Description of a surgical repair of a perineal hernia in an Asian elephant in Myanmar. Oo Z.M., et al. 2016 Surgical treatment of a cervico-vaginal prolapse in an Asian elephant in Myanmar. Gajah 44, 36-39 To perineal hernia Case report Perineal hernia, surgical repair Date: 2016 Place: Myanmar Data provided by: Gajah History A 47 years old captive female Asian elephant, working in the logging industry, displayed a large bulging mass below the tail since 10 years, which increased in diameter from 4 inches to 22 inches. At ultrasonographic examination it was diagnosed as a cervico-vaginal prolaps. Better described as perineal hernia with cervico-vaginal involvement (WS). The elephant did not suffer of any limitations in relationship to this condition. Previous cases in other female elephants of the company had died of this condition. As the mass was increasing in size, it was decided to perform a correcting surgery. At the time of surgery the animal was in healthy condition. Read the article in Gajah Treatment A standing sedation procedure with xylazine and ketamine was used. A incision of the skin and the vaginal vestibule was made 7 inches lateral to the perineal midline, allowing manual passage to explore the subcutaneous area. A catheter (1/4 inch diameter) was advanced into the uterus (or was it into the urthra?) and the uterus (bladder?) could be drained. The catheter was replaced by a larger catherter (½ inch) and more fluid, which contained several stones was drained. The the cervix uteri and the vagina (bladder?) were pushed back into the p elvic cavity through the herniated pelvic diafragm using the arm that was advanced into the vagina . The vaginal vestibule was closed using a continuous catgut suture. The skin was closed using 3 continuous nylon matrass sutures. At each knot a protecting plastic plate was placed underneath the knot to protect the skin from perforation by the nylon suture. Finally, a wooden block with a foamy protection layer was tightly tied to the (formerly) bulging area using ropes around the elephants body. Treatment results The ropes and the supporting block remained in place and the hernia did not recur between surgery and publication of this paper. However, it is not know for how long this animal stayed without recurrence. Complete report: Oo Z.M., et al. 2016 Surgical treatment of a cervico-vaginal prolapse in an Asian elephant in Myanmar. Gajah 44, 36-39 To page top

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Prevalence, symptoms, treatment and vaccination against anthrax (Bacillus anthracis) in elephants are described. To infectious diseases Anthrax This figure gives a nice overview of the epidemiological cycles of Bacillus anthracis (https://anipedia.org/resources/anthrax/1203 ). Anthrax infection in humans Human anthrax infections are often contracted during work activities in oneofthe following fields: Tanneries Wool sorters Bone processors Slaughterhouses Laboratory workers When humans become infected, the disease is usually presented as skin wounds that heal very slowly. The bacteria can penetrate the skin if they come in contact with a fresh skin wound. These photos demonstrate the type of wound that results from such an infection. The person with the wound on the left image worked on a cattle carcass that died from anthrax. While handling this carcass, he injured himself by a sharp bone fragment that was infected with the anthrax bacteria ( https://www.microbiologybook.org/ghaffar/anthrax-pennsylvania.htm ). The person on the right image is probably a tannery worker, who infected himself by rubbing his knuckles on the skin of an animal that died of anthrax ( http://www.fao.org/ag/againfo/programmes/en/empres/news_070212.html ) If untreated or if the infected wound is big, the bacteria can spread in a large area around the wound, as is shown here. This severe wound needs immediate treatment with the right antibiotic. Anthrax spores can also be inhaled. In the lungs the anthrax bacteria can cause a very severe inflammation. On the left image you can see an X-ray of healthy lungs, with a clear heart shadow. On the right X-ray you can distinguish a big mass in the thorax that does not allow the x-rays to get through. If this disease is left untreated until obvious symptoms occur, it is usually fatal. This patient had a business where he made drums using cattle and goat skins imported from Africa. He died 1 day after this X-ray was made. (https://www.microbiologybook.org/ghaffar/anthrax-pennsylvania.htm ) Anthrax infection in animals Typically, the incubation period is 3–7 days (range 1−14 days). The clinical course ranges from peracute to chronic. The peracute form (common in cattle and sheep) is characterized by sudden onset and a rapidly fatal course. Staggering, dyspnea, trembling, collapse, a few convulsive movements, and death may occur in cattle, sheep, or goats with only a brief evidence of illness. Often, the course of disease is so rapid that illness is not observed and animals are found dead. A very characteristic feature of acute anthrax is free non-coagulating blood running from body openings, due to the disturbed coagulation. The disease in horses may be acute. Signs may include fever, chills, severe colic, anorexia, depression, weakness, bloody diarrhea, and swellings of the neck, sternum, lower abdomen, and external genitalia. Death usually occurs within 2–3 days of onset. Anthrax bacteria disturb the natural blood coagulation. This results in bleedings in the skin and all internal organs. Non-coagulated blood is collected in the lymph nodes, while free-running blood appears from all openings. These symptoms usually lead to a sudden death. Anthrax can affect multiple species, like cattle and wild ruminants (greater kudu), zebras as well as predators (lion). Note the running blood from nostrils or eyes in all these animals and the small bleedings in the skin of the kudu. (https://anipedia.org/resources/1203) Multiple outbreaks of anthrax in wild hippopothamus has been reported in several southern African countries ( https://www.sciencealert.com/anthrax-outbreak-suspected-to-have-killed-more-than-100-hippos-in-namibia ) Animal to animal transmission There are several ways of transmission of B. anthracis between animals. Animals grazing in areas where anthrax victims have been buried, can be infected when the carcass remnants are digged up either by the feeding animal or through human activities (road or building constructions). Flies that have fed on an anthrax-carcass can easily spread the bacteria through their droppings that remain on leaves. Predators (big cats) that feed on infected carcasses can become infected and die of anthrax. Flies that feed on an infected carcass may spread B.anthacis through their droppings as illustrated on these images (https://anipedia.org ). Anthrax in elephants Anthrax in elephants is usually a gastro-intestinal infection. The animal ingests the spores while feeding food or water contaminated with spores. After an incubation period of a few days, the animal dies of septicemia. Multiple cases have been reported from several range countries in Asia (Kumaraguru A. et al. 2015). In some areas Asian elephants play a role in the transmission of anthrax between wildlife and farm animals (Walsh M.G. et al. 2019). Like in other mammals, symptoms consist of rapid detoriation after infection. Usually the elephant is found dead before symptoms were observed. Running blood from the trunk, mouth, eyes, rectum or vagina should alarm the finder of the carcass for this being an anthrax case. Anti-PA antibodies were detected in elephants, which suggests that they can mount adaptive immune responses against anthrax. In addition, these results suggest that elephants can be infected with anthrax and survive infection under some circumstances (Cizauskas et al. 2014). A fatal case of anthrax in a 15-yrs-old African elephant was reported from a wildlife park in Nigeria (Okewole, 1993). Frequent urination, restlessness and weakeness of the hind quarters were observed prior to death. A co-infection with Cowdria ruminatium was diagnosed at post mortem examination (coccoid intracytoplasmatic bodies in the endothelia of the brain). A wild Asian elephant that died of anthrax in the forest of Myanmar. Note the amount of free running blood around the head. Photo courtesy: Myanmar Forest Police A wild Asian elephant that died of anthrax in the forest in India (Kumaraguru A. 2015) Treatment Early detection of the disease is essential, though difficult. Multiple classes of antibiotics can be used if treatment is started in time: oxytetracycline, penicillins, aminoglycosides, fluoroquinolones, macrolides, and sulfonamides. Dosages can be obtained from the website of Elephant Care International: https://www.elephantcare.org/formulary Diagnosis (post-mortem) Post mortem findings in elephants are: Bleedings In and under the skin Around muscles In organs Free blood in the intestines Free blood in the lungs Free blood