Wasting Syndrome in a Bull African Elephant (Loxodonta africana)
American Association of Zoo Veterinarians Conference 2002
Karen A. Emanuelson1, DVM; Dalen W. Agnew2, DVM
1Oakland Zoo, Oakland, CA, USA; 2Department of Pathology, Microbiology, and Immunology, School of Veterinary Medicine, University of California, Davis, CA, USA


In May 2000, Oakland Zoo’s 28-year-old breeding bull African elephant (Loxodonta africana) was evaluated for mild diarrhea of four days’ duration. Historically, the bull had few, relatively minor, medical problems, including a low serum vitamin E level in 1998. At the current presentation, the physical examination was within normal limits, and body weight was adequate (7590 kg). In the following 7 months, the bull continued to have diarrhea and abdominal distention of varying severity, with a consistent, normal appetite. Diarrhea ranged from barely softened fecal boluses to severe hematochezia and liquefied feces with colic. Diagnostic testing performed during this time was extensive. Weekly complete blood counts and serum chemistries, and urinalyses were completed. Persistent mild anemia, hyponatremia and hypochloremia were the only abnormalities found. Additionally, feed analysis, over 30 successive negative stool cultures for Salmonella, repeated negative stool cultures for other pathogens such as Clostridium, Campylobacter, and other enterics, including E. coli (serotyped), as well as Mycobacterium paratuberculosis and M. tuberculosis were completed and did not provide an etiologic agent. Serial fecal parasite evaluations, fecal electron microscopy for viruses, and serum tests for Leptospira sp., malignant catarrhal fever, bovine viral diarrhea, bluetongue, and infectious bovine rhinotracheitis, Ehrlichia risticii were also negative and no abnormalities of heavy metals (copper, magnesium, lead, iron, selenium, zinc) were found. Lastly, feces evaluated for E. risticii by PCR were found negative and rectal biopsies collected by endoscopy were non-diagnostic. Fecal occult blood and cytology were routinely performed during the illness and were abnormal only during the period of hematochezia and liquefied feces.

Treatments in the first few months consisted primarily of feed exclusion trials, when soft feces and minor abdominal distension were the only clinical signs. Produce, concentrates (Mazuri Elephant Pellets without vitamin E, PMI Nutritional International Inc., Brentwood, MO, USA), and browse were eliminated from the diet, reducing the diet to hay only; produce was eliminated immediately followed by gradual reduction of browse and concentrates for over 6 months. Hay types were then changed to try Sudan, oat, wheat, and meadow grass; feeding each type of hay lasted approximately 4–6 weeks. Feeding frequency was increased to every 20–60 minutes during the day, with two night feedings, to decrease the volume of each meal. This final method of feeding did reduce the diarrhea slightly. After several months, browse then concentrates were slowly added back into the diet, as their elimination had no beneficial effect.

Two sudden, severe exacerbations of clinical signs occurred in the third and fifth month of the illness. In August 2000, the bull developed a sudden onset of bloody diarrhea, tenesmus, extreme bilateral bloat, anorexia, adipsia, weakness, and lethargy. Parenteral treatment with amikacin (Butler Co., Columbus, OH, USA; 5 mg/kg SID IM for 10 days) and ceftiofur (Naxcel, The Upjohn Co., Kalamazoo, MI, USA; 1 mg/kg SID IM for 10 days) resolved clinical signs in eight days. However, the diarrhea slowly returned after cessation of treatment. During the fourth month, treatments included a course of rectally administered metronidazole (500 mg tablets, 15 mg/kg rectally SID for 10 days); ivermectin (Eqvalan paste, Merck Sharp and Dohme Quimica de Puerto Rico Inc., Barceloneta, Puerto Rico, 0.065 mg/kg orally once); fenbendazole (Panacur granules, Global Pharmaceuticals Inc., Don Mills, ON, Canada, 5 mg/kg orally SID for 5 days); lactobacillus equine supplements (Probios gel, CHR Hansen Biosystems, Milwaukee, WI, USA); elephant fecal supplements; continued salt supplementation, and frequent feeding and watering. In October, a sudden onset of watery diarrhea and weakness occurred. The elephant was again treated with parenteral amikacin (6 mg/kg SID IV for 14 days, with the dose administered IM on day 13 and 14) and ceftiofur (2.2 mg/kg SID IM for 14 days). Treatment also included ranitidine (Zantac, Glaxo Wellcome, Research Triangle Park, NC, USA, 0.5 mg/kg orally BID for 14 days), and flunixin meglumine (Banamine paste, Schering-Plough Animal Health Corp., Union, NJ, USA, 0.5–1.1 mg/kg orally SID to BID for 6 days). The weakness resolved, but diarrhea continued.

In the ninth month, January 2001, the bull’s clinical signs progressed to include ventral peripheral edema, multiple areas of dermatologic pustules, lethargy, reluctance to move, weight loss, episcleritis, and oral ulcerations. Due to the noted weight loss, concentrates and hay were increased in the diet. In February 2001, endoscopic intestinal biopsy provided little informative histopathology. Due to progression of disease and poor response to the second course of antimicrobials, oral prednisone was initiated on March 10; however, only a single oral dose was administered. On March 11, 10 months after the onset of illness, the elephant collapsed. Despite supportive treatment for shock and assisted lifting with a crane, the elephant could not rise, and died 15 hours after onset of recumbency.


The cause of chronic diarrhea, debilitation, and, ultimately, death, in this middle-aged bull African elephant was not determined after extensive clinical diagnostics and a complete necropsy. Pronounced pigmentation was noted in the liver, kidney, and brain, and to a lesser extent in the heart, skeletal muscle, and mesenteric ganglia. This unusual pigment was partially PAS-positive, slightly acid-fast, and possessed autofluorescence, similar to lipofuscin. It was negative with iron, copper, bile, and melanin stains. By transmission electron microscopy, hepatocytes were packed with mitochondria often containing lipid and electron dense bodies, and other membrane-bound electron dense bodies were consistent with lipofuscin.1 These lipofuscin aggregates appeared the result of terminal mitochondrial degeneration. Lipofuscin pigments are seen in aging animals of many species by post-mitotic cell lysosomes accumulating cellular materials of autophagocytosis.11 It is commonly reported in large amounts within the liver, brain, heart, and skeletal muscle of captive aged elephants.5

Lipofuscin accumulation in the brain or liver or both may also be associated with genetic enzyme deficiencies, such as those seen in South Hampshire sheep or Devon cattle.8,9 Alternatively, it may be seen in association with plant intoxications, such as Gomen disease in horses or Acacia aneura, Trachyandra divaricata, or Trachyandra laxa in sheep, goats, horses, and pigs.4 T. divaricata and T. laxa also produce marked lipofuscinosis in the renal cortex and lymph nodes. Interestingly, the degree of histologic neuroaxonal damage noted in sheep and horses intoxicated with T. divaricata and T. laxa is not consistent with the degree of clinical signs. This suggests some unrecognized sublethal effect, leading to the marked lipofuscinosis and neurologic disease.3,7,10 In this elephant, the degree of lipofuscinosis in the liver, kidney, and brain far exceeds that seen even in considerably older elephants (R.J. Montali, personal communication). Genetic or enzyme mutations have not been reported in elephants, this animal was wild caught, and presumed not inbred, so genetic causes are considered unlikely. Investigations into its diet reveal that it had consumed large amounts of browse offered by the zoo for behavioral enrichment and more balanced nutrition. This browse was mixed, but contained large amounts of Acacia melanoxylon, Acacia dealbata, and Chinese elm (Ulmus parvifolia). Reports of toxicity from these plants are limited, but A. melanoxylon does contain polyphenols, which may reduce protein and nitrogen digestion and assimilation. Other Acacia sp. have been associated with posterior paralysis, but not lipofuscinosis.2,6 Lipofuscin deposition is considered to be a result of oxidative damage accumulated over a period of time. This elephant was reportedly deficient in vitamin E 2 years earlier, but its diet had been successfully corrected long before clinical signs began. It is possible, however, that lipofuscin had accumulated during the deficient period. Vitamin E deficiency has, however, been reported in mammals, without observation of similar clinical signs or pathologic lesions.

A definitive etiology for this elephant’s marked lipofuscinosis is currently unknown. Archived necropsy materials from African and Asian elephants with similar clinical signs may provide additional etiologic information once reviewed.


We would like to thank Dr. Linda Lowenstine, Dr. Dick Montali, Dr. Freeland Dunker, Dr. Gary Magdesian, Dr. Larry Gallupo, Dr. Jim Oosterhuis, and Dr. Birgit Puschner for their assistance and interest in the case; the elephant care staff at the Oakland Zoo for their countless hours of effort on this animal’s behalf; as well as the elephant himself, for his greatness and stoicism throughout the ordeal.

Literature Cited

1.  Cheville N.F. 1994. Ultrastructural Pathology: An Introduction to Interpretation. Iowa State University Press, Ames, Iowa. Pp. 377–335.

2.  Clement B.A., C.M. Goff, and T.D.A. Forbes. 1997. Toxic amines and alkaloids from Acacia berlandieri. Phytochem. 46: 249–254.

3.  Huxtable C.R. 1987. Neurological disease and lipofuscinosis in horses and sheep grazing Trachyandra divaricata (branched onion weed) in south Western Australia. Aust. Vet. J. 64:105–108.

4.  Le Gonidec,G. 1981. Letters to the editor: a neurologic disease of horses in New Caledonia. Aust. Vet. J. 57:194–195.

5.  McGavin, M.D., R.D. Walker, E.C. Schroeder, C.S. Patton, and M.D. McCracken. 1983. Death of an African elephant from probable toxemia attributed to chronic pulpitis. J. Am. Vet. Med. Assoc. 183:1269–1273.

6.  Nantoume H., T.D.A. Forbes, C.M. Hensarling, and S.S. Sieckenius. 2001. Nutritive value and palatability of guajillo (Acacia berlandieri) as a component of goat diets. Small Ruminant Res. 40:139–148.

7.  Newsholme, S.J. 1985. A suspected lipofuscin storage disease of sheep associated with ingestion of the plant,

8.  Trachyandra divaricata (Jacq.) Kunth. Onderstopoort J. Vet. Res. 52: 87–92.

9.  Nordstoga, K. 1990. Hepatic lipofuscinosis in healthy Norwegian sheep. Acta Vet. Scand. 31:73–78.

10.  Read, W.K., and C.H. Bridges. 1968. Neuronal lipodystrophy: occurrence in an inbred strain of cattle. Vet. Pathol. 6:235–243.

11.  Winter H. 1966. Autofluorescence in hepatic lipofuscinosis of sheep. Aust. Vet. J. 42:40–41.

12.  Yin, D. 1996. Biochemical basis of lipofuscin, ceroid, and age pigment-like fluorophores. Free Radical Biol. Med. 21:871–888.


Speaker Information
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Karen A. Emanuelson, DVM
Oakland Zoo
Oakland, CA, USA

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