The general aim in maintaining marine mammals in captivity is to duplicate their natural environment as closely as possible. In the US, the Marine Mammal Protection Act of 1972 specifies that coliform bacterial counts of water for captive marine mammals must be ≤1,000 most probable number (MPN)/100 mL, although systems should routinely be maintained well below that number.
Marine mammals housed in the extremes of their temperature tolerance range are more susceptible to environmental and infectious disease. In general, cetaceans, pinnipeds, otters, and polar bears are better adapted to cold than to heat, but species-specific tolerances differ. Sirenians are generally adapted to warmer waters and become hypothermic in cold-water conditions. Inappropriately combining different species for display purposes can result in compromises that jeopardize the health of some species.
Good air quality, especially in indoor facilities (10–20 air changes/hour) is as important as good water quality. Outdoor exhibit detritus should be kept away, and environment drains should not empty into pools. Photoperiods, light spectral and intensity requirements, sound tolerances, and flight distance requirements are not well-established for any marine mammal. Extremes in any of these factors should be considered detrimental in the absence of specific data.
Cetaceans: Most cetaceans live in marine habitats, although some species migrate into freshwater, and five species of river dolphins have adapted completely to freshwater habitats. Marine cetaceans must be kept in water with a salinity of 25–35 g/L, preferably using balanced sea salts. Water for captive marine cetaceans should be maintained as close to the pH of mid-ocean waters (8–8.3) as possible. Freshwater cetaceans require water similar to that of their natural habitat. Variation in pool depth and a nonuniform environment are recommended for a more acoustically enriching environment.
Pinnipeds: Environmental requirements of pinnipeds are similar to those of cetaceans, except that pinnipeds commonly haul out (ie, temporarily leave the water). Although captive pinnipeds can be kept in freshwater, saltwater pools that meet the specifications listed for cetaceans are preferred, except for the freshwater Baikal seals (Phoca sibirica). Most pinnipeds obtain their metabolic water requirements in food and do not require access to freshwater if provided fish with a high fat content. However, it is common practice to allow pinnipeds access to potable water. Pools for captive pinnipeds should provide shelter from wind and some shade. Haul-out requirements are different for each species, but several haul-out options should be provided to allow animals to separate themselves on land.
Sirenians: Manatee subspecies vary in the time they spend in freshwater, but the dugong (Dugong dugong) is completely marine. Sirenians are tropical and subtropical species with water quality requirements similar to those of cetaceans and will become hypothermic with prolonged exposure to water that is too cold (below ~20°C). The most common sirenian in the US, the Florida manatee (Trichechus manatus latirostris), migrates between marine and freshwater environments seasonally. Manatees do better in captivity if salinity is changed seasonally to match migration in the wild.
Sea otters: Captive sea otters must be housed in a cold marine water system. As sea otters do not have blubber, fur is their major protection against hypothermia; thus, the water must be kept free of oils and organic material that could mat or damage the coat. Sea otters should also have dry haul-out areas available.
Polar bears: Polar bears naturally live on arctic and subarctic ice. Polar bears have successfully adapted to subtropical climates in captivity but are more susceptible to dermatologic disease in warm climates. Polar bears traditionally have been provided with freshwater in captivity, but salt or brackish water is beneficial. Proper attention to filtration and water quality is beneficial, and variable terrain and ample pool space and sunlight should be provided. A secluded den area is required for pregnant females.
Restraint of Marine Mammals
Various restraint techniques are used to perform a thorough examination of marine mammals, including training (behavioral), manual, mechanical, and chemical restraint. Behavioral restraint is widely used to facilitate examination and collection of diagnostic samples; these techniques are highly advantageous as they can reduce potential injury and stress to both animals and humans. For these animals, the presence of familiar attendants is important. Examples of diagnostic samples collected from trained animals include blood, nasal or respiratory samples, gastric, sputum, ocular and skin samples, urine, and feces. Diagnostic procedures can include numerous elements of the physical examination, ultrasonography, radiography, and intraocular pressure measurement. However, alternative methods of restraint must be available for emergencies as sick or injured animals may not respond to normal training cues.
Cetaceans and sirenians: For complex procedures or untrained animals, the safest approach to restraining a cetacean or sirenian is to remove it from the water. Captive enclosures should allow water drainage so that animals can be stranded without the use of nets; a built-in lift floor is the most rapid way to accomplish this. As the animal begins to lose buoyancy in the draining water, it should be positioned over thick foam pads to minimize struggling and injury. Nets are an alternative for corralling small cetaceans and sirenians kept in sea pens or encountered in the wild; however, experienced personnel are required to minimize the risk of drowning or injury to the animal or staff. Netted animals are placed on foam or specially designed stretchers or floats that can suspend the animal partially within or above water level.