in the abdomen Edematous swelling of the skin Swollen spleen with bleedings Liver and lymph nodes are swollen and contain a lot of blood Disposal of an anthrax carcass If a dead elephant is suspected of anthrax, a full necropsy is not recommended. A blood smear from a small incision made in an ear should first be made and examined microscopically for the presence of Gram-positive stained rods, lying in chains, sometimes accompanied by spores. The carcass should be disposed off as soon as possible in a proper way. The disposal must be done following the next steps: To minimize the spread of blood, you should try to plug the openings (trunk, ears, mouth, anus, vulva) with non-absorbent material. You can also wrap the head of the elephant in plastic and tape it with duct tape or tie it with a rope to the skin of the neck. Don’t move the animal around Incinerate the carcass if possible If incineration is not possible, burry the carcass as deep as possible. Use heavy excavating equipment (backhoe loader) to dig a deep, large hole, at least 2 meters deep Disinfect all materials that have been in contact: 10% formalin or 5% lime solution (sodium hydroxide) Necropsies of any animal should always be performed with great care. If there are signs of anthrax, a peripheral blood smear should always be examined first. If accidently the diagnose was missed, any signs of internal bleedings should alarm the prosector. B. anthacis can be cultured quite easily. Every necropsy should be performed with adequate body protection: proper eye protection, a respiration mask, long gloves, rubber boots and protective clothing. Vaccination Elephants can be vaccinated against anthrax with commercially available vaccines. This is highly recommended in areas where anthrax is seen in farm animals or if there is a history of anthrax in elephants in that area. References/further reading Berry HH. 1993. Surveillance and control of anthrax and rabies in wild herbivores and carnivores in Namibia. Rev Sci Tech 12(1):137–146.Cizauskas CA, Bellan SE, Turner WC, Vance RE, Getz WM. 2014. Frequent and seasonally variable sublethal anthrax infections are accompanied by short-lived immunity in an endemic system. J Anim Ecol 83(5):1078–1090 Hanna P., 1998. Anthrax pathogenesis and host response. Curr Top Microbiol Immunol 225:13–35 Turnbull PC, Bell RH, Saigawa K, Munyenyembe FE, Mulenga CK, Makala LH. 1991. Anthrax in wildlife in the Luangwa Valley, Zambia. Vet Rec 128(17):399–403. Kumaraguru A., Kumaraguru Arumugam , N.S. Manoharan , Ramakrishnan Balasundaram . 2015. Prevalence and disease management with reference to anthrax in the Asian elephant (Elephas maximus) in the Sathyamangalam Wildlife Santuary, Tamil Nadu, India & Indash; A case study. Scientific Transactions in Environment and Technovation, 5(1): 48-51. Okewole P.A., Oyetunde I.L., Irikanulo E.A., Chima J.C., Nwankpa N., Laleye Y., Bot C. 1993. Anthrax and cowdriosis in an African elephant (Loxodonta africana). Walsh, M.G., Mor, S.M., Hossain, S., 2019. The elephant–livestock interface modulates anthrax suitability in India. Proceedings of the Royal Society B: Biological Sciences 286 EAZWV Transmissible Disease Fact Sheet ANTHRAX American Association of Zoo Veterinarians Infectious Disease Manual ANTHRAX Recommended websites: Merck Veterinary Manual. 2021. https://www.merckvetmanual.com/generalized-conditions/anthrax/overview-of-anthrax OIE (Organization for Animal Health: https://anipedia.org/resources/anthrax/1203 FAO: http://www.fao.org/home/search/en/?q=anthrax Microbiology and Immunology On-line: https://www.microbiologybook.org/ghaffar/anthrax-pennsylvania.htm To page top General information Merck Veterinary Manual (2021) : Anthrax is a zoonotic disease caused by the sporeforming bacterium Bacillus anthracis . Anthrax is most common in wild and domestic herbivores (eg, cattle, sheep, goats, camels, antelopes) but can also be seen in people exposed to tissue from infected animals, to contaminated animal products, or directly to B anthracis spores under certain conditions. Depending on the route of infection, host factors, and potentially strain-specific factors, anthrax can have several different clinical presentations. In herbivores, anthrax commonly presents as an acute septicemia with a high fatality rate, often accompanied by hemorrhagic lymphadenitis. In dogs, people, horses, and pigs, it is usually less acute although still potentially fatal. Toxins are the source of most of the disease symptoms associated with