1. Wear clean, wet exam gloves when handling fish.
   1. Nitrile gloves which are latex- and powder-free are preferred.  If gloves with powder are used they should be rinsed thoroughly to remove residue before handling fish.
   2. Moisten gloves with clean dechlorinated water, or water from the animal's home tank or environment.
   3. In some field conditions, clean wet hands (without gloves) are acceptable.
2. Use nets to handle fish whenever possible as fish have a protective mucus layer on their skin and are slippery and difficult to hold. Always use a net to restrain non-sedated fish.
3. Work surfaces for medical/surgical procedures must be non-abrasive but also prevent the fish from sliding, such as a soft moist cloth or sponge.
4. Anesthetize fish during any procedure that is stressful or likely to cause pain.
5. Ensure fish remain wet during procedures out of the water. Water can be dripped, poured, or sprayed on exposed areas of the fish.
   1. Use water from the home tank wherever possible; clean non-chlorinated water is also acceptable.

1. General Considerations
   1. There is tremendous inter-species variation, and anesthetic protocols that work for one type of fish may not be effective in another. Prior to use of any anesthetic regimen it is recommended to consult the literature and investigators with experience in the species of interest. For anesthetic regimens in unfamiliar species or use of agents with little information available in fish species, a pilot study with a small number of animals is strongly advised prior to initiating larger scale administration. Neiffer (2009) is a good starting resource for individual species options. Refer to Appendix A for general recommendations.
   2. Consider withholding food from fish for up to 24 hours prior to anesthetic events. Regurgitation can interfere with gill function and contaminate the surgical area. Contact ULAM veterinary personnel for fasting recommendations of specific species.
2. Delivery Methods
   1. Immersion
      1. This is analogous to inhalant anesthesia in air-breathing species and occurs when a diluted anesthetic is absorbed by the gills and/or the skin to produce anesthesia
         1. Advantages of immersion anesthesia include consistent response and a relatively high safety margin, rapid recovery following removal to fresh water, and the lack of physical restraint for induction
         2. Disadvantages include difficulty changing the anesthetic concentration in the water during the procedure, lack of suitability for longer procedures (>10 minutes), and the irritant nature of some anesthetics. Despite these potential disadvantages, anesthesia by immersion is generally the anesthetic method of choice.
      2. Induce anesthesia in an anesthetic-treated tank. Monitor the animal to determine an appropriate depth of anesthesia has been reached (see Procedures section 2.c).
      3. Use buffers to establish and maintain a physiologically appropriate pH as several anesthetics can significantly alter the pH of water.
      4. Use water from the animal's home tank in the anesthesia and recovery tanks whenever possible.  Fish are very sensitive to changes in water pH, temperature, and mineral content, and consistent water quality between the fish's normal holding tank, and the anesthetic and recovery tanks is extremely important.
   2. Flow Anesthesia
      1. For procedures out of water lasting more than a few minutes, recirculating or continuous flow systems using aerated water containing anesthetic can be used to provide both continuous anesthesia and artificial ventilation.
      2. Induce anesthesia using the immersion method above when using flow anesthesia.  Transition the animal to either of the methods listed below when appropriate level of anesthesia is reached.
         1. Intubation: The recirculating delivery system utilizes a submersible pump that is placed within a tank containing the anesthetic solution. The anesthetic water is pumped from this tank, into the animal's mouth, over the gills and out through the opercula. The animal is positioned on a fenestrated board above the anesthetic water tank. This method is commonly used in larger fish but has also been described in small fish such as zebrafish (26).
         2. Drip: The non-recirculating method uses a (new) IV bag and drip set as the anesthetic water reservoir and delivery method. The size of the bag and the drip rate depend on the size of the animal being anesthetized (see Stetter 2001 for specific size recommendations).
         3. Adjust the rate such that water flows into the mouth, gently over the gills and out the opercula for either flow anesthesia approach. Inappropriate flow rates can interfere with gas exchange or force water into the gastrointestinal tract.
   3. Parenteral
      1. Administer IM injections into the epaxial muscles. There may be leakage of anesthetic as the muscles contract post- injection. Insert the needle between scales.
