Fish Anesthesia

Adequate oxygenation is necessary for maintenance of normal physiologic homeostasis during fish anesthesia. Oxygen should be supplied, via an air stone or similar device, to all water chambers used during transport, anesthesia and recovery. Ideally, water taken from the original fish holding tank should be used in transport, anesthetic and recovery chambers. If water from another source is used, water quality parameters (i.e., chlorine, temperature, pH and ammonia) should closely duplicate that of the original holding tank. Fish are often fasted for 12-24 hours prior to anesthesia to reduce the risk of regurgitation of food that may lodge in the gills or foul the water. Fasting also decreases fecal contamination of water used for anesthesia and recovery. When working with a new anesthetic protocol or species it is advisable to anesthetize a few fish and follow them through full recovery to insure drug dosages and techniques are safe and provide sufficient anesthetic depth for the intended procedures.

When a fish reaches the level of anesthesia sufficient to perform surgery, there is a total loss of equilibrium and muscle tone, decreased respiratory rate and no response to stimuli. A firm squeeze at the base of the tail may be used to determine response to stimuli. Respiratory rate may be evaluated by observing movement of the operculum (rigid flap that covers the gills) as it opens and closes. Gill color should be dark pink to light red. If respirations become extremely slow or stop, the fish may be placed in anesthetic-free recovery water until respirations resume. Water temperature should be maintained at the species normal optimum during both anesthesia and recovery.

Anesthetic Agents Used in Fish

Anesthetic Agent	Dose	Comments
MS-222 (tricaine methanesulfonate)	75-125 mg/l (induction) and 50-75 mg/l (maintenance)	Sodium bicarbonate should be added to stock solution to maintain neutral pH. Only FDA approved anesthetic for fish (21 day withdrawal).
Clove oil	40-100 mg/l (depending on fish species and anesthetic depth)	Active ingredient is eugenol. Clove oil stock solution (100mg/ml) made with 95% ethanol. Stock solution is added to induction chamber at 40-100 mg/l. Not approved for use in fish that may be used for food or released into public waterways.
Benzocaine hydrochloride	25-100 mg/l	Sodium bicarbonate may need to be added to stock solution to maintain neutral pH. Small margin of safety between effective and lethal doses.

 

 

 

 

 

 

 

 

 

 

 

For additional information regarding anesthetic use in fish please consult an ARP veterinarian or the following references.

1. Stetter MD: Fish and Amphibian Anesthesia. In Veterinary Clinics of North America: Exotic Animal Practice 4(1), January 2001; pp. 69-82.
2. Burka JF, et al: Drugs in Salmonid Aquaculture - A Review. J. Vet Pharmacol. Therap. 20, 1997; pp. 333-349.
3. Bowser PR: Anesthetic Options for Fish. In Recent Advances in Veterinary Anesthesia and Analgesia: Companion Animals, 2001. International Veterinary Information Service, Ithaca, NY.

             

Amphibian Anesthesia

Amphibians in the early stages of their life cycle (e.g., tadpoles) are entirely aquatic and have gills for respiration. Most amphibians lose their gills during metamorphosis and develop lungs. Most importantly, amphibian skin acts as a semipermeable membrane that allows for respiration (cutaneous respiration) and absorption of substances through the skin. Depending on life stage, amphibians may be anesthetized by immersion in an anesthetic solution, placement in an anesthetic gas induction chamber or by anesthetic preparations applied to the skin. Fasting for 12-24 hours prior to anesthesia is recommended to decrease the incidence of regurgitation which will foul the water of the anesthetic or recovery container. Amphibians may also show an excitement phase during anesthetic induction. It is important to induce anesthesia in a container that will prevent injury due to the animal jumping or falling out.

General Anesthesia Use in Amphibians

Stages of Anesthesia in Amphibians
Induction	Light anesthesia	Surgical anesthesia
Decreased gular movement and diminished withdrawal reflex	Loss of righting reflex and absence of abdominal respiration	No withdrawal reflex (toe pinch) and gular movements cease

 

 

 

 

 

After the amphibian reaches the appropriate level of anesthesia for the planned procedure, it should be removed from the anesthetic bath and rinsed with fresh water or the topical preparation removed by rinsing with fresh water. The animal will remain anesthetized for 10-80 minutes, depending on the method and drug concentration used (see below for anesthetic drugs and dosages). Pulmonary respiration will cease during anesthesia in amphibians and cannot be used to monitor anesthetic depth. Cutaneous respiration is sufficient to prevent clinical hypoxia during anesthesia. Heart rate may be monitored during anesthesia by direct observation (ventral midline, caudal to the shoulders), ECG, ultrasonagraphy or a Doppler flow detector. Normal values for heart rates have not been published. Recovery from anesthesia (righting reflex returns and animal able to move normally) will take 30-90 minutes after the animal is rinsed with fresh water.

• Do not raise the amphibian's body temperature above that of normal room temperature in an attempt to speed recovery. Increasing the body temperature will increase metabolism and oxygen requirements. Cutaneous respiration may not be sufficient to maintain adequate oxygenation.
• Do not apply alcohol or other preparations that contain alcohol directly to the skin of an amphibian. It may be absorbed through the skin and may dissolve normal secretions that protect the animal from dehydration and infections.

 

Anesthetic Agents Used in Amphibians

Anesthetic agent	Dose	Comments
MS-222 (tricaine methanesulfonate). Wide margin of safety. Anesthetic solutions must be buffered with sodium bicarbonate to maintain neutral pH.	250-500mg/l  1g/l 2-3 g/l	Tadpoles and newts Frogs and salamanders Large specimens and toads
Benzocaine (powder or hydrochloride). Powder must be dissolved in ethanol to create a stock solution. Solutions must be buffered with sodium bicarbonate to maintain neutral pH.	2 gm/l	True toads, spadefoots and large salamanders
Isoflurane may be mixed into a viscous solution with KY jelly and water or injected into an absorbent pad and applied directly to the dorsum of the animal.	Variable	Applied to skin in patch or viscous gel

 

 

 

 

 

 

 

 

 

For additional information regarding anesthetic use in amphibians please consult an ARP veterinarian or the following references.

1. Stetter MD: Fish and Amphibian Anesthesia. In Veterinary Clinics of North America: Exotic Animal Practice 4(1), January 2001; pp. 69-82.
2. Green DE: Anesthesia of Amphibians in the Field. Amphibian Research and Monitoring Initiative Standard Operating Procedure, March 2001. National Wildlife Health Center, United States Geological Service.
3. Smith JM and Stump KC: Isoflurane Anesthesia in the African Clawed Frog ( Xenopus laevis ). Contemporary Topics, 39(6), November 2000; pp. 39-42.
4. Wright KM and BR Whitaker: Amphibian Medicine and Captive Husbandry. 2001; pp. 115-121. Krieger: Malabar, Florida. 

             

Anesthesia & Analgesia In Wildlife

Due to the wide variation in anesthetic and analgesic drug use requirements in wildlife species specific drug names and dosages are not listed on this website. Please contact an ARP veterinarian for information on anesthesia and analgesia when using wildlife in research or teaching protocols.

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The information and drug dosages presented in this website are intended as a resource for Pennsylvania State University research investigators. No guarantee of drug efficacy or safety is made nor must information obtained from this site be substituted for professional veterinary advice.