Preventive Medicine for Zoo Animals

The examination of animals leaving or entering a collection is a fundamental part of zoo medicine and consists of preshipment examination and quarantine, respectively.

Preshipment examinations are performed at the facility of origin prior to shipment to the receiving institution. The purpose of the preshipment examination is to ensure that the animal is healthy enough to withstand transport, to plan for appropriate transport in light of the animal's health status, and to prevent the spread of transmissible diseases across institutions.

The components of a preshipment examination vary depending on the species, and additional testing or treatment can be requested by the receiving institution or determined by state, federal, and international regulations. Strategic use of preshipment examinations can decrease the time an animal must spend under quarantine, because the risk of disease is mitigated prior to the animal's arrival.

Quarantine is the period during which newly acquired animals are kept separate from the rest of the collection to prevent the introduction of transmissible diseases. Historically, quarantine protocols were strictly standardized for all incoming animals, as many animals came from the wild, and their medical history was limited. Now, most animals come from captive-bred populations with well-documented medical histories.

Accordingly, risk-based quarantine protocols are recommended in zoo animal practice. Veterinarians review the medical history of the individual and of the populations of the animal's origin and identify any transmissible diseases that could be introduced along with them and what consequences the introduction of these diseases would have on their collection. On the basis of the risk analysis, the duration of quarantine, diagnostic tests, and treatments to be carried out during quarantine are determined on an individual basis.

International transfer of animals between facilities is an exception, as these are subject to legislation that can determine many of the quarantine conditions. In the US, imported animals must undergo quarantine in facilities specifically designed according to USDA regulations under strict protocols aimed at preventing the introduction of foreign animal diseases. In the EU, quarantine conditions are detailed in the Balai Directive.

Zoos must obtain import permits in accordance with national and international laws for the acquisition of animals from nonmember states. For example, given the risk of the spread of zoonotic diseases, all primates coming from overseas must undergo quarantine in accordance with the World Organisation for Animal Health (WOAH, formerly the Office International des Epizooties, OIE) Terrestrial Animal Health Code.

Birds originating from areas classified by WOAH as infected zones must undergo indoor quarantine without access to mosquitoes or other potential arthropod vectors. Ventilation systems must be properly filtered to prevent the introduction of vector-borne or airborne diseases.

Ungulates housed in open paddocks during quarantine must have at least two escape-proof fences located at least three meters from other animal enclosures. If kept in herds, the facilities must allow for the separation of individuals from the herd if medical intervention is required.

Generally, quarantine should effectively keep animals out of direct or indirect contact with the rest of the collection by careful planning of footpaths, assignment of dedicated personnel, appropriate use of personal protective equipment, and use of designated tools and equipment. Facilities should allow for proper sanitization and handling of animals for any diagnostics or treatments indicated prior to the end of quarantine, which must be approved by the veterinarian. Shipping crates should be disinfected prior to being removed from the quarantined area.

While under quarantine, animals must be closely monitored for clinical signs of disease or maladaptation related to the stress of transport and husbandry changes. Social enrichment must be provided to social species (eg, nonhuman primates, penguins, flamingos, other birds). Identification should be verified upon entrance to and exit from quarantine. If no identifier is present, one should be placed (eg, tattoo, tag, band, ear tag, passive integrated transponder) to ensure future identification. When new animals are introduced to enclosures, caution and forethought are necessary to prevent self-induced trauma. Visual barriers can prevent injury and promote acclimation of an animal newly introduced to an exhibit.

Comprehensive parasite control is an integral part of any zoo preventive-medicine program. Zoo animals are vulnerable to ecto- and endoparasites, and parasite management includes veterinary and husbandry interventions. Veterinary intervention has typically relied on antiparasitic drugs, with medications selected carefully because of species-specific sensitivities (eg, fenbendazole toxicity in pelicans (1), porcupines (2), pigeons (3), and others), whereas husbandry interventions focus on hygiene, pasture maintenance, and pest control.