Small cetaceans such as dolphins can often be restrained by the weight of three or four attendants: one person controls the peduncle of the tail fluke and the others apply weight alongside the animal’s body, although care must be taken to avoid restricting normal respiration. Additional personnel can be used for larger species such as beluga whales. The pectoral fins should be tucked alongside the animal in a natural position to avoid permanent damage and should never be used to position an animal. In larger cetaceans, the powerful tail fluke may be secured with mechanical restraints, although this can also result in damage to the animal and humans; thus, excessive caution is warranted. Sirenians are relatively docile but have a very powerful tail fluke; problems in restraint are generally due to their bulk and weight, and caution is recommended because they tend to roll.
Pinnipeds and sea otters: Capturing pinnipeds is easier and safer on dry ground, although small animals can be captured in the water with end-release hoop nets. Larger animals should not be netted in water but should be coaxed or driven from the water or have the water drained from their pool. On land, hoop nets can be used on larger animals. Cargo nets, baffle boards, and snare poles (come-along poles) also can be helpful. Once captured, smaller pinnipeds can be restrained for some shorter procedures by an experienced handler sitting on the seal’s back and holding the head; the handlers should rest the most of their weight on their knees and use their legs to restrain to avoid restricting respiration. Larger pinnipeds or more complex procedures require an appropriately designed squeeze cage or chemical restraint. Sea otters can be removed from pools with hoop nets. Once they are out of the water, well-padded restraint bags, squeeze boxes, or other restraint devices for small wild carnivores can be used.
Polar bears: Polar bears are large and dangerous, and manual restraint is not advised. Some training techniques can be used, especially if combined with squeeze cages or similar, but protected contact is strongly recommended.
Anesthesia of Marine Mammals
Physiologic adaptations to diving and the aquatic environment can make general anesthesia of marine mammals challenging. Anesthetic drugs commonly used in other animals often have narrow margins of safety or cause unexpected reactions in marine mammals. Numerous tranquilizers, sedatives, and anesthetics have been used in different species but should be administered to marine mammals only by personnel experienced in their use. Careful planning and training is of critical importance as morbidity and mortality associated with anesthesia can be high in these species. Reversible drugs are desirable as they may decrease recovery time. Close monitoring of heart rate and quality (ECG), respiratory rate, temperature, end-tidal carbon dioxide, and anesthetic depth and perfusion is required. Marine mammals are prone to atelectasis, and many drugs reduce respiration rate and quality or result in apnea. Some common anesthetic agents and dosages are provided in the table Common Injectable Sedative and Anesthetic Agents used in Mixed Pinnipeds and Cetaceans.
Common Injectable Sedative and Anesthetic Agents used in Mixed Pinnipeds and Cetaceans
Drug | Otariids | Phocids | Cetaceans |
|---|---|---|---|
Alfaxalone | Aa: 2–3 mg/kg, IM Zc: 1–2 mg/kg, IM, with midazolam (0.1–0.2 mg/kg) Recommend in combination with a benzodiazepine | Pv: 1–1.5 mg/kg, IM or IV Better effects in combination with a benzodiazepine | — |
Butorphanol | Ej: 0.1–0.2 mg/kg, IM Zc: 0.1–0.3 mg/kg, IM Reversible | Hl: 0.05–0.17 mg/kg, IM Pv: 0.1–0.3 mg/kg, IM or IV Reversible | Dl, Lo, Oo, or Tt: 0.05–0.15 mg/kg, IM |
Diazepam | Aph: 0.04–0.28 mg/kg, IM or IV Zc: 0.05–0.3 mg/kg, PO Reversible | Hg: 0.3 mg/kg, IM Lc: 0.1–0.25 mg/kg, IM Ml: 0.02–0.35, IV or IM Pv: 0.05–0.32 mg/kg, IV or IM Reversible | Dl: 0.16 mg/kg Oo: 0.25–1 mg/kg, PO Tt: 0.1–0.22 mg/kg, PO or IM Reversible |
Ketamine | Aa: 1 mg/kg, IM Ag: 6–7 mg/kg, IM Aph: 2–6 mg/kg At: 1.9–2.8 mg/kg, IM Zc: 1–4 mg/kg, IM Better effects in combination with a benzodiazepine | Hg: 6 mg/kg, IM Lw: 1.6–2.4 mg/kg, IM Lc: 2.9–7.7 mg/kg Ml: 1.8–3.4 mg/kg, IV, or 2–8 mg/kg, IM Better effects in combination with a benzodiazepine | 1.1 mg/kg |
Medetomidine | Cu: 0.01–0.04 mg/kg, IM Ej: 0.02–0.06 mg/kg, IM Zc: 0.01–0.07 mg/kg, IM Reversal recommended | Ml: 0.012–0.027 mg/kg, IM Reversal recommended | — |
Meperidine (pethidine) | 0.1–0.45 mg/kg, IM Respiratory depression at higher dosage | Hg: 2 mg/kg, IM Hl: 1–1.5 mg/kg, IM Lc: 1–3 mg/kg, IM Ml: 1.2–6.7 mg/kg, IM Reversible | 0.1–0.5 mg/kg, IM |
Midazolam | Aa: 0.1–0.7 mg/kg, IM Ej: 0.2–0.25 mg/kg, IM Zc: 0.1–0.3 mg/kg, IM Reversible | Hg: 0.2–0.3 mg/kg, IM Hl: 0.16–0.38 mg/kg, IM Lw: 0.08–0.5 mg/kg, IM Lc: 0.15–0.85 mg/kg, IM Ml: 0.02–0.07, IM Pg: 0.2 mg/kg, IM Pv: 0.1–0.3 mg/kg, IM or IV Reversible | Tt: 0.05–0.15 mg/kg, IM or IV Reversible |
Propofol | Zc: 2–5 mg/kg, IV (anesthetic induction) | Pv: 1.25–2 mg/kg, IV; 2-6 mg/kg, IV (anesthetic induction) | Tt or Lo: 3–5.5 mg/kg, IV (anesthetic induction) |
Tiletamine-zolazepam | Aa: 1–1.4 mg/kg, IM Cu: 0.77–1.14 mg/kg, IM Ej: 1.8–2.5 mg/kg, IM Ph: 1.6–2 mg/kg, IM Zc: 1 mg/kg, IM | Hg: 0.5–1.6 mg/kg, IM Hl: 1.2–1.4 mg/kg Lw: 0.3–1.1 mg/kg, IV or IM Ma or Ml: 0.4–1.2 mg/kg, IV or IM | — |
Species Key | Aa: Arctocephalus australis (South American fur seal) Ag: Arctocephalus gazella (Antarctic fur seal) Aph: Arctocephalus phillipi (Juan Fernandez fur seal) Apu: Arctocephalus pusillus (South African fur seal) At: Arctocephalus tropicalis (subantarctic fur seal) Cu: Callorhinus ursinus (northern fur seal) Ej: Eumetopias jubatus (Steller's sea lion) Ph: Phocarctos hookeri (New Zealand sea lion) Zc: Zalophus californianus (California sea lion) | Hg: Halichoerus grypus (grey seal) Hl: Hydrurga leptonyx (leopard seal) Lw: Leptonychotes weddellii (Weddel seal) Lc: Lobodon carcinophagus (crabeater seal) Ma: Mirounga angustirostris (northern elephant seal) Ml: Mirounga leonina (southern elephant seal) Pg: Phoca groenlandica (harp seal) Pv: Phoca vitulina (harbor seal) | Dl: Delphinapterus leucas (beluga whale) Lo: Lagenorhynchus obliquidens (Pacific white-sided dolphin) Oo: Orcinus orca (killer whale Tt: Tursiops truncatus (bottlenose dolphin) |
Cetaceans: General anesthesia is not routinely performed in these species, particularly as many procedures can be accomplished with training or sedation, in conjunction with local anesthetics, with the animal out of water. Specialized anesthetic machines and respirators (apneustic plateau) are required for cetaceans. General anesthesia can be induced with propofol administered IV or other agents, maintained with injectable agents or inhalant anesthesia, and requires endotracheal intubation. Following the loss of reflexes, including jaw tone, the mouth is held open, the modified larynx (goosebeak) is manually palpated and dislocated cranially from the nasopharyngeal sphincter, and an endotracheal tube is placed into the trachea. The trachea is very short; thus, length of the endotracheal tube insertion must be closely checked. The goosebeak must be manually replaced into proper position during recovery, and the cetacean must be closely monitored until normal respirations have fully resumed as renarcotization is common.
Pinnipeds: Otariids, phocids, and odobenids are routinely anesthetized both in captivity and in the field for various procedures. A variety of injectable agents have been successfully used, although reversible agents may be particularly desirable. Inhalant anesthetics, usually isoflurane and sevoflurane, have also been used for maintenance or induction of anesthesia in smaller animals. Endotracheal intubation is recommended, as apnea and atelectasis are very common in these species. The trachea bifurcates prior to entering the thorax in otariids; thus, caution must be used with endotracheal tube placement. Walrus have a pharyngeal pouch that can be accidentally intubated, so use of a long laryngoscope or endoscope to visualize the larynx is recommended. Mechanical ventilation is recommended for longer procedures, and particularly when animals are positioned in dorsal recumbency as hypotension is common. Ephedrine is effective in treating hypotension.
Sirenians: Sirenians rarely require general anesthesia, but sedation (usually with benzodiazepines) may be recommended for some diagnostic tests and treatments. Endotracheal intubation is done through the nasal cavity and is best accomplished by use of endoscopy, either by visualizing the larynx through one nare and inserting the endotracheal tube through the other, or placing the endotracheal tube over the endoscope directly and visualizing the larynx. Similar to cetaceans, the sirenian trachea is also short. A mechanical ventilator should be used to ensure adequate ventilation, and a large ventilator system should be used.
Sea otters: In both wild and captive situations, sea otters are most commonly anesthetized with fentanyl, IM, in combination with midazolam or diazepam. Endotracheal intubation is recommended for longer procedures, and the appropriate endotracheal tube size is generally smaller than would be expected for terrestrial animals of similar size. Inhalant anesthetics such as sevoflurane and isoflurane can be used to supplement injectable anesthetics. Narcotic recycling has been documented; thus, opioid reversal with naltrexone is recommended. Close monitoring of temperature is particularly important in sea otters.
Polar bears: Polar bears are routinely immobilized with etorphine, tiletamine-zolazepam with or without medetomidine, ketamine with xylazine, or a variety of other agents administered IM. The required dose is highly dependent on the individual animal and environment. Once anesthetized, precautions should be taken to ensure human safety in case of unexpected or sudden anesthetic recovery.
Sample Collection in Marine Mammals
Blood Collection
Marine mammals have vascular adaptations that can make blood collection and sample interpretation challenging, particularly arteriovenous anastomoses within the peripheral vasculature that often result in mixed arteriovenous blood. Ultrasonography can be helpful in locating vessels in many species and is highly recommended for catheterization. Aseptic technique should always be followed when collecting blood samples or placing catheters.
Cetaceans: Blood is collected most commonly through the ventral or dorsal fluke vessels of the periarteriole venous rete with a butterfly needle and vacuum blood collection tube or with a needle and syringe. A slight furrow or indentation of the vascular tree can usually be seen or palpated. Blood can also be collected from the periarteriole venous rete in the dorsal fin or foreflipper in some species. The peduncle vein can be accessed via a lateral or ventral approach, and this site is usually used for larger blood samples or catheterization.
Pinnipeds: The most common blood collection sites from phocids and walruses are the epidural intervertebral vein or sinus and a plantar interdigital vein. The epidural intervertebral vein is generally accessed from the dorsal midline, 1–3 intervertebral spaces cranial to the pelvis, and is reliable for large sample collection, whereas the interdigital vein usually yields smaller samples. In otariids, the caudal gluteal vein, interdigital vessels of the hind flippers, and ventral brachial vein of the front flippers are most commonly used. The caudal gluteal vein is caudal to the anterior pelvis, approximately one-third to half the distance from femoral trochanter to the base of the tail and just lateral to the sacral vertebrae. It can be accessed with either a vacuum blood collection tube and needle holder setup or with a needle and syringe. The interdigital vessels vary in size and prominence among species, and pressure (eg, a tourniquet around the tarsus) and local warming of the flipper may help open the vessel, which can be directly seen and palpated. Similar techniques may enable brachial venous access. The jugular vein and subclavian vein can also be used for IV access in these species but generally when under anesthesia.
Sirenians: Sirenian blood is usually collected from the brachial vascular bundle of the pectoral flipper, located between the radius and ulna, with either a medial or lateral approach.
Sea otters: Blood is usually collected from the popliteal or femoral vein in conscious sea otters and can also be collected from the jugular and cephalic vein in anesthetized otters.
Polar bears: Polar bear blood is usually collected from the jugular, femoral, saphenous, or cephalic veins under anesthesia, although small amounts can be collected from conscious trained bears from the digital vessels.
Urine Collection
As with other species, urine is most commonly collected via cystocentesis, catheterization, or free catch. Some dolphins and other cetaceans have been trained for urine collection. It may be difficult to collect urine via catheter or cystocentesis from some species, such as manatees, in which case free catch is valuable. A shallow dish can be placed under the urinary opening in manatees in ventral recumbency, and pinnipeds and otters can be held temporarily in cages lined with bars or grates on the bottom, allowing urine to fall through.
Gastric Sample Collection
Gastric samples for cytologic evaluation and to assess pH and other parameters are common in captive dolphins. Many dolphins are trained to allow passage of a flexible gastric tube to both deliver oral hydration and collect samples. Most dolphins are amenable to this training technique given the unique structure of their modified larynx and esophagus; however, some pinnipeds have also been trained to allow gastric tube insertion and sample collection.
Respiratory Sample Collection
Respiratory samples are also routinely collected from captive cetaceans to establish baseline cytologic profiles and monitor for potential disease or the presence of new pathogens, such as fungi. Dolphins are generally trained to exhale strongly (ie, chuff) upon request. A petri dish or other sampling medium is held above the blowhole to capture exhaled particles for evaluation.
