for color to return to an external capillary bed after it has been blanched by the application of pressure over the area. An applicator stick or a finger is pressed on the gums, pinna, or nail beds of the anesthetized animals. The number of seconds that it takes for the blanched area to return to a pink color should not be more than 1-2 seconds. An extended refill time suggests a reduction in heart rate or strength of cardiac contractions, indicating the animal may be too deeply anesthetized and near death.
It is important to utilize several different parameters to assess 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. Use alternate sites for toe and ear pinch assessments. Anesthetic depth should be reassessed every 10-30 minutes throughout the surgery.2
Studies have shown that there are cardiorespiratory changes in an anesthetized animal. While anesthetized with injectable drugs, the animals experience a stable respiratory rate; however, they demonstrate variability in cardiac output. The response to injectable anesthetics has been reported to vary greatly between different strains, thus it is difficult to standardize the dosage.7 Inhalant agents tend to decrease the respiratory rate but have a lesser impact on the cardiovascular system. As the dosage of inhalant anesthesia is easily adjusted throughout the duration of the procedure, it is often the preferred method.
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!
The proper use of anesthetics for surgery, or other potentially painful procedures, is crucial not only for the animal's wellbeing, but also for the integrity of the scientific data collected during the procedure. There are many variables that factor into choosing the appropriate anesthetic regiment. The depth of anesthesia must be closely monitored, as each individual animal can respond differently to the drug. With the use of the proper anesthetic and careful monitoring, painful procedures can be accomplished with no pain and minimal physiological changes in the animal.
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!