In closed pastures, ungulates are susceptible to diseases caused by trichostrongylid parasites, especially Haemonchus spp, a nematode that resides in the abomasum of ruminants and compartment 3 of the stomach in camelids. These parasites have direct life cycles, persist in the environment, and can rapidly develop resistance to anthelmintics, complicating their control in zoological collections. Risk factors for a high trichostrongylid load in pastures include temperatures above 10°C (50°F), > 5 cm (> 2 inches) of rainfall per month, > 8–12 hours per day spent in pasture, grazing < 7.6 cm (< 3 inches) from the ground, herd density > 5 animals per acre, and immunosuppression or malnutrition of susceptible hosts (4).

Objective monitoring of trichostrongylid egg shedding using modified McMaster chambers for fecal egg counts (FECs) should be performed at least every 6–12 months. An FEC reduction test should be performed to assess for anthelmintic resistance in the parasite population of a specific host by performing FEC before starting treatment and 10–14 days after treatment in individual animals. The emergence of resistance is defined by a < 95% decrease in FEC after anthelmintic treatment (4).

Treatment for trichostrongylid parasites should take a comprehensive approach, emphasizing pasture management, control of parasite populations, and treatment of infected and susceptible hosts. To avoid the development of resistance, only one drug class should be used, or two that act synergistically (eg, benzimidazoles and avermectins). One drug class should be reserved for use only in emergency situations. Fasting animals for 24 hours before treatment decreases rumen flow and can increase drug bioavailability and contact time with nematodes in the small intestine (5).

Refugia (parasite populations not exposed to treatment) allow nematodes that have survived anthelmintic treatment to disseminate resistance genes. The number of refugia can increase because of the presence of other host species (ie, those not susceptible to infection by the parasite) in mixed-species pastures and by not treating animals with an appreciable parasite load that do not show clinical signs of disease during quarantine.

Dietary concentrated tannins or fresh or pelleted common lespedeza plants (Lespedeza cuneata) can decrease nematode fecundity. Duddingtonia flagrans is a fungus that kills free-living nematode larvae and can be administered orally with feed. The fungal spores are eliminated in the feces, where they germinate and kill nematode larvae in the pasture without having any biological effect on the animal.

Treatment with copper oxide wire particles (COWPs) does not contribute to the development of resistance. The COWPs are ingested and retained in the folds of the abomasum, where the low pH releases soluble copper ions. These ions kill nematodes such as Haemonchus spp through an unknown mechanism of action. Because only abomasal nematodes and not intestinal nematodes are susceptible to treatment, a larval development test should be performed before treatment to confirm the parasite's identification.

Treatment with COWPs can decrease FECs by up to 90% in 3–4 weeks, although the response to treatment varies depending on the species (6). Adverse effects include liver disease or anemia related to copper toxicity. To decrease the risk of toxicosis, treatment should be limited to a maximum frequency of every 6–12 months.

The most common parasitic diseases of importance in nondomesticated carnivores are the same as for domestic dogs and cats and include GI parasites (roundworms hookworms, whipworms, and tapeworms), parasitic protozoa (coccidia, Giardia), heartworm (Dirofilaria immitis), and various metazoan parasites, including fleas, ticks, and mites. Fecal flotation is recommended 2–4 times a year, and monthly heartworm prevention is recommended in endemic areas.

Treatment and prophylaxis of both ecto- and endoparasites follow the same general principles as in domestic dogs and cats. Avermectins and isoxazolines, such as fluralaner and sarolaner, have been used in zoo animals. Veterinarians are advised to stay abreast of the latest publications to decide whether a new product should be included in their preventive-medicine program for zoo carnivores.

For avian collections, vector-borne diseases and parasites with intermediate hosts are of special concern:

• Black flies (Simuliidae) carry Leukocytozoonspp, a haemosporidian parasite that can cause disease (hemorrhages, anaphylactic reactions, or death) through simuliotoxicosis from excessive biting.

