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Rodent Anesthesia

Rodent Anesthesia Wetlab
26th Annual Conference and Trade Show
Saskatchewan Association of Veterinary Technologists Inc.
November 5-7, 2010; Saskatoon

Dr Colette Wheler, DVM
Ms Peggy Nelles, RVT RMLAT
Ms Nadine Schueller, RVT
Ms Carla Hudy, RVT

The goal of this wetlab is to provide the basic skills necessary to safely anesthetize a rat or a mouse. At the completion of this session, participants should be able to:

1. Perform a physical examination on a rat or mouse, including weight
2. Develop an injectable anesthetic protocol, including pre-medication and post-operative analgesia for a rat and mouse, and calculate correct drug dosages
3. Carry out an injectable anesthetic on a rat
4. Provide appropriate support to the animal while it is anesthetized
5. Monitor reflexes and vital signs of the anesthetized animal
6. Provide an appropriate recovery environment for the animal
7. Develop a plan for appropriate post-operative care
   
   In addition, inhalational anesthesia will be demonstrated on a rat and a mouse.

 GENERAL INFORMATION

Anesthesia:

• a type of chemical restraint used for medical and surgical procedures
• allows us to safely perform a variety of manipulations and procedures on animals without causing them unnecessary pain or distress.
• a state of controllable, reversible insensibility in which sensory perception and motor responses are markedly depressed.
• causes CNS (central nervous system) depression
• affects the cardiovascular, respiratory and thermoregulatory mechanisms

Support of these body functions is necessary during the anesthetic episode, and forms part of the overall anesthetic management of the patient.
 

PRE-OPERATIVE CARE

• Rats and mice are unable to vomit so it is not necessary to withhold food and water from them prior to anesthesia.
• The animal should always be handled gently and calmly in order to minimize struggling and fright. Prolonged excitement will disturb the circulatory and metabolic state of the animal. This will increase the risk of anesthetic complications.
• Ensure all drugs and equipment are readily available for the anesthetic procedure.
• Animals should be examined just prior to anesthesia for any signs of breathing or heart problems, including:
  • Respiration rate and character
  • Heart rate
  • Mucous membrane (gum) color
  • Discharge from the eyes or nose
  • Hydration status (Checked by pinching up the skin)

Normal Physiological Parameters for Rats and Mice
Species	Rectal Temperature	Respiratory Rate	Heart Rate
Rat	37.0 °C	70-115 breaths/min	250-450 beats/min
Mouse	37.5 °C	94-163 breaths/min	325-780 beats/min

Assessment of Hydration Status

A determination of hydration status can be made by performing a “skin tent”, where the elasticityof the skin is evaluated. The skin over the nape of the neck is gently pinched together and pulledaway from the body. In a normal animal, the skin will immediately fall back into its normalposition. In a severely dehydrated animal, the skin will remain tented up, as it has lost all of its elasticity due to dehydration.

 

% Dehydration	Detectable Signs
<5%	Not detectable
5%	Subtle loss of skin elasticity
6-8%	Definite delay in return of skin to normal position
10-12%	Tented skin stands in place, eyes are sunken
12-15%	Signs of collapse and severe depression; on the verge of death

Weight: Determination of the weight of the animal is one of the most important things to do!! The weight of the animal is used to calculate drug dosages, and is also used for postoperative monitoring of the animal, to ensure it is eating and drinking after surgery.

PRE-ANESTHETIC MEDICATION
Depending on the situation, some drugs, such as analgesics, anti-cholinergics and sedatives may be given before the anesthetic agent. The selection of a “pre-med” depends on the species of animal, the other anesthetic agents to be used, and personal preference. Analgesics are the most common type of pre-med given to rats and mice.

GENERAL ANESTHESIA

General anesthesia causes:

• loss of consciousness
• suppression of reflexes
• muscle relaxation
• analgesia (suppression of pain perception)

It is important to remember that the amount of pain relief (analgesia) will vary depending on the specific anesthetic agent used. The entire anesthetic protocol (ie pre-med, general anesthetic and post-operative care) must provide an appropriate level of analgesia for the procedure being performed. If anesthesia is being used to provide humane restraint for non-painful procedures, such as radiology, then only light anesthesia with minimal analgesia will be needed. However, if a painful surgical procedure, such as a fracture repair, is being performed, complete suppression of pain perception is needed. This will involve pre-medication with analgesics, a deep level of anesthesia, and administration of post-operative analgesics, often for several days afterwards.
 

In rodents, general anesthetics can be given by injection or inhalation. There are a variety of anesthetics available, each with their own advantages and disadvantages.

General Anesthesia – Injectable

• injectable anesthesia requires very simple equipment
• drugs are usually given intraperitoneally (IP) in rodents
• the appropriate size of syringe and needle is important for accurate dosing and patient comfort; 1cc insulin syringes and 25 or 27 g needles are recommended
• the dosage of anesthetic must be carefully calculated to avoid an overdose
• a fresh, sterile needle and syringe should be always be used to draw up the anesthetic in order to avoid bacterial contamination of the drug; needles must be disposed of in the appropriate “sharps” container, and NEVER discarded in the garbage
• the needle should be protected from contamination at all times, ie do not touch it or lay it down on a non-sterile surface
• in rats and mice, the needle should be introduced at an angle of about 30o, slightly to the right of midline, about one third to one half way between the hind legs and the end of the ribcage
• the plunger should be gently pulled back prior to injection to ensure the needle has not accidentally entered a blood vessel, or an abdominal organ
• if blood, urine or other material enters the syringe when the plunger is pulled back, the
  needle should be withdrawn and the syringe, needle and contents discarded. The injection can be tried again with a new needle and syringe.

IP injection in a mouse	IP injection in a rat

• complications may include:
  • injection of the drug into the intestines
  • injection of the drug into the urinary bladder
  • injection of drug subcutaneously or intramucularly
  • injection into abdominal organs, such as liver or spleen
  • laceration of a blood vessel or organ resulting in hemorrhage
• the rate of drug absorption, and the anesthetic effect may vary considerably; the anesthetic response will also vary according to the animal’s strain, sex, and age
• one disadvantage of administering drugs by the IP route as a single dose is that it is impossible to adjust the dose according to the individual animal’s response, so inadvertent overdosing and underdosing may occur
• it is also possible to inject anesthetic agents intravenously into the tail vein, however a greater degree of skill is necessary
• the intramuscular (IM) route is not recommended in rodents due to problems with tissue damage and self-mutilation

Once the anesthetic is injected, the animal should be placed by itself in an empty cage lined with paper towel and watched closely until it becomes anesthetized. Noises and other stimulation should be kept to a minimum to facilitate a smooth induction of anesthesia.

Injectable Anesthetic Drugs
A variety of injectable anesthetics and anesthetic combinations may be used to anesthetize the animal. Factors to consider when choosing which drugs to use include:

• species and strain of animal
• affect of the drug(s) on the animal
• length and invasiveness of the procedure
• availability of the drugs
• personal preference

A list of injectable anesthetics and dosages for rats and mice can be found in Appendix 1

General Anesthesia - Inhalational

• volatile drug inhaled and absorbed through lungs
• can provide anesthesia for as long as needed
• need specially designed anesthetic equipment
• can ventilate the drug out of the body with O2
• best method for long procedures
• an anesthetic machine is necessary to delivery oxygen and volatile anesthetic agent
• an anesthetic machine has four basic components:
  • A compressed oxygen source
  • A regulator to ensure a constant pressure of gas flow
  • Flow meters to control the flow of gas
  • A vaporizer, which is a device for turning the volatile anesthetic agent into a vapour and delivering an accurate concentration to the patient

The anesthetic system (the part that attaches to the anesthetic machine) depends somewhat on the size of the animal and the type of equipment used. All anesthetic breathing systems must deliver enough anesthetic gas and oxygen to meet the animal’s ventilatory requirements and to remove exhaled gases, which will contain carbon dioxide. If these expired gases are not removed, they are breathed in again by the animal, but may not contain enough oxygen to support the animal’s life.

Most rodents are induced with either injectable drugs or in a chamber. Commercially available anesthetic chambers for mice and rats are available or a chamber can be made out of glass or plexiglass. The anesthetic gases are delivered into the bottom of the chamber, and the waste gases are removed from the top of the chamber. For rapid induction, the entire chamber should be quickly filled with anesthetic.

Anesthetic Chamber for Rodents

Once the animal is induced it is removed and a facemask is placed over its nose and mouth, which is an open breathing system. Expired gases pass around the edges of the mask, and, as long as the oxygen flow rate is high enough, very little rebreathing of these gases occurs. If the oxygen flow is too low, the animal will breath in room air from around the edges of the mask, which will dilute the anesthetic and may cause the animal to become too lightly anesthetized. The most serious disadvantage with a face mask is that it is not possible to ventilate the animal artificially, should the animal stop breathing. For human safety, exhaled gases should be scavenged and removed from the operating area.

Rodent Ansthesia Using a Face Mask (open breathing system)

A Modified Bain System is commonly attached to the face mask and used for rodents. A rebreathe bag acts as a reservoir for gases. Other systems are also available depending on the species of animal being anesthetized.

Modified Bain System

It is also possible to place an endotracheal tube into both rats and mice, however, this requires a great deal of skill and some specialized equipment. The benefit of placing an endotracheal tube is that the anesthetic is delivered directly into the lungs, so less is needed; the animal’s lungs can be directly inflated, should the animal stop breathing; and the airway is protected from fluids.
 

Summary of Inhalational Anesthetic Technique

 

IP Injection	OR        Gas Induction in Chamber

Induction:

The animal is induced either by injecting anesthetic intraperitoneally, or by using gas anesthetic
in a chamber.

Maintenance:
Once the animal is induced:

• it is placed on a heated surface
• lubricant is put in the eyes
• a face mask is placed over its nose and mouth
• the anesthetic gas and oxygen are turned on to maintain anesthesia (a modified Bain system is often used for rodents)
• reflexes are checked to ensure the animal is anesthetized deeply enough for the procedure
• vital signs are monitored for the duration of anesthesia.

Drugs Used
Isoflurane is the most common inhalational anesthetic used in rats and mice.

Isoflurane

Advantages:

• Rapid induction and recovery
• Depth of anesthesia can be altered quickly
• Non-irritant, non-explosive, non-flammable
• Very little biotransformation; almost completed eliminated in exhaled air
• Very little effect on liver microsomal enzymes

Disadvantages:
 

• Slightly more respiratory depression than halothane but less CV depression
• Pungent odor
• No residual analgesia upon recovery

Stages of Anesthesia
The following signs are associated with various levels of anaesthesia:

Sedation	Tail down, head down, locomotion slows and stops, respiration slows but breathing remains regular
Light Anesthesia	Righting reflex lost, responds markedly to painful stimuli
Surgical Anesthesia	Withdrawal reflex and tail pinch response absent, limb muscle tone decreased/absent, heart and respiratory rates slower and regular
Overdose	Deep irregular breaths, cyanosis (blue color) on footpads, ears, tail, pupils fail to respond to light, slow irregular heart beat

Even if injectable anesthesia is used, it is a good idea to supply the animal with oxygen to provide support to the respiratory system during anesthestia.

Anesthesia is not a natural state, ie the animal is not simply sleeping, and everyone must be aware of this. The amount of risk involved depends upon the health of the animal, the experience of the surgeon/anesthetist and the facilities available.
 

MONITORING

• the animal must be monitored to ensure that sufficient analgesia and unconsciousness has been produced for the procedure to be performed – is the animal deep enough?
• because anesthetic agents commonly affect the cardiovascular, respiratory, and thermoregulatory systems, vital signs must be monitored to detect and correct any problems that might occur – is the animal experiencing an adverse reaction to the anesthetic

It is important to try to maintain heart rate, respiratory rate and body temperature within normal physiological limits during anesthesia.

Parameters Used to Monitor Anesthetic Depth in Rodents

• Depth and rate of respiration
  • watch for expansion of the chest cavity
  • watch for secretions at the back of the throat (a gurgling sound) – it may be necessary to suction or swab the throat to clear the airway
• Heart rate
  • gently feeling the sides of the chest will give an indication of heart rate
  • it will be impossible to count the heart beats, but you can get an idea of rate and regularity
  • a stethoscope, ECG, Doppler or pulse oximeter can be used to monitor heart rate, if available 
• Mucous membrane color
  • Gums should remain pink; a bluish color (cyanosis) indicates hypoxia (oxygen levels are too low)
  • Capillary refill time is useful indicator of blood pressure. This is performed by pressing on the gums to blanch them, then watching to see how quickly the blood refills the tissues, which normally occurs in less than 2 seconds
• Body Temperature
  
  • Anesthesia diminishes the body’s ability to thermoregulate
  • Rodents will chill very quickly due to their small body mass
  • The most common cause of death in anesthetized rodents is hypothermia
  • Hypothermic animals will take longer to recover from anesthesia
  • Thermal support must be provided to maintain normothermia
  • Options include;
    • circulating warm water blanket – safest method to avoid burns
    • electrical heating pad – ensure adequate insulation by placing a drape folded several times between the animal and the heating pad to help avoid accidental burns
    • loosely wrap the animal in bubble wrap
    • heat lamp – avoid placing it too close to the animal to avoid burns and always make sure a portion of the cage is shielded from the heat lamp to allow the animal to have a cooler place to move to in order to avoid hyperthermia
    • place padding between the animal and the work surface (newspaper, paper towel, cloth towel, fleece)
    • hot water bottle or equivalent (ensure the water is not too hot or burns may occur!!!)
  • It can be difficult to take the temperature of a rodent; small rectal thermometers may be used in rats; fine rectal probes, implants in mice  
• Reflexes – used to measure the degree of central nervous system depression
  
  • Pedal reflex (digital withdrawal)
    • performed by extending a limb and using your fingers to pinch the web between the toes (not the toe itself)
    • if the limb is withdrawn, muscles twitch or the animal cries out, the animal is not sufficiently anesthetized
    • lack of response indicates deep anesthesia
    • works best with barbiturate anesthesia; not as effective with inhalational anesthetics (disappears at a lighter plane of anesthesia)
  • Tail pinch: used in small rodents only when the foot is too small to perform the
    pedal reflex
  • Palpebral reflex (blink reflex):
    • performed by lightly tapping the skin at the medial canthus of the eye, or
      running the finger along the eyelashes
    • if the animal blinks or the lid moves, the animal is not sufficiently
      anesthetized
    • this reflex disappears in light to medium anesthesia
    • this reflex cannot be used when ketamine has been administered, as ketamine inhibits the blink reflex in animals before they reach a surgical plane of anesthesia
  • Anal reflex
    • contraction of the anal sphincter muscle on sudden manipulation of the anus
    • disappears with medium to deep anesthesia
• Other
  • Lacrimation (tear production)
    • lacrimation is reduced in the deeper stages of surgical anesthesia, which
    • leads to drying of the cornea and possible keratitis or ulceration
    • **ophthalmic ointment or sterile mineral oil must be instilled into the
    • conjunctival sac to prevent drying and damage to the cornea**
  • Muscle relaxation
    • ocurs with most anesthetics, except ketamine, which causes rigidity
    • jaw tone can be used to monitor anesthetic depth – it should be abolished
    • at the surgical plane of anesthesia with halothane and isoflurane

A variety of instruments are available that can be used to monitor an animal’s vital signs while under anesthetic, such as blood pressure monitors, EKG machines, pulse oximeters, dopplers, etc. However, most investigators do not have the necessary equipment for use in rodents.

RECOVERY

• recovery refers to the period from the end of the procedure performed under anesthetic until the animal is
  • conscious
  • can maintain normal body position unassisted
  • is unhooked from all monitoring devices
  • has regained normal physiological functions (eg. drinking, urinating, eating, defecating)
• the time frame for recovery may be as little as a fraction of an hour to several hours, to more than 24 hours, depending on the length of the anesthesia and the invasiveness of the surgical procedure
• continuous monitoring should be provided during the immediate post-anesthetic period
• if the animal(s) are not completely recovered by the end of the normal working day, arrangements must be made for regular checks by qualified people during the evening and night for this reason, it is a good idea to schedule surgeries early in the day, so that there is sufficient time to monitor the animals postoperatively
• each animal should be held in a separate cage until allowed to recover
• under no circumstances should a partially recovered animal be placed back into a cage with other animals as cannibalism may occur
• the animal should be placed in a body postion to allow for normal breathing
• rodents should not be placed directly on particulate bedding that could obstruct the nares; paper or cloth based bedding that is absorbent and warm should be used
• care must also be taken to avoid placing the rodent so that the nose is up against the side of the cage
• thermal support must be continued during the recovery period; animals wake up more quickly if their body temperature is maintained in the normal range
• post-operative/post-anesthetic heat supplementation must be maintained until the animal is fully recovered from the effects of the anesthetic, which may be several hours after the return of the righting reflex

MAINTENANCE OF HYDRATION

• surgery may result in excessive fluid loss
• on recovery, the animal may not drink for some time after recovery
• due to their small body size, rodents are prone to dehydration in the post-anesthetic period
• fluids, warmed to body temperature, should always be given to replace estimated losses
• the fluid requirement during surgery is approximately 10 ml/kg/hr
• fluids may be given subcutaneously or intraperitoneally while they are still anesthetized; oral fluids may be given if the animal is conscious
• monitoring the animal’s body weight before and after surgery can provide a good estimate of loss of body fluids

POST-OPERATIVE MONITORING AND ANALGESIA

• careful monitoring of the animal must occur during the immediate post-operative period and for several days after
• the weight and body temperature of the animal provide good, objective data that is very useful
• other observations, such as whether the animal is hunched over, huddled in a corner, depressed, not eating or drinking, licking at the surgery site, etc, are important to determine how well the animal is recovering from the surgery, and for how long to continue analgesics
• fluid therapy and supplemental feeding may be necessary to help the animal recover and heal
• analgesics should be given at the recommended dose and frequency during the immediate postoperative period
• the animal should be closely observed for changes in physiology and/or behaviour which might indicate that pain is present

Clinical Signs of Pain in Rodents

 

Clinical Sign	Example
Impaired activity	Soiling of animal due to lack of grooming; reluctance to move; limping; sitting in corner of cage away from others
Changes in temperament	More or less docile or aggressive
Restlessness	Shaking or scratching a particular area; hyperactive or circling behaviour, skin twitching
Decreased intake	Food and/or water
Abnormal posture	Hunched back (abdominal pain); standing with front legs apart (thoracic pain)
Self-mutilation	Excessive licking or chewing of an appendage
Changes in bowel or urinary activity	Constipation; impaction; increased or decreased urination; incontinence
General appearance	Rough hair coat; dull eyes

GOOD NURSING CARE IS VERY IMPORTANT FOR POST-OPERATIVE SURVIVAL!!!!!!!!

 

Appendix 1
Some Drugs Used for Anesthesia/Analgesia of Rats and Mice
(IP = intraperitoneal; SC = subcutaneous; PO = per os (by mouth); IV = intravenous)

  

Last Updated: November 28, 2010