anthrax. Edema toxin complex (EdTx) causes the fluid and edema seen in cutaneous anthrax infections, and lethal toxin complex (LeTx) causes shock and death from systemic anthrax (Hanna, 1998). B anthracis spores can remain viable in soil for many years. During this time, they are a potential source of infection for grazing livestock but generally do not represent a direct risk of infection for people. Grazing animals may become infected when they ingest sufficient quantities of these spores from the soil. In addition to direct transmission, biting flies may mechanically transmit B anthracis spores from one animal to another. The latter follows when there have been rains encouraging a high fly hatch and reporting has been delayed on the index ranch, such that there are 4–6 moribund or dead cattle for the flies to feed on. Feed contaminated with bone or other meal from infected animals can serve as a source of infection for livestock, as can hay muddy with contaminated soil. Raw or poorly cooked contaminated meat is a source of infection for zoo carnivores and omnivores; anthrax resulting from contaminated meat consumption has been reported in pigs, dogs, cats, mink, wild carnivores, and people. Human cases may follow contact with contaminated carcasses or animal products (raw meat, skins of animals that died of anthrax). Flies that have fed on a carcass from an anthrax victim can spread the disease over longer distances. Diagnosis Anthrax can be diagnosed in fresh blood smears taken from the ear. Microscopically, B. anthracis can be recognized as long chains of Gram-positive bacteria. If the smear has been exposed to air, the bacteria may have formed spores that can be easily detected. B. anthracis differs in shape from other Bacillus species, that may contaminate the sample in case the animal has been dead for a longer period. Whereas the bacteria chains of B. anthracis seem to be sharply cut off with a knife, the chains of B. cereus have round edges.

Rabies has been diagnosed in a few elephants and was fatal in all reported cases. The source of the infection was attributed to canids (Wimalaratne et al. 1999, Nanayakkara et al. 2003, Sharma et al. 2005, Aravind et al. 2006). The incubation time for rabies in elephants is unknown. Depending on the distance between the bite wound and the elephant's brain, a long incubation period can be expected. Rabies should be included in the differential diagnosis whenever there are neurological signs. To infectious diseases Rabies General information Rabies is a viral disease that is usually fatal. It is caused by a neurotropic Lyssa virus. Several species of Lyssaviruses have been identified, of which the rabies virus (worldwide), the Mokola virus (Africa), the Duvenhage virus (South Africa) and the European and Australian bat lyssaviruses are responsible for fatal encephalomyelitis. Rabies is transmitted by a percutaneous bite from a rabies-infected animal or by wound contamination with saliva from a rabid animal (MSD, 2021). Air born infection through aerosols can occur when visiting bat caves. Reservoir hosts vary geographically. In the U.S. hosts include skunks, bats, raccoons, foxes, and coyotes. Civets, mongooses and hyenas are the main hosts in Africa; domestic dogs are hosts in Asia, South America, and Africa. Certain bat species in southern Africa are host for a Lyssa virus After infection of the bite wound, the virus migrates from the wound to the brain via the nerves that run from the wound area, resulting in neurological signs. Virus replication takes place in the brain, from where the virus migrates to the salivary glands. The incubation period varies from 3 weeks to many months. In most species affected by rabies, the animal shows an increase of aggressive behaviour and will try to bite other animals. Hydrophobia can be one of the accompanying symptoms. In a few species (e.g. domestic cat), the animal becomes more quite or even soporous. Once the virus has reached the brain, there is no cure. Treatment is only possible in the short period between infection and start of the migration. In this short time window, the animal should be treated daily with a rabies vaccine (described below). If available, locally anti-rabies serum should be injected around the wound area. Diagnose of rabies is based on PCR or histopathology of brain tissue by demonstrating the presence of typical Negri bodies using a special stain especially in pyramidal cells within the Ammon's horn of the hippocampus. Wound