      2. IV injection produces rapid induction of anesthesia but requires manual restraint or prior sedative administration. The caudal vein lies ventral to the tail vertebrae and can be accessed by a ventral mid line or lateral approach.
      3. IC (intracoelemic) administration is analogous to IP injections in mammals. It is effective but induction times are inconsistent and there is risk of visceral damage.
      4. Additional considerations when working with injectable anesthetics include inconsistent sedation or anesthesia that may require supplementation with immersion techniques, and the potential for prolonged recovery requiring artificial ventilation.
3. Monitoring Anesthesia
   1. The depth of anesthesia in fish can be monitored by observing the behavior of the fish in water. While appropriate monitoring parameters can vary based upon anesthetics and species, several general guidelines can be used for monitoring anesthetic depth. See Procedures section 2.c.ii for the stages of anesthesia and Appendix A for anesthesia in fish.​
      1. Activity decreases, the righting reflex is lost, and muscle tone decreases as fish become anesthetized.
      2. Opercular movement (respiratory rate) progressively decreases with deepening anesthesia.
         1. Hypoxemia can occur in fish despite good ventilatory efforts and can be recognized by pallor of the gills and the fin margins.
      3. The heart rate can be directly monitored using a Doppler blood flow probe or ECG leads.
         1. Monitor trends when using the Doppler or ECG; reference ranges are not known for many fish species.
         2. Surgical planes of anesthesia can be confirmed by a lack of response to a firm squeeze at the base of the tail.
   2. Fish Stages of Anesthesia

d. Anesthetic Agents

1. Immersion/Flow Anesthetics
   1. MS-222 (Syncaine, TMS, tricaine methanesulfonate) is the recommended agent. 
      1. Ensure MS-222 is pharmaceutical-grade before using. Currently there is only one pharmaceutical-grade MS-222 preparation available:
         1. Syncaine (MS-222) – manufactured by Syndel.
      2. Dissolve MS-222, a chemical powder, in water (fresh or salt) prior to usage. MS-222 is acidic in solution and must be buffered to a physiologic pH (7.0-7.4) prior to usage.
      3. Store MS-222 powder in the original sealed container in a dry location at room temperature until the expiration date noted by manufacturer on packaging. Ideally MS-222 stock solutions are utilized the same day as preparation per vendor recommendation. When necessary, stock solutions of MS-222 may be kept up to 30 days. They must be refrigerated and stored in tinted (amber) or opaque bottles. Stock solutions of MS-222 that are older than 30 days, or that have not been properly stored must not be used. All MS-222 powder and stock solution containers must be appropriately stored, labeled (concentration and preparation or expiration date), and used prior to expiration date. (Alpharma, 2001 and Pharmaq, 2010)
      4. Wear nitrile gloves, lab coat, and goggles and utilize a biosafety cabinet or chemical fume hood when handling MS-222 as it is hazardous to humans. Alternatively use a top loading balance with a clear plastic wind guard. Extended direct contact to skin can cause a reversible retinopathy.
      5. Contact the University of Michigan’s Department of Environment, Health & Safety (734-647-1143) for appropriate disposal methods as MS-222 solutions cannot be poured down the drain or introduced into the general water supply.
      6. The only substance approved by the FDA for field sedation of fishes is MS-222. However, MS-222 use in the field is limited because of an FDA requirement that food fish, including feral fishes that may be caught and eaten by humans, must go through a 21-day withdrawal period prior to release or slaughter for human consumption (American Fisheries Society, 2014; 21CFR529.2503). Therefore, any wild fish anesthetized with MS-222 that are of a species consumed by humans and of legal size (see table 4 in Western Chemical, 2014) must either be held for the 21-day withdrawal period prior to release or euthanized. Alternatively, other methods of anesthesia or sedation requiring special permitting can be used. Please contact ULAM veterinary personnel if anesthesia or sedation of wild fish that will be released back into the wild is required.
      7. If the animal begins to recover prior to completion of the procedure, there are several options:
         1. Apply a paper towel or gauze soaked in the original MS-222 solution directly to skin with care to avoid the surgical site.
         2. Drip the MS-222 solution directly onto the skin with care to avoid the surgical site.
         3. Place the animal back into 50% of the original concentration of MS-222 if there is no open incision.
   2. Benzocaine
      1. Local anesthetic and parent compound of MS-222, which is less water soluble and less acidic.
      2. Low toxicity in humans and improved safety in species sensitive to MS-222.
      3. Fat solubility may result in prolonged recovery, especially in old or gravid fish.
   3. Lidocaine
      1. Local anesthetic that has been used for surgical anesthesia in some fish species, including zebrafish and medaka.
      2. Recovery time may be longer than with MS-222.
      3. May be an effective anesthetic in combination with propofol in some species (zebrafish).
   4. Metomidate (Aquacalm)
      1. Nonbarbituate hypnotic used for sedation in ornamental finfish (not suitable for surgical levels). Primarily used outside the US.
      2. Longer recovery compared to MS-222. Not FDA approved.
   5. Etomidate
      1. Ultra-short acting non-barbiturate agent that provides rapid induction, but prolonged recovery. There is no analgesic effect.
      2. May be used for sedation or light anesthesia in adult zebrafish. May cause minor post-procedural effects on behavior. (Jorge et al, 2021).
   6. Alfaxalone
      1. Synthetic neurosteroid anesthetic agent. The multidose formulation (Alfaxan Multidose) has a shelf life of 28 days once the vial is breached.
      2. Used an an immersion anesthetic in some fish.  Induction and recovery times may be longer than with MS-222, but it is considered a safe and viable alternative in species in which it has been tested.
   7. Clove Oil (Eugenol/Iso-eugenol)
      1.  A natural anesthetic used in aquaculture industry to reduce handling stress.  Active ingredients are eugenol and isoeugenol. Clove oil stock solution (100mg/ml) made with 95% ethanol. Stock solution is added to induction chamber at 40-100 mg/l.
      2. There is a ready-made solution (AQUI-S 20E) with the active ingredient iso-eugenol that is sold in the USA under the Investigational New Aquaculture Drug (INAD) program. It can be mixed with treatment water and used for sedation prior to handling or minor procedures (27).
      3. Not approved for use in fish that may be used for food or released into public waterways.
   8. Gradual Cooling
      1. Shown to be useful in zebrafish for short-term, minor procedures as well as surgical procedures.
      2. Faster recovery than with MS-222 in zebrafish and no distressful behaviors observed.
      3. Home tank/system water should be placed in a container within a larger container containing cold tap water. Ice is added to the outer tank and a slurry formed until inner tank temperature reaches 17°C and place fish into inner container.  Add more ice incrementally to the outer container until fish are anesthetized (typically occurs at about 10°C).
2. Parenteral Anesthetics - dosing depends upon species and route of administration
   1. Ketamine: Use in combination with a 2-agonists (dexmedetomidine).
      1. Provides safe and effective anesthesia in some species; can cause respiratory depression, bradycardia, and poor immobilization in others
      2. Consistently effective as an aid to restraint or capture and for short, minor procedures.
   2. α~2~-agonists:
      1. Medetomidine, in conjunction with ketamine, has been used in fish but currently has limited availability. Dexmedetomidine has not been evaluated in fish, but can be used at half the recommended medetomidine dose in other species.
         1. Usually used in combination with ketamine to improve muscle relaxation and anesthetic duration and depth.
         2. Effects can be reversed with the administration of atipamezole.
         3. Xylazine can produce apnea and convulsant activity and is not recommended.
3. For anesthetic agents for use in fish, see Appendix A
4. Recovery/Post-Anesthetic Care
   1. Recover fish in aerated, untreated water.
   2. During recovery, a reversal of the stages of anesthesia should occur with a gradual increase in opercular movement, return of equilibrium, and eventual resumption of normal swimming.
   3. Most fish are fully recovered from immersion anesthetics within 5 minutes of placement in fresh water. Prolonged recoveries (> 10 minutes) indicate excessive anesthetic depth or a compromised animal. Recovery from parenteral anesthetics can be highly variable.
   4. Fish may pass through an excitement phase during recovery and may attempt to escape from the recovery tank. Stimulation can exacerbate the excitement phase and once fish are showing progressive signs of recovery (increased opercular and fin movement, increased muscle tone, and a return of equilibrium) it may help to cover the tank with a lid. Occasionally, fish demonstrate vigorous movement and may need to be restrained to prevent self-injury.
5. Emergency Care
   1. Immediately place the animal in untreated, aerated water and initiate forced ventilation if respiration stops altogether. Forced ventilation stimulates the buccal flow/heart rate reflex and provides support while speeding the elimination of the anesthetic.
      1. Forced ventilation is accomplished by moving the fish gently and slowly FORWARD in a circular path through the water. Dragging the fish backward through the water may result in damage to the gills. For any fish with questionable opercular movement, gentle forced ventilation can help speed initial recovery. Forced ventilation enhances opercular movement and increase the passage of fresh, oxygenated water over the gills.
   2. Respiratory arrest can precede cardiac arrest by several minutes; continue resuscitation efforts for several minutes despite a lack of immediate improvement.

1. General Considerations
   1. Pain and the use of analgesia remains a controversial issue in fish medicine. Marked differences in neuroanatomy and behavior exist between fish and higher, homeothermic vertebrates, and analgesic use in fish has not been systematically evaluated. While currently there is not overwhelming evidence of efficacy for most analgesics in fish, some beneficial effects on behavior and physiologic parameters have been reported in some instances, and the drugs that have been tested did not have significant adverse side effects. However, it is important to note that there may be considerable interspecies variation in response to analgesics. In the absence of contrary evidence, conditions or procedures that are expected to cause pain or distress in a human are to be considered painful or distressing to fish and alleviated accordingly. Consult veterinarians on specific recommendations.
2. Analgesic Agents
   1. Opioids - Fish have mu and kappa receptors suggesting a role for opioids. Suggested opioids and doses are in Appendix B.
   2. NSAIDs - The analgesic effects of NSAIDs in fish are questionable, but the anti-inflammatory effects may provide benefit. See Appendix B.
   3. Topical analgesia - amino amides (lidocaine)
3. For analgesic agents for use in fish, see Appendix B
4. Signs of Stress. Distress, and Pain
   1. Fish react to noxious stimuli, distress, and discomfort. Signs of distress and discomfort in fish may include, but are not limited to, the following:
      1. Listlessness
      2. Decreased appetite
      3. Loss of body condition
      4. Abnormal behavior
      5. Social isolation
      6. Abnormal orientation
      7. Darker coloring
      8. Rapid opercular movement
      9. Rapid gill or mouth movement
      10. Agitated swimming

1. Preparation of the Surgical Area
   1. According to the Guide for the Care and Use of Laboratory Animals: Eighth Edition, "For most survival surgery performed on rodents and other small species...the space should be dedicated to surgery and related activities when used for this purpose, and managed to minimize contamination from other activities conducted in the room at other times." (pg. 144)
      1. The surgical area should be a room or a portion of a room that is easily sanitized and not used for any other purpose during the time of surgery.
      2. Clean and disinfect the surface upon which the surgery is performed with an approved environmental disinfectant before beginning the surgical procedure.
2. Preparation of Surgical Supplies
   1. Surgical Instruments
      1. Use sealed aseptic surgical supplies whenever possible.
         1. Initial steam sterilization (autoclaving), plasma vapor sterilization, vaporized hydrogen peroxide, or ethylene oxide sterilization (for heat or pressure sensitive items) is required for all surgical instruments and items to be implanted.
3. Preparation of the Animal
   1. Fasting
      1. Consider fasting animals for up to 24 hours prior to anesthesia depending on animal size, species, and procedure. Consult ULAM veterinary personnel to determine if fasting is appropriate/recommended.
