Acup 110 Fish And Amphibian Anesthesia - Research

    Drowning Fish. Anesthesia procedures for fish and amphibians. investigators and staff who have approval to anesthetize fish and amphibians in their . accidental drowning. www.research.cornell.edu.

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Acup 110 Fish And Amphibian Anesthesia - Research
ACUP110.01 CONTROLLED DOCUMENT Page 1 of 7
Document: ACUP110.01
Issue Date: 01 OCT 15; Effective Date 01 OCT 15
Authorization: Dr. D. Winkler, IACUC Chair
Author: Dr. J. Gourdon
FISH AND AMPHIBIAN ANESTHESIA
1. PURPOSE
1.1. The intent of this Animal Care and Use Procedure (ACUP) is to describe common
anesthesia procedures for fish and amphibians. This ACUP is approved by the
Cornell IACUC. Any deviation must be approved by the IACUC prior to its
implementation.
2. SCOPE
2.1. This ACUP is intended for use by investigators and staff who have approval to
anesthetize fish and amphibians in their study protocol.
3. INTRODUCTION
3.1. This document provides guidelines for fish and amphibian anesthesia, procedures,
and post-anesthetic care and monitoring. Contact CARE at [email protected] for
more information or for other assistance.
4. MATERIALS AND EQUIPMENT
4.1. Anesthetic agent (e.g. pharmaceutical grade tricaine methanesulfonate [MS-222]),
sodium bicarbonate (if using MS-222).
4.2. Gloves (powder-free, pre-moistened).
4.3. Transport, anesthetic, and recovery tanks.
4.4. Oxygenation equipment (e.g., air pump, tubing, and air stone).
5. PROCEDURE
5.1. General Considerations
5.1.1. If using a new anesthetic protocol or species, anesthetize a small cohort of
animals. Follow them through to recovery to ensure drug dosages and
techniques are safe and provide sufficient anesthetic depth for the intended
procedures.
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5.1.2. Do not disturb the mucus layer of fish or amphibians. When handling animals,
wear powder-free gloves pre-moistened with distilled or dechlorinated water.
Do not apply detergents or solvents to the animal’s skin, and limit contact with
abrasive materials (e.g. dry paper towels).
5.1.3. Ensure that all water used for fish and amphibians is well-oxygenated, chlorine-
free (dechlorinated or distilled), and within the normal temperature range for the
species.
5.1.4. For help designing an anesthetic protocol contact CARE at [email protected].
5.2. Fish Anesthesia
5.2.1. Fast fish for 12–24 hours prior to anesthesia.
5.2.2. When possible, use water taken from original fish holding tank for transport,
anesthetic, and recovery chambers. If using another water source, closely
duplicate the water quality parameters (i.e., chlorine, temperature, pH and
ammonia) of the original holding tank.
5.2.3. Anesthesia is achieved by immersing the animal in an anesthetic solution (see
Appendix for a partial list of agents and doses).
5.2.4. Stages of anesthesia in fish:
Stage 1:
Deep Sedation
Stage 2:
Deep Narcosis
Stage 3:
Surgical Anesthesia
• Cessation of voluntary
swimming
• Decreased response
to stimuli.
• Decreased muscle
tone
• Equilibrium loss
• Appropriate level for
fin and gill biopsies.
• Decreased respiration
and heart rate
• Total loss of response
to stimuli
• Firmly squeeze at the
base of the tail to
determine response
to stimuli.
5.2.5. Allow animal to reach appropriate level of anesthesia prior to beginning
procedures.
5.2.6. While performing procedures, keep the fish’s skin moist and the gills
submerged or regularly flushed with well-oxygenated water.
5.2.7. Evaluate respiratory rate and gill color throughout anesthesia:
5.2.7.1. Observe movement of the operculum (rigid flap that covers the gills) as it
opens and closes to assess rate.
5.2.7.2. Observe gill color; should be dark pink to light red.
5.2.7.3. If respirations become extremely slow or stop, place the fish in
anesthetic-free recovery water until respirations resume. If the fish must
be re-anesthetized after recovery, proceed with a decreased
concentration of MS-22 and monitor respiration closely.
5.3. Amphibian Anesthesia
5.3.1. Fast for 12–24 hours prior to anesthesia.
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ACUP110.01 CONTROLLED DOCUMENT Page 3 of 7
5.3.2. When possible, use water taken from the original holding tank for transport,
anesthetic and recovery chambers. If using another water source, closely
duplicate the water quality parameters (i.e., chlorine, temperature, pH and
ammonia) of the original holding tank.
5.3.3. Anesthetic induction may produce an excitement phase and must be performed
in a container that will prevent the animal from jumping or falling out.
5.3.4. Anesthesia is achieved by immersing the animal in an anesthetic solution (see
Appendix for a partial list of agents and doses).
5.3.4.1. If alternative methods of anesthesia are desired, contact CARE at
[email protected] for assistance.
NOTE: When inducing a terrestrial amphibian in an immersion anesthetic-
bath, keep the animal’s head and nares above the water line (to prevent
accidental drowning).
5.3.5. Stages of anesthesia in amphibians:
Induction
Light Anesthesia
Surgical Anesthesia
• Decreased gular
movements
• Diminished
withdrawal reflex
• Loss of righting reflex
• Absence of abdominal
respiration
• Loss of withdrawal
reflex (toe pinch)
• Absent Gular
movements
5.3.6. Allow animal to reach appropriate level of anesthesia for planned procedures.
5.3.7. Remove the animal from the anesthetic bath after appropriate level of
anesthesia is reached. Keep the amphibian’s skin moist throughout
procedures.
5.3.8. The animal will remain anesthetized for 10–80 minutes, depending on the
method and drug concentration used.
5.3.9. If supplemental anesthesia is needed, anesthetic solution can be dripped onto
the animal’s skin to effect.
5.3.10. Monitor heart rate during anesthesia (e.g. direct observation [ventral midline,
caudal to the shoulders], Electrocardiogram [ECG], Doppler flow detector).
NOTE: Normal values for heart rates are not well established in amphibians.
5.4. Post Anesthetic Care
5.4.1. General Considerations
5.4.1.1. Closely monitor fish/amphibians recovering from anesthesia until they are
swimming/moving normally and have completely regained their righting
response.
5.4.2. Fish
5.4.2.1. Place the fish in un-medicated water in a holding tank.
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5.4.2.2. To speed recovery, create a flow of oxygenated water over the gills by
either:
5.4.2.2.1. Gently moving the fish back and forth in the water or
5.4.2.2.2. Opening and closing the mouth several times.
5.4.3. Amphibians
5.4.3.1. After procedures are completed, thoroughly rinse the animal with fresh
water.
5.4.3.2. Recovery chamber:
5.4.3.2.1. Aquatic Species – place animal in well-oxygenated, un-medicated
water in a holding tank.
5.4.3.2.2. Terrestrial Species - place animal in a container lined with wet
towels until fully recovered.
5.4.3.3. Do not raise the amphibian's body temperature above that of the species’
normal range in an attempt to speed recovery.
6. PERSONNEL SAFETY
6.1. Medical Emergencies:
CALL 911.
6.2. When working with animals wear appropriate PPE, observe proper hygiene, and be
aware of allergy, zoonosis, and injury risks. Refer to the CARE Occupational Health
and Safety webpage for more information.
6.3. MS-222 (tricaine methanesulfonate) safe practices:
6.3.1. Wear protective clothing, gloves, and goggles when handling MS-222 powder or
animals exposed to MS-222.
6.3.2. If possible, work inside a fume hood to prepare a concentrated stock. Mix MS-
222 powder in a volume of water appropriate to obtain the desired concentration
and based on manufacturer’s recommendations. Wear gloves and use a utensil
to stir until all powder is dissolved.
6.3.3. Dispose of MS-222 waste by flushing down the drain to a sanitary sewer with an
excess of water. Do not discard MS-222 directly into surface water, storm water
conveyances or catch basins.
6.3.4. If in a remote location where a sewer may not be readily available, further dilute
the solution with water and dump wastes on land, in a location away from water.
7. ANIMAL RELATED CONTINGENCIES
7.1. Post contact information for emergency assistance in a conspicuous location within
the animal facility.
7.2. Non-emergency veterinary questions and requests for care, email CARE veterinary
staff at [email protected].
7.3. Emergency veterinary care is available at all times including after working hours and
on weekends and holidays by calling the CARE pager (1-800-329-2456).
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8. REFERENCES
8.1. Neiffer, D.L., Stamper, M.A. 2009. Fish sedation, anesthesia, analgesia, and
euthanasia: Considerations, methods, and types of drugs. ILAR Journal, 50(4), 343-
360.
8.2. Fish, R.E., Brown, M.J., Danneman, P.J., Karas, A.Z. (Eds). 2008. Anesthesia and
Analgesia in Laboratory Animals, 2
nd
Edition. Academic Press, New York.
8.3. Carpenter, J.W. (Ed). 2005. Exotic Animal Formulary, 3
rd
Edition. Saunders, St.
Louis, MO.
8.4. Mitchell, M.A. 2009. Anesthetic Considerations for Amphibians. Journal of Exotic Pet
Medicine, 18(1), 40-49.
8.5. CARE Occupational Health and Safety webpage:
http://www.research.cornell.edu/care/OHS.html
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9. APPENDIX
9.1. Immersion anesthetic agents used in fish and amphibian species
Fish
Anesthetic Agent
Comments
(tricaine methanesulfonate)
and
50–75 (maintenance)
Buffer with equal weight
of sodium bicarbonate
This is the only FDA
approved anesthetic for fish
(21-day withdrawal).
Small margin of safety
between effective and lethal
doses.
Buffer solution with
sodium bicarbonate to
maintain neutral pH
Amphibians
Anesthetic Agent
Comments
(tricaine methanesulfonate)
250-500 mg/L of buffered
aqueous solution
1-2 g/L of buffered aqueous
solution
2-3 g/L of buffered aqueous
solution
Tadpoles
Frogs and salamanders
Toads
Buffer with equal weight
of sodium bicarbonate
hydrochloride)
solution
True toads, spadefoots, and
large salamanders
Buffer solution with
sodium bicarbonate to
maintain neutral pH
9.1.1. MS-222:
9.1.1.1. Always buffer solution with an equal weight of sodium bicarbonate
to maintain neutral pH
9.1.1.2. In solution, MS-222 will lose efficacy if kept longer than 7 days.
9.1.1.3. MS-222 is a light-sensitive chemical and must be kept in a dark container
or in a cabinet/drawer.
9.1.1.4. MS-22 has a wide margin of safety.
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9.1.2. Benzocaine:
9.1.2.1. Always buffer solution with sodium bicarbonate to maintain neutral
pH
9.1.2.2. Dissolve powder in appropriate solvent (e.g. water, or ethanol) to create a
stock solution.
10. HISTORY
Date:
Event:
03 NOV 15
New Format – Converted by: J. Kirby
01 OCT 15
Most Recent Annual Review – Reviewed by: Dr. D. Jeffery
31 JAN 04
New Issued – Original Author: Dr. J. Gourdon; Referee: G. Wooster