tissue, saliva and cerebrospinal fluid can be used when the animal is still alive (CDC, 2021). Rabies in elephants Rabies has been diagnosed in a few elephants and was fatal in all reported cases. The source of the infection was attributed to canids ( Wimalaratne et al. 1999, Nanayakkara et al. 2003, Sharma et al. 2005, Aravind et al. 2006). The incubation time for rabies in elephants is unknown. Depending on the distance between the bite wound and the elephant's brain, a long incubation period can be expected. Rabies should be included in the differential diagnosis whenever there are neurological signs. The initial signs of rabies in elephants may be vague but most often the elephant (FAO 2005): Is listless. Prefers to stay in dark places. Eats very little. As the disease progresses the elephant likely: Writhes in pain. Does not recognize the mahout. Chases and attacks humans and animals. Has eyes that roll and wander. Does not eat. Walks unsteadily and the legs lose strength. Goes to the ground in paralysis. Has locked jaws and the tail hangs still. Has saliva flowing continuously. Death may shortly follow the appearance of these more severe signs. The differential diagnose in case of rabies comprises any disease that can cause central nervous system symptoms, like: Tetanus Trauma Snakebite Toxicity (e.g. heavy metal; pesticide) Anytime an elephant is bitten, particularly if the bite has drawn blood, the mahout and owner should take four actions: 1. Write the day on a calendar; then you will be able to predict when the elephant may show clinical signs if it was infected. 2. Talk to people who know the dog and ask about its behaviour in the days prior to the attack; if the dog has been acting strangely (staring fixedly, foaming at the mouth, etc.) there is a good chance the dog is rabid. 3. Inform everybody in the community of the health hazard, because the disease also attacks humans, and ask them to help track down the dog. 4. Very carefully capture the dog, confine it securely, and observe its condition; if after ten days it is normal then the elephant does not have rabies. If the elephant dies, consult Disposal of carcasses, page 55 of the FAO manual . Treatment of rabies in elephants Immediately after a suspected dog bite, wash the wound intensively with soap and water. Then apply tincture of iodine or Povidone-iodine 1% in and around the wound (FAO 2005). There is no effective treatment once the symptoms have appeared. Even though the disease is not contagious to other elephants, separate the elephant, taking it to a shady, clean and quiet place. Make sure the elephant is chained tightly and securely (FAO 2005). Although there is no report on post-exposure vaccination, emergency vaccination of the elephant can be considered, if a rabies vaccine is available. One study describes the successful post-exposure treatment in pigs that where heavily exposed to rabies. The affected pigs were repeatedly vaccinated with an inactivated rabies vaccine 0, 3, 7, 14 and 30 days after the bite incident, while equine rabies immune globulins were injected in and around the bite wound (Mitmoonpitak et al. 2002). When an elephant is exposed to rabies, consider to inject the animal intramuscularly with a 2 ml dose of an (inactivated = killed) rabies vaccine as soon as possible after it was bitten by a rabid animal. These vaccinations should to be given daily for at least 5 days. During this period the elephant should be kept under close observation. Anytime an elephant is bitten, particularly if the bite has drawn blood, the mahout and owner should take four actions: Write the day on a calendar; then you will be able to predict when the elephant may show clinical signs if it was infected. Talk to people who know the dog and ask about its behaviour in the days prior to the attack; if the dog has been acting strangely (staring fixedly, foaming at the mouth, etc.) there is a good chance the dog is rabid. Inform everybody in the community of the health hazard, because the disease also attacks humans, and ask them to help track down the dog. Very carefully capture the dog, confine it securely, and observe its condition; if after ten days it is normal then the elephant does not have rabies. If the elephant dies, consult Disposal of carcasses, page 55 of the FAO manual . Prevention Regular rabies vaccination is recommended for all elephants kept under human care in areas where rabies is endemic. Because rabies is incurable the best prevention is to annually vaccