Veterinary Clinical Services

Anesthesia and Analgesia

Recommended Drugs and Dosage for each Species

Anesthesia Guidelines

Legal Requirements
General Considerations

What is Anesthesia?
Monitoring and Maintenance of Normal Physiology

Respiratory Support
Fluid Therapy/Cardiovascular Support
Thermoregulation
Monitoring

Muscle Relaxants

A. Legal Requirements

Federal Guidelines

The Federal Animal Welfare Act (Public Laws 89-544, 91-579 and 98-198) require that guidelines and consultation be provided to research personnel on the type and amount of anesthetics, analgesics and tranquilizers for each species of animal used. Proper precautions, records and handling of controlled substances are the responsibility of the principal investigator.

B. General Considerations

1. What is Anesthesia?

Anesthesia is a state of unconsciousness induced in an animal. The three components of anesthesia are analgesia (pain relief), amnesia (loss of memory) and immobilization. The drugs used to achieve anesthesia usually have varying effects in each of these areas. Some drugs may be used individually to achieve all three. Others have only analgesic or sedative properties and may be used individually for these purposes or in combination with other drugs to achieve full anesthesia.

It is important to realize that anesthesia is not a simple thing. It has profound effects on an animal's physiology because of the generalized central nervous system effects as well as specific effects on all other body systems. Thus, while anesthesia is required by law to prevent pain or distress in research animals, it must not be ventured into lightly. It is important to learn about the drugs you will be using and about the physiology of the animal you will be monitoring.

2. Monitoring and Maintenance of Normal Physiology

a. Respiratory Support

Most anesthetics cause direct depression of the respiratory center in the brain and reduce ventilation. This is complicated by other factors that may interfere with respiration. When an animal is in lateral recumbency the lung that is down is being compressed by the rest of the body. Likewise, animals in dorsal recumbency may experience compression of the diaphragm by abdominal viscera. The airway may be compromised by regurgitated food or pharyngeal and tracheal secretions that normally would be removed by reflex swallowing or coughing. These reflexes are lost during anesthesia. There are several ways to monitor and support the ventilation of an anesthetized animal.

Withhold food and water from large animals for 12 h prior to anesthesia and from small animals for 2 h to prevent regurgitation. It is not necessary to withhold food and water from rodents or rabbits prior to anesthesia. Prolonged food or water deprivation are distressful to animals and are rarely necessary.
Premedication with atropine or glycopyrrolate (anticholinergics) may reduce the respiratory tract secretions in some animals
Intubate the trachea whenever possible, even if injectable anesthetics are being used. Intubation can be achieved on animals as small as a rat. This will prevent aspiration pneumonia and allow you to assist respiration if the animal stops breathing.
Assist respiration during the procedure. This can be done with a mechanical ventilator. However, mechanical ventilation is rarely needed (unless a thoracotomy or diaphragmectomy is being performed) and can be detrimental to the animal if over-done. Attaching an AMBU bag to the endotracheal tube or using an anesthetic machine's rebreathing bag will allow you to administer a deep breath every 2-5 min during the procedure. This will inflate all areas of the lungs and improve gas exchange. If the animal is not intubated, ventilation can be performed using a nose cone or face mask (may cause overinflation of stomach).
Protect the eyes from drying out using an ophthalmic ointment and protect them from being contaminated with surgical scrub solutions. Also protect pressure points, such as bony protrusions, from pressure necrosis or peripheral nerve damage by providing padding between the animal and the table.
Monitor respiratory function throughout the procedure and recovery.

Monitor respiratory rate and depth (compare to normal for the species. You can expect them to be slightly decreased). Observe, or use a stethoscope or esophageal stethoscope.
Monitor the color of the mucous membranes (gums, conjunctiva, vulvar mucosa). A bluish color means the animal is not getting enough oxygen, ventilate!
Red-tinged foam present in the airway along with dyspnea (difficulty breathing) may indicate pulmonary edema. This can result from overventilation or overhydration. A diuretic like furosemide can be administered, but prognosis is poor.
Sophisticated respiratory monitoring can be achieved by measuring blood gasses, or expired oxygen and carbon dioxide concentration or by use of a pulse oximeter.

b. Fluid Therapy/Cardiovascular Support

Many anesthetics have direct effects on the heart or vasculature, decreasing cardiac output and blood pressure. This is further complicated by increased fluid requirements during anesthesia and surgery that may result in hypovolemia. Fluid requirements are increased because: breathing dry, cold oxygen (if inhalant anesthesia is used) increases respiratory fluid loss; the animal has not received its normal fluid intake since it was fasted; fluid may be lost through hemorrhage or exposure of moist viscera to room air; many anesthetics are metabolized in the kidney (creating a slight diuresis minimizes renal toxicity).

To minimize the effects of surgery and anesthesia on hydration:

Place an intravenous catheter whenever possible to provide access for fluids and medications
Supplement fluids, intravenously if possible; otherwise intraperitoneally or subcutaneously

Fluid should be supplemented at the rate of 5-10 ml/kg/hour during anesthesia
Monitor hydration status- Overhydration results in frequent urination and pulmonary edema, underhydration results in sticky mucous membranes, loss of skin elasticity, the eyes sinking into the orbit, decrease in blood pressure and increase in heart rate
To replace blood loss with saline or lactated ringers, administer 3X the volume of blood lost by slow IV drip. Monitor the hematocrit. If it drops below 20%, whole blood replacement may be necessary.
Monitor cardiovascular function by monitoring one or more of the following:

Mucous membrane color and capillary refill time (the time it takes for the mucous membranes to regain their normal color after pressure is applied)
Heart rate and rhythm- stethoscope or esophageal stethoscope
Pulse rate and pressure- using your fingers
Blood pressure- arterial catheter or Doppler cuff required
EKG

If the animal has pale mucous membranes, the capillary refill time is greater than 2 seconds, or if the other cardiovascular parameters are out of normal range (determine normal for the species you are using!) you may have a cardiovascular emergency. Increasing the rate of intravenous fluid administration will improve cardiac output temporarily. However the depth of anesthesia will need to be reduced and if there is a primary cardiac problem it will require specific treatment.

c. Thermoregulation

Animals frequently become hypothermic during anesthesia because of inhalation of cold gasses, exposure of body cavities to the room air, and loss of normal thermoregulatory mechanisms and behaviors. Hypothermia depresses all physiologic functions, including respiration and cardiac function, slows the metabolism of anesthetics and results in prolonged recoveries. All of these can contribute to anesthetic death. Hyperthermia is less common, but may occur because of excessive application of heat, hot surgery lights or malignant hyperthermia in genetically pre-disposed animals. To thermoregulate your patient:

Monitor the body temperature frequently using a thermometer during the procedure and during anesthetic recovery. While animal normals vary from species- to-species, in general, when body temperature drops below 99° F, an animal is considered hypothermic. Below 95-96° F an animal cannot regain normal body temperature without supplementation.
Prevent heat loss by insulating cold surfaces with a blanket
Prevent heat loss during gas anesthesia by utilizing low flow techniques that conserve heat
Supplement heat with a water-driven heating pad (do not use electric heating pads, they will cause severe burns) and/or with pre-warmed fluids
Treat hyperthermia by administering intravenous fluids or applying water to foot pads or exposed skin. Only use an ice bath as a last resort, as it may cause cardiovascular shock.

d. Monitoring Anesthesia

The depth of anesthesia must be monitored carefully. Animals that are too light will experience pain and may move during the procedure. Animals that are too deep run the risk of experiencing cardiopulmonary arrest. If an animal is too light the anesthesia should be supplemented, if too deep, animals on gas anesthesia can be turned down. Animals given injectable anesthetics can not be lightened directly. Instead respiratory and cardiovascular support must be administered until the anesthetic is metabolized and the animal begins to lighten on its own.

To monitor the depth of anesthesia, perform the following:

Reflexes- these reflexes disappear as the animal becomes deeper in the following order:

Palpebral reflex- touching the eyelids causes blinking. The animal is light if it is blinking.
Toe pinch reflex- pinching the toe or foot web will cause a pain response. If the animal withdraws the toe it is not deep enough. If it doesn't, it is not sensing pain.
Corneal reflex- touching the cornea of the eye with a tuft of cotton results in a blink. Once the animal has lost its corneal reflex, it is too deep.

Muscle tone decreases as the depth of anesthesia decreases, unless the animal is receiving a cataleptic drug like ketamine in the absence of a sedative. Test muscle tone by pulling on the lower jaw or a limb. Rigid tone indicates inadequate depth of anesthesia.
Monitor cardiopulmonary function and body temperature. As an animal becomes too deeply anesthetized, respiration and cardiac output decrease, resulting in poor blood oxygenation and tissue perfusion and decreased blood pressure and temperature. Likewise, elevations in heart rate and blood pressure may be indications that an animal may be feeling pain and is anesthetized too lightly. Monitor as previously described.

3. Skeletal Muscle Relaxants (Curariform Drugs)

Curariform drugs induce complete muscle relaxation (paralysis) by acting at the neuromuscular junction. They are often classified as depolarizing (e.g. succinylcholine, decamethonium) or non-depolarizing (e.g. curare, gallamine, pancuronium). These drugs are occasionally used in conjunction with a general anesthetic (particularly barbiturates or halothane) to induce complete muscle relaxation which may be needed during certain parts of specific surgical procedures. Artificial positive pressure respiration must be used because the respiratory muscles are also paralyzed.

These drugs are not anesthetic and have no analgesic effect. Thus, though an animal may be completely paralyzed by a muscle relaxant, it is fully conscious and feels all stimuli, painful and otherwise. For this reason, these drugs must be used only in animals under general anesthesia. However, a serious problem of using muscle relaxants with anesthetics is that it is difficult to assess the adequacy of the depth of general anesthesia in the presence of a muscle relaxant drug. If it is not possible to choose an anesthetic drug whose duration of anesthetic action is known to be longer than that of the muscle relaxant drug, it is essential to properly monitor the animals throughout the entire procedure. Monitoring the effect of painful stimuli (e.g. toe pinch) applied at frequent intervals (10-15 minutes) on EEG, EKG or blood pressure is usually considered sufficient. It is recommended that the procedure is performed without paralytic as a trial run.

Analgesia Guidelines

Animal welfare guidelines recommend that all vertebrates that undergo procedures that might cause more than momentary pain or distress be treated with analgesics, unless it can be scientifically justified that the treatment will frustrate experimental objectives.

IACUC requires that analgesics be administered to all USDA covered species that experience procedures that may cause more than momentary pain or distress to animals, unless it is scientifically justified that doing so will frustrate experimental objectives. This justification must be included in the IACUC protocol. IACUC also recommends that analgesia be administered to rodents undergoing comparable procedures.

Analgesics should be administered immediately post-procedurally, and dosing repeated at intervals appropriate for the analgesic and the procedure. For example, a simple skin incision may only require 24 hours of analgesic administration, while orthopedic procedures may need analgesics for up to a week. Rodents experiencing abdominal surgery or similar procedures normally only require analgesic administration for the first 12 hours. Large animals undergoing similar procedures will normally be given analgesics for 72 hours. If analgesia is withheld until the animal is fully recovered to prevent interference with recovery, long acting local anesthetics should be used to assure that the animal remains pain free during this period. It is not appropriate to wait until signs of pain or distress are demonstrated before administering analgesics. Once pain is perceived, it requires more analgesics to relieve pain than if the analgesic was given prior to pain onset. In fact, in rodents, the signs of pain following surgery are manifested as decreases in food and water consumption and weight gain, usually within the first 12 hours following the procedure. These effects are not normally observed unless they are specifically measured. Administration of opiates will ameliorate these effects. During this time animals are also experiencing normal changes in behavior consequent to anesthetic administration. For these reasons, monitoring for the presence of clinical signs of pain is an unreliable method of determining if analgesics are required.

Mice

Injectable:
100mg/kg Ketamine + 10mg/kg Xylazine SQ or IP: 1ml Ketamine (100mg/ml) + 0.5ml Xylazine (Rompun) (20mg/ml) + 8.5ml saline; inject 0.1ml/10g bodyweight. Duration 30-45 minutes.

Inhalation:
for short term anesthesia:
2-4% Methoxyflurane (Metofane): may be used in bell jar or open drop nose cone to effect. Duration 1-20+ minutes.
or
80% CO₂ + 20% O₂ mix: ideal for eye-bleeds and tail-biopsies. Duration 30 seconds to 2 minutes (if the animals remains immersed in the gas mixture).
for long term anesthesia:
2-3% Isoflurane and Halothane: require a precision vaporizer (spontaneous evaporation at room temperature reaches lethal concentrations of 20-30% --> do not use in bell jar). Duration as desired.

Analgesics:
Post-OperativeAnalgesia in Rodents Policy

Rats

Injectable:
75mg/kg Ketamine + 5mg/kg Xylazine SQ or IP: 3ml Ketamine (100mg/ml) + 1ml Rompun (20mg/ml); inject 0.1ml/100g bodyweight. Mixture may be used for up to 6 months if prepared and used using sterile technique. Duration of anesthesia is approximately 30 minutes. If needed, it can be redosed with 1/3 to 1/2 of the original dose to prolong anesthesia time.

Inhalation:
for short term anesthesia:
2-4% Methoxyflurane (Metofane): may be used in bell jar or open drop nose cone to effect. Duration 1-20+ minutes.
or
80% CO₂ + 20% O₂ mix: ideal for eye-bleeds and tail-cutting. Duration 30sec - 2minutes (if the animals remains immersed in the gas mixture).
for long term anesthesia:
2-3% Isoflurane and Halothane: require a precision vaporizer (spontaneous evaporation at room temperature reaches concentrations of 20-30% --> do not use in bell jar). Duration as desired.

Analgesics:
Post-OperativeAnalgesia in Rodents Policy

Rabbit

Injectable:
35mg/kg Ketamine + 5mg/kg Xylazine SQ (IM causes muscle necrosis and may lead to auto-mutilation)

Analgesia:
1mg/lb Carprofen PO BID
or
0.01-0.03mg/kg Buprenorphine IM BID