   2. Anesthesia
      1. Use an approved agent appropriate for the species AND the procedure.
      2. Maintain the animal in a surgical plane of anesthesia throughout the duration of the procedure
      3. Skin disinfection
         1. Due to the protective mucus layer present on fish skin, aggressive cleansing is not recommended. The skin should be prepared with a single wipe using sterile saline or a dilute povidone iodine or chlorhexiderm solution immediately before surgery. Removal of scales is generally not recommended and should be avoided or minimized when possible.
4. Surgeon Preparation
   1. Wash hands thoroughly with a disinfecting soap such as chlorhexidine or iodine based surgical scrubs or 3M Avaguard^® hand antiseptic.
   2. Required attire for the surgeon during the surgical procedure:
      1. Mask
      2. Sterile or clean gloves
         1. Clean gloves include unused standard latex or nitrile lab gloves stored in a sealable bag or container to minimize dust and debris contamination.
      3. Clean scrub top, clean disposable PPE gown, or clean lab jacket
      4. Refer to EHS Animal Handler PPE Chart if alternate PPE accommodations are necessary.
5. Performing Multiple Surgeries in Series
   1. Begin with at least 2 sets of sterile instruments.
   2. Clean the instruments and sterilize with a hot bead sterilizer between each animal.
      1. It is imperative that tools are completely cooled after sterilization to avoid thermal damage to the animal.
      2. Cold sterilization products require prolonged contact times, present health hazards the animals and/or surgeons, and are not recommended.
   3. No more than 5 animals should be operated on per pack of sterile instruments.
   4. Use new clean or sterile gloves for each animal.
   5. Clean the surgical area with an appropriate disinfectant between animals.
6. Suture Materials and Wound Closure
   1. Use monofilament suture material (e.g. PDS, Maxon, or Ethilon) to minimize tissue reactions and infection that could slow healing time. Braided materials such as vicryl or silk are generally contraindicated for skin closure in fish despite their historical use. Tie sutures snugly for a watertight seal. Post-operative swelling in fish is minimal, unlike mammals.
   2. Absorbable suture materials have been recommended as a means of avoiding additional handling for removal. Remove all skin sutures (absorbable or nonabsorbable) by 14 days after surgery if they are still present (provided the skin incision is adequately healed), unless described otherwise in an IACUC approved protocol or as recommended by ULAM veterinary personnel to necessitate complete wound healing.
   3. Tissue adhesives (e.g. cyanoacrylate) are not recommended for use in fish (associated with tissue reactions and wound dehiscence). Similarly, surgical staples are regarded as inferior to appropriately placed sutures for wound closure.
   4. Routine prophylactic antimicrobial use is generally not necessary if appropriate aseptic technique is utilized. Contact ULAM veterinary personnel for cases that may warrant antibiotic treatment (contaminated wound repair, inadvertent bowel perforation, other break in aseptic technique).
7. Post-Operative Monitoring and Care
   1. Monitor animals every 15 minutes during recovery from anesthesia until they resume normal swimming (see Procedures section 2.d.iv).
      1. Return animal to its home tank when normal swimming/activity has resumed.
   2. Post-operative medications including analgesics, antibiotics and/or anesthetic reversals should be administered during the early recovery period and according to the approved protocol or the advice of ULAM veterinary personnel.
   3. Affix a yellow acetate with a Surgery Observation Sticker (SOS) to the tank unless approved by the IACUC to use a different form of cage labeling.  Keep the label on the tank for 7 days after surgery or until the skin sutures are removed (if applicable), whichever is longer.
      1. The label should include the date of surgery.
   4. Records should document animal condition, pre and post-operative drugs, concerns noted during surgery and post-operative recovery notes. Post-operative monitoring can be in the form of a post-surgical record or as poorly recovering post-operative fish clinically appear similar to a spontaneously ill fish, the post-operative monitoring can be incorporated into the daily health check process with associated daily documentation.
      1. Records must be kept near the animal, either in the animal housing room or in an adjacent procedure area.
   5. Monitoring should continue until skin sutures are removed, or if sutures are not placed, fish should be monitored specifically for post-operative complications in the first 24 hours following surgery and after that time their post-operative monitoring can be incorporated into daily health checks.