• Midges (Ceratopogonidae) and louse flies (Hippoboscidae) can transmit Haemoproteus.

• Mosquitoes (Culicidae) can carry Plasmodium spp, the causative agent of avian malaria, to which some species (such as penguins) are particularly susceptible.

• These arthropods can also serve as mechanical vectors for the transmission of fowlpox (poxvirus), to which young animals are particularly vulnerable.

• Other diseases transmitted by arthropods include West Nile virus (WNV) and Eastern (EEE) and Western (WEE) equine encephalitis viruses, to which some birds (eg, penguins, corvids, ratites, cranes, storks, and others) are particularly susceptible.

Vector control includes removing standing water wherever possible, treating standing water and drains with larvicides during peak mosquito seasons, removing vegetation near enclosures, holding birds indoors at dawn and dusk (times of peak mosquito activity), and the use of nets, screens, and fans. Routine malaria prophylaxis (eg, with oral primaquine) should be considered for penguins that have outdoor access.

Spirurian nematodes (eg, Serratospiculum), Capillaria spp, air sac trematodes (Szidatitrema spp, among others) and renal trematodes (eg, Paratanaisia bragai) are parasites of concern among zoo-housed birds. Air sac parasites, such as Serratospiculum and Serratospiculoides, can cause respiratory and neurological disease in raptors. They have an indirect life cycle and are transmitted by intermediate arthropod hosts (eg, Tenebrionidae beetles, Eubelidae woodlice).

In addition to arthropod control, treatment for parasites of zoo-housed birds can be achieved through the use of avermectins. Benzimidazoles (eg, albendazole, fenbendazole) should be used with caution in birds, as cases of toxicosis have been reported in several species (1, 3).

Air sac and renal trematodes are not species specific and infest a variety of avian hosts. They are transmitted by ingestion of intermediate mollusk hosts (snails and slugs) and can be particularly problematic in aviaries with natural vegetation and soil substrate. Monitoring is by routine fecal flotation and fecal (and urate) sediment examinations using composite samples collected over several days. Treatment does not usually eliminate infection; however, it can mitigate morbidity and mortality in the collection. Treatment involves systemic praziquantel, physical removal of trematodes by celioscopy, and control of mollusks.

Direct fecal examinations and fecal flotation should be routinely performed on herpetofauna under managed care. This is especially important for animals under quarantine that might have originated from free-ranging populations.

Parasites with direct life cycles are especially important because animals can develop high parasite loads. In reptiles, pinworms (Oxyuridae) are commensals of the lower GI tract and can even aid in digestion. Pinworm eggs are sticky and relatively resistant to disinfection. A high parasite load can be associated with GI inflammation, diarrhea, or cloacal prolapse.

Amphibians often carry rhabditiform nematodes, such as Rhabdias spp, the lungworm of anurans (frogs and toads). Frogs become infected percutaneously from third-stage larvae, and severe infections can cause morbidity and mortality in collections. In both cases, treatment is administration of anthelmintic agents to each animal and disinfection of the environment. The goal of treatment is not to completely eliminate the parasites but to decrease the parasite load to decrease morbidity and mortality in the population.

Vaccination is an essential part of any prevention program. Available vaccines have been shown to induce protective immunity against natural exposure in the select domestic species for which they are labeled. However, such protection has not been demonstrated for most nondomestic species in zoos. Therefore, vaccination is only one aspect of a holistic preventive-medicine program that should also include biosecurity and quarantine, integrated pest management, and vector control.

Vaccination schedules must be adapted to each situation, and recommendations to vaccinate or not should be on the basis of the relative risk of a specific infectious disease in a given collection. Caution should be exercised when using modified live vaccines (MLV) in species for which they are not labeled, because of the risk of reversion to virulence.

Recommendations for vaccinating specific taxa can be found in the management manuals and best practice guidelines published by the Association of Zoos and Aquariums and the European Association of Zoos and Aquariums, which are freely available. The decision to vaccinate zoo animals for less common diseases for which a vaccine is available should be made on an individual basis, with considerations for geographic area and disease risk.

All canids, procyonids, mustelids, and other carnivores (hyenas, bears, some felids, viverrids), peccaries, and some xenarthrans are considered susceptible to canine distemper virus. The use of recombinant, inactivated, or subunit vaccines against canine distemper virus is recommended in some susceptible zoo animals, although the induction and duration of immunity can vary among species and type of vaccine used.

Live attenuated vaccines often incite a greater immune response; however, they have been associated with clinical canine distemper infection related to reversion to virulence, and they are generally not recommended in zoo animals. Vaccination schedules follow those of domestic animals: after an initial series, annual revaccination is recommended. The vaccine can be used in pinnipeds to offer cross-protection against phocine distemper virus.

All canids are considered susceptible to canine adenovirus (CAV). Bears, skunks, mustelids, and procyonids can also be susceptible. CAV vaccination is considered a core vaccine in nondomestic canids. Unlike other modified live vaccines, CAV-2 vaccines have not shown reversion to virulence when used in nondomestic canids, with the exception of a case of CAV-2 hepatitis in a maned wolf (Chrysocyon brachyurus) (7).

Parvoviruses include canine parvovirus and feline panleukopenia virus, and most carnivores are susceptible to disease caused by one or both viruses. Canine parvovirus vaccination is generally recommended for canids, procyonids, and viverrids, with the exception of the red panda (Ailurus fulgens) (8).

Feline herpesvirus-1 causes severe disease in several exotic felids, including the cheetah (Acinonyx jubatus) and the Pallas's cat (Otocolobus manul), and feline calicivirus can cause life-threatening systemic infections in nondomestic felids. Feline panleukopenia, herpesvirus, and calicivirus are considered core vaccines for felids, and in all cases, the use of modified live vaccines is contraindicated because of risk of reversion to virulence.

Feline leukemia virus has been associated with lymphoma in cheetahs and with mortality in free-ranging populations of Iberian lynx (Lynx pardinus), puma (Puma concolor), and Florida panther (Puma concolor coryi). Most felids are considered susceptible to Chlamydia felis. Vaccines labeled for domestic cats are available for both of these diseases; however, infection in zoo animals is uncommon, and vaccination is not usually recommended.

Leptospirosisis caused by one of the many serovars of the spirochete Leptospira interrogans, and all mammals are susceptible. Prevention in zoos is achieved mainly by pest-rodent control and elimination of stagnant water. Vaccination limits disease severity and shedding by infected individuals. Vaccines are inactivated, multivalent, and approved for use in domestic dogs; however, it is not a core vaccine in zoo animal practice.

All mammals are susceptible torabies virus, and infection should always be considered fatal. Rabies is therefore of great zoonotic importance, especially in endemic areas. Marsupials are considered unlikely to be vectors because of their low body temperature, which makes it difficult for the virus to replicate.

Mesocarnivores, such as raccoons, foxes, jackals, or skunks, are common rabies vectors in wildlife communities and at the human–wildlife–domestic animal interface. The rabies vaccine is an inactivated vaccine administered annually to all carnivores and herbivores, including elephants, camelids, and ruminants, in endemic areas. Vaccination should also be considered for primates with access to outdoor enclosures in endemic areas and all mammals used in educational programs that are handled frequently or interface directly with the public.

Many Clostridium species have been associated with disease in mammals and birds, especially ruminants, equids, primates, mustelids, and waterfowl. Clostridium botulinum produces neurotoxins (called toxins A, B, C, D, E, F, and G) that commonly cause flaccid paralysis in affected animals. Macropods, elephants, and hoofstock (perissodactyls and artiodactyls) are vaccinated against clostridial toxins with commercially available multivalent vaccines labeled for use in domestic animals.

Herbivorous mammals and primates are considered especially susceptible to tetanus (Clostridium tetani), although all warm-blooded animals can be infected. Tetanus toxoid labeled for horses can be administered to susceptible, at-risk zoo animals as an initial series of two doses, 3–6 weeks apart, with revaccination annually. Elephants can be vaccinated at 3–4 months old, at 1 year old, and every 5 years thereafter. Nonhuman primates can be vaccinated with the product labeled for humans following similar protocols.

Canine parainfluenza virus can affect nondomesticated canids. However, it is not commonly observed in zoo animal practice. Many great apes are susceptible to human parainfluenza virus. No vaccine is available; therefore, prevention is primarily by biosecurity.

Many bird and mammal species are susceptible to influenza viruses. Influenza viruses are subject to antigenic drift and recombination, so the effectiveness of vaccination decreases over time, because naturally occurring strains do not always match those used in vaccines. Seasonal vaccination with the human product is generally recommended for great apes.

Equine influenza virus (EIV) causes respiratory disease in horses and nondomestic equids and can also infect humans and domestic carnivores. EIV vaccination is recommended for nondomestic equids before transport to other facilities. Vaccination of birds against avian influenza virus is currently not permitted in the US.

Erysipelas is the disease caused by Erysipelothrix rhusiopathiae (also known as Erysipelothrix insidiosa) in domestic pigs (Sus scrofa), prosimians, Old World and New World monkeys, cetaceans, and other marine mammals. Primates, cetaceans and suids can be vaccinated with the erysipelas vaccine labeled for use in domestic pigs.

Many nonhuman primates are susceptible to diseases for which vaccines have been developed for use in human medicine, including hepatitis B virus, measles, rubella, and pertussis. In these cases, vaccines are usually administered following schedules recommended for humans by the American Academy of Pediatrics and other professional associations for physicians.

Macaques (especially crab-eating macaques, Macaca fascicularis) and chimpanzees (Pan troglodytes) are susceptible to infection with hepatitis B virus. Although there are no documented cases of transmission between animals and humans, it is presumed to be a zoonotic disease. Seronegative primates can be vaccinated following the schedule recommended for humans.

Nonhuman primates in zoos are typically exposed to measles virus through contact with unvaccinated humans and ill primates, including great apes and Old and New World monkeys. Because of widespread human vaccination, nonhuman primates in zoos are not routinely vaccinated for measles. Policies can be reconsidered, however, in light of vaccine hesitancy among humans leading to measles outbreaks in human populations. The human vaccine can be used in nonhuman primates; however, caution is advised, because most measles vaccines contain live attenuated virus strains. The vaccination schedule begins at approximately 6 months old in most monkeys and at 15 months old in apes, following the guidelines for human vaccination.

Nonhuman primates can be susceptible to infection with rubella virus, like with measles, and exposure is generally achieved through contact with unvaccinated humans. However, incidence in human populations is low because of the widespread availability of vaccination and the use of personal protective equipment in zoos. When performed, primary vaccination of nonhuman primates is between 12 and 15 months old, with a booster administered at 4–6 years old.

Bordetella bronchiseptica is a well-known pathogen of domestic dogs and guinea pigs; however, in zoos it is of great concern for nonhuman primates and prosimians. Bordetella pertussis is the causative agent of pertussis in humans and has been described in captive chimpanzees. Great apes can be vaccinated against pertussis at 2, 4, and 6 months old, with revaccination 1 year later and every 10 years thereafter.

Streptococcus pneumoniae is the most common causative agent of meningitis in nonhuman primates (pneumococcal infection). Although not a core vaccine of great apes, the commercially available human vaccine can be used when indicated.

West Nile Virus (WNV) is a vector-borne flavivirus primarily transmitted by mosquitoes that first emerged in the US in 1999. It has since been detected in multiple orders of birds, mammals, and reptiles. Perissodactyls (including equids and rhinoceroses), marine mammals (harbor seals, Phoca vitulina, and killer whales, Orcinus orca), and many bird species (montane and non–African species, corvids, birds of prey, ratites, and penguins, among others) are particularly susceptible. Vaccination is with a monovalent WNV vaccine or with a polyvalent combination vaccine that includes Eastern and Western Equine Encephalitis viruses, to which many of these species are also susceptible. Duration of immunity is highly variable among species. Vaccination is recommended at least annually, prior to the start of the expected mosquito season (later winter or early autumn in temperate zones). Crocodilians are also susceptible to WNV, and a vaccine labeled for use in American alligators (Alligator mississippiensis) is commercially available. However, this is used mostly in the alligator ranching industry and is rarely used in zoos.

Postmortem examination is an essential part of zoo animal health programs. All mortalities must be recorded.

Postmortem examination includes gross examination (necropsy) and histological evaluation of tissues. At a minimum, signalment, suspected cause of death, and circumstances around death (eg, natural death versus humane euthanasia, whether an animal was under treatment at time of death) should be recorded.

Every collection animal that dies (or subsets of populations for colony-housed animals) should receive postmortem evaluation. Some exceptions include cases of severe autolysis that precludes diagnostic examination. Morbidity and mortality must be reviewed periodically in order to assess for trends, identify threats to animal health, and suggest ways to improve husbandry and preventive care of the collection.

When possible, biological samples, subsets of tissues, or slides for histological examination should be catalogued and banked for potential future examination. Evaluation of free-ranging wildlife found dead on zoo grounds should be considered also. This allows for surveillance of infectious and noninfectious diseases that can affect the health of collection animals, zoo staff, or guests. Given the limited existing knowledge regarding some species in zoo collections, complete postmortem examinations often add to the canon of literature on which to base evidence-based medicine and husbandry.

The health of the animals in a collection can be affected by both pests, which are often important vectors of infectious disease, and the methods used to control them. The objective of integrated pest management is to decrease the population of pests as effectively as possible with minimal risk to the zoo's animal collection. Pest-control programs should include both direct and indirect suppression methods.

Indirect suppression is considered the most important strategy for long-term pest control and works by eliminating pest attractants, such as sources of food and shelter. Examples include improving seals around doors, using tight-fitting lids for garbage bins, promptly removing trash from the premises, eliminating standing water, elevating feeding dishes or using exclusion devices to prevent pests from accessing the food of collection animals, and maintaining landscaping devoid of brush and organic litter.

Direct suppression includes exclusion of pests and the use of baits, traps, and pesticides. Of these, exclusion is the most important method, although the most difficult to implement. Baits, traps, and pesticides can be used when pest population is high or is suspected of carrying transmissible infectious diseases. Lethal traps are generally preferable to lethal baits, as the latter carry the risk of accidentally poisoning nontarget species.

In the absence of a qualified veterinary nutritionist or consultant, zoo veterinarians should oversee diet formulation and help manage, along with husbandry staff, diet preparation and delivery. Protocols should be developed for formulating diets, tracking changes to the diet, and monitoring all items that animals might consume, including foods that are used as rewards for training, foods offered as part of guest experiences, and foods that can be foraged on exhibit (eg, grass from pasture).

When formulating diets, needs specific to the species and to the individual must be considered. Software is commercially available to help ensure that diets are formulated to meet the nutritional needs of the animals. A holistic approach to nutrition is recommended, and animal body condition, diet consumption (or refusal), nutritional diseases or conditions, and the integration of diets into behavioral husbandry programs (eg, foraging enrichment, positive rewards for training) should be routinely evaluated. The Association of Zoos and Aquariums Nutrition Advisory Group is an excellent resource for clinicians working in zoo practice.

There are a number of nutritional diseases of zoo animals that clinicians should be familiar with:

Peracute mortality syndrome of giraffes is characterized by sudden death and believed to be associated with diets high in phosphorus, low in fiber, and high in concentrated feed. Clinical signs, when present, can include those typical of urinary tract obstruction observed in other ruminants, including pollakiuria and stranguria. On necropsy, affected animals typically exhibit poor body condition, serous atrophy of fat, and calcium carbonate urolithiasis with or without urinary obstruction.

The natural diet of the maned wolf (Chrysocyon brachyurus), a South American canid, is higher in fiber and lower in protein than that of most canids. Under human care, maned wolves are prone to inflammatory bowel disease (IBD) and cystinuria. Although it likely has a multifactorial etiology, diet plays a pronounced role in the pathogenesis of IBD. Hypoalbuminemia and secondary ascites can occur because of protein-losing enteropathy or maldigestion/malabsorption, and it is associated with a poor prognosis. Cystinuria has been observed in both wild and captive maned wolves.

The use of soy-based feed can improve fecal consistency and body condition. Surgery can be indicated for the removal of bladder stones or in cases of urinary obstruction. Medical treatment includes oral potassium citrate supplementation to alkalinize the urine, decreasing dietary sodium because it stimulates cysteine ​​excretion, and decreasing dietary animal protein because it contains more sulfur-containing amino acids. Administration of thiol-based drugs such as D-penicillamine can increase cysteine ​​solubility.

Iron storage disease (hemochromatosis) is an important nutritional disease in a number of zoo-housed species, including Sumatran and black rhinoceroses, tapirs, some frugivorous lemur species, dolphins (Tursiops truncatus), and some pinnipeds, as well as birds of the families Ramphastidae (eg, toucans, toucanets, and aracaris) and Bucerotidae (Asian hornbills), mynah birds, and some species of starlings, among others.

Ferroportin is a transmembrane protein that allows dietary iron from meat (heme iron) and plants (nonheme iron) to enter and exit cells. It is expressed in hepatocytes and enterocytes, making the liver the first organ affected in cases of iron overload. Within the cytoplasm, iron binds to ferritin for storage. It is transported throughout the body bound to transferrin in the bloodstream. Hepcidin is an acute-phase protein found in fish, herpetofauna, and mammals that mediates anemia associated with chronic disease. It is released in response to increased iron storage and decreases ferroportin production to decrease its absorption and facilitate its elimination through the natural turnover of enterocytes.

Clinical signs of iron storage disease are often associated with liver failure and can include ascites, weight loss, hepatomegaly, icterus, and anorexia. Diagnosis is challenging. Serum ferritin and transferrin saturation percentage can be used to diagnose and monitor the disease in some mammal species. In birds and many mammal species, a liver biopsy is necessary for antemortem diagnosis, although MRI of the liver has also been used.

Prevention of iron storage disease begins with dietary modification, limiting the intake of iron as well as of vitamin C and acidic foods that promote the uptake of dietary iron. Tannic acid (tannins) can be added to the diet of frugivorous animals to limit iron absorption from the diet. The primary treatment involves therapeutic phlebotomy. Iron chelators, such as parenteral deferoxamine or oral deferiprone, are being studied in rhinoceroses and other species.

Veterinarians should review nutritional supplementation of all taxa, with particular attention to marine mammals and piscivorous animals in general. Many fish contain thiaminase in their tissues, which metabolizes vitamin B1 over time. Fresh frozen fish is recommended for piscivorous animals; however, the diet should be supplemented with vitamin B1 (thiamine), nonetheless.

Vitamin E is an antioxidant that prevents the oxidation of fats that causes rancidity. Given the high lipid content of piscivorous diets, these should also be supplemented with vitamin E. Vitamin E deficiency can lead to steatitis or fat necrosis, as has been described in many piscivorous species under human care, including teleosts and crocodilians (9).

Cetaceans, like primates, guinea pigs, and bony and cartilaginous fish, cannot produce endogenous vitamin C, and dietary supplementation is recommended. Updated recommendations for handling fish fed to piscivorous animals can be found through the Association of Zoos and Aquariums Nutrition Advisory Group.

• Nutrition Advisory Group. Association of Zoos and Aquariums.

• Maslanka M, Watts J, Bissell H, et al. Handling Fish Fed to Fish-Eating Animals. 2nd ed. Association of Zoos and Aquariums Nutrition Advisory Group. 2023.

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