Challenges with Elasmobranch Capture and Anesthesia in Large Aquariums
American Association of Zoo Veterinarians Conference 2012

Natalie D. Mylniczenko, DVM, MS, DACZM

Disney’s Animals, Science and Environment, Bay Lake, FL, USA


Immobilization of large elasmobranchs in multimillion-gallon aquatic systems provides many technical difficulties and requires special skill sets. Prior to embarking on an anesthetic event with large elasmobranchs, it is highly recommended that a pre-procedure briefing occurs with clear goals and assigned staff roles, that safety issues are recognized and that an emergency protocol is in place in the event of a human safety incident (on land and underwater). The procedure itself should be as quick as possible without comprising animal or human safety.

Immediate Areas of Concern

1.  Human safety (below)

2.  Target animal safety

a.  Potential risk of anesthesia

b.  Attention must be paid to conspecifics that may be aggressive and damage the sedate individual

c.  Focal damage from darts or darting devices and equipment or net damage

d.  The larger the animal the more possible it is to inflict damage to the animal itself (e.g., spinal damage) and organ damage (flipping large rays can result in hepatic fracture if not done carefully).

3.  “Collateral damage”

a.  Non-target animals

i.  Exposure to anesthetic drugs

ii.  Traumatizing/stressing them due to proximity or presence

b.  Exhibit space damage

i.  Glass or plexiglass cracks or scratches by anesthesia or restraint equipment

ii.  ‘Furniture’ in the exhibit can be broken, such as coral heads.

When thinking about human safety, handlers should be experienced with elasmobranchs; if they are divers, they should be physically fit and outfitted with proper protective gear. Whether they are on land or underwater, constant attention must be paid to the location of the oral cavity of the animal, to the skin of the sharks and to the barbs of stingrays (even if trimmed). Protective items can include poles or baffles to keep animals at a distance, Kevlar™ or chain mail gloves or clothing where indicated, protective tubing over barbs/tail protrusions (especially with freshwater rays), and nose devices for sawfishes. There should also be a plan in place for accidental exposure of staff members to anesthetic drugs (as with zoo hoofstock). Ultrapotent narcotics are not typically used; however, alpha 2 agonists at dosages used for large elasmobranchs are a significant concern.

Removal from Enclosure

How do you remove a single animal from a mixed species large aquarium? Some basic categories for approaching these animals are:

1.  Training and removal from the aquarium

a.  The animal is trained into a device which can be removed from the water (a cage, a sling, a net).

2.  Training animals into smaller areas for delivery of anesthetic drugs or for manual restraint. The animal is:

a.  Targeted into a device or smaller space (a medical pool) for net or manual restraint

b.  Cornered or baffled and anesthetic drugs are delivered

c.  Is moved into an even smaller device (a swimming pool) for use of immersion drugs

3.  Surprise catch

a.  Scoop method: The animals are manually caught in a net or other restraint device by catching them off the surface during a feed or patterned swim movement.

b.  Catching the animals under water without sedation: manually or with nets.

i.  In some cases, a net across the entire tank that is capable of moving toward a wall and can be used to ‘push’ animals and isolate the target animal or group.

a)  This requires a great deal of dive staff, depending on the size of the enclosure, maneuverability around the exhibit and you can get many non-target species.

b)  This can be a very effective and rapid technique with a skilled team.

4.  Anesthetic pre-sedation or tranquilization and then restraint.

a.  Drugs

i.  Oral (fed out): often requires very high dosages and many classic mammalian drugs have little or no overt effect (needs more research)

ii.  Injectable: see section below

iii.  Immersion: smaller area, animal separated (e.g., swimming pool), versus whole tank exposure.

a)  Obvious difficulties depending on total animal numbers and volume of water

b)  Very good at moving an entire collection of animals if appropriate drugs are used

c)  Can require a lot of staff

d)  Need to consider effluents

e)  Drugs:

(1)  Tricaine methane sulfonate (MS-222)

(a)  Varied doses

(b)  Graded anesthesia (high induction, lower maintenance)

(2)  2-Phenoxyethanol

(3)  Eugenol

iv.  Over the gill: high doses of immersive drugs, most typically tricaine methane sulfonate delivered focally over the gills or through the mouth for induction

b.  Restraint devices

i.  Sling

ii.  Sock

iii.  Box

iv.  Box net

v.  Hoop net, large

vi.  Manual restraint

Injections-Technical Aspects

When a procedure requires injection of the animal underwater while it is free swimming then a number of additional considerations must be had. What tools are available?

1.  Hand syringe

a.  Results in very close contact with the animal

b.  Must use proper angle and strength

2.  Pole syringe

a.  Must use proper angle and strength; need to be quick

3.  Dart guns (e.g., laser aimed or other similar underwater gun)

a.  Practical range of 8 feet

b.  Compression of air at depth alters discharge

c.  Usually used at the surface or in shallow water

d.  Pushing dart through the water is harder than it seems, difficult to gauge pressures

e.  Cost of procuring or producing an underwater dart gun

4.  Hawaiian sling/other similar (e.g., speargun)

a.  Very difficult to judge projectile strength

b.  Recovery procedure versus euthanasia

What are some of the technical difficulties with injections into elasmobranchs?

1.  Thickness of skin, very abrasive skin, dulls needles

2.  Potentially large volumes of drugs

a.  Concentrated drugs are recommended

b.  Fish muscles are not elastic and cannot hold as much as mammal muscles

3.  Location of injection

a.  Intramuscular

i.  Red muscle (aerobic) is best but is less likely for injection than white muscle (anaerobic)

ii.  Location on the body: the ‘saddle’ is the reputed best spot

b.  Intravenous

i.  Intended for

a)  Conditioned animals (stingrays) or slow-moving larger elasmobranchs (whale sharks)

b)  Manual restraint

4.  Leakage of drug

a.  The dart should stay in animal for several minutes to allow sufficient discharge and to prevent flowback (out of the injection site).

i.  The animal’s response to the dart being removed is a useful indicator of the level of induction.

b.  Leakage post injection can be as much as 20% (pers. comm. M. Andrew Stamper).

5.  Darts not going off at depth.

6.  Human limitations

a.  Assumption of distance underwater is challenging

b.  Compensation for the angle of refraction of the mask

Some Successful Intramuscular Drug Combinations

What is successful? The expectations of sedation are important. Most reports of using injectable anesthetics have resulted in highly varied results ranging from no sedation to sedation that lasted for several days and in some cases, death. Mostly however, the general result is an animal that is notably affected by the drug (slow, less likely to respond to a human and mildly ataxic) and can be led into a safer restraint device. Some drug combinations that have been successful (see references for details on dosages and responses):

  • Ketamine and xylazine ± midazolam
  • Ketamine and medetomidine (or dexmedetomidine)
  • Etomidate
  • Butorphanol and medetomidine
  • Medetomidine or dexmedetomidine alone
  • Alphaxalone-alphadolone (Saffan) (no longer available)

Considerations for Induction

Regardless of the methods chosen, efficiency and speed are key, particularly with pelagic species. Elasmobranchs follow three general lifestyles: benthic, intermediate and pelagic. The more pelagic the species, the more physiologic stress that animal will endure with an ensuing lactic acidosis. However, consideration for how much struggling even a benthic animal does during the initial capture and or induction is also an important consideration. Habituated or trained animals tend to handle stressors better than newly caught or naïve animals. Obligate ram ventilators must have a continual stream of water pumping over the gills, which may mean that underwater pumps must be available to dive staff. The pelagic animals (e.g., Carcharhinus limbatus and C. acronotus) are strict ram ventilators and must be ventilated as soon as possible as they are very susceptible to hypoxia and capture stresses. These animals are not good candidates for underwater injections or long capture attempts. The intermediate species, such as C. plumbeus, C. melanopterus, and C. taurus tolerate handling and short periods of poor ventilation well. Benthic species such as Ginglymostoma cirratum are highly tolerant. Stingrays, in general, even pelagic rays, are seemingly more tolerant of handling stresses. Exertional rhabdomyolysis can occur, though the more immediate and life-threatening blood gas fluctuations are more common. The lifestyle of the animal should guide how to best immobilize elasmobranchs.

Induction quality and length of time vary with the choice of drugs and modalities of capture. Injectable anesthetics can take as long as an hour for induction. Typically, these animals begin to have an awkward gait and start bumping into exhibit materials. Where safe and possible, guiding the animal away from such objects is warranted. Under some circumstances, there need to be divers prepared with baffles and restrictive devices to keep other animals away from the animal that is being induced. Naturally curious or well-trained animals may prove to be a nuisance under these circumstances. Animals, like sawfish, can be hazardous to divers or to large nets and must be kept away from the focus animal and anesthetic equipment.

Anesthetic Maintenance

Once animals are induced and considered safe to handle, following other basic principles of fish and elasmobranch anesthesia are prudent and can be found in the below references. For elasmobranchs, key elements for stable anesthesias will include continued ventilation (from beginning to full recovery), good water quality, and periodic blood gas evaluation (getting a baseline is fundamental). Ultrasound evaluation of cardiac contractility with skilled eyes offers another method to determine how well animal is performing under anesthesia.


As above, safety is key and all the above points bear weight during recovery. It is preferable to recover animals in an isolated area, if not a medical pool then a penned area. If this is not possible, then retaining the animal until it is capable of evading tankmates is necessary. It is a judgment call of when to release an animal, as they are rarely ready at the first notice of voluntary motion. This can be difficult with pelagic species as they need continual ventilation until the last moment. A common issue with fish handlers, is the desire to “walk” animals or to place their oral cavity into an outflow of water. This is not an efficient method of gas exchange and can be detrimental during the recovery phase of anesthesia. The most effective method of ventilation (as evidenced by blood gasses) is with a low flow pump that is directed over both gill arches. This becomes a human safety risk as the hands are near the oral cavity at this timeframe and sudden movements from the animals can result in trauma to the handler, therefore proper protective gear is important. Rays and certain sharks have crushing plates that can inflict serious injury and the shark mouth is itself a dangerous area even for an incidental scratch by casual contact with the teeth. Once the animal is released, it is not uncommon for them to fall to the bottom of the enclosure; depending on the stage of recovery, it may be appropriate to leave the animal. However, if the animal is a ram ventilator or still does not have the ability to propel itself well in the water column, it must be retrieved and further supported until it is ready to swim. If possible, the animal can be positioned in the direct path of a water inlet pipe or with a water pump to ensure steady flow of water over the gills during the recovery phase.


Special thanks to the aquarium and hospital staff at Disney’s The Seas with Nemo and Friends® at Epcot®, Disney’s Animal Health Department, and especially Dr. Mark Penning. Additional thanks to the aquarium and hospital staff at the John G. Shedd Aquarium and Tonya Clauss, DVM, MS, Director of Animal Health, Georgia Aquarium.

Literature Cited

1.  Penning, M.R. 2012. Immobilizing marine fish for transport or surgery—from angelfish to tiger sharks, whether individuals or multi species groups. North American Veterinary Conference, Orlando, FL.

2.  Clauss, T. Berliner, A. Brainard, B. 2011. Pilot studies with select sedatives & anesthetics in bonnethead sharks (Sphyrna tiburo). International Association for Aquatic Animal Medicine conference, Las Vegas, Nevada.

3.  Clauss, TM, Mylniczenko, ND, Stamper, MA. 2012, in preparation. cartilaginous fishes: elasmobranchs & holocephalans. In West, G, Heard, D, Caulkett, N. (Eds). Zoo Animal and Wildlife Immobilization and Anesthesia Cartilaginous Fishes: Elasmobranchs & Holocephalans.

4.  Smith, M.F.L., Marshall, A., Correia, J.P., Rupp, J. Chapter 8 Elasmobranch Transport Techniques and Equipment. In: Smith, M., D. Warmolts, D. Thoney, and R. Hueter (eds). The Elasmobranch Husbandry Manual: Captive Care of Sharks, Rays and their Relatives. Ohio Biological Survey: 2004;105–131.

5.  Vaughan D.B., Penning M.R., Christison K.W. 2-phenoxyethanol as anaesthetic in removing and relocating 102 species of fishes representing from Sea World to uShaka Marine World South Africa. Onderstepoort Journal of Veterinary Research. 2008;75:189–198.


Speaker Information
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Natalie D. Mylniczenko, MS, DVM, DACZM
Disney's Animals, Science, and Environment
Bay Lake, FL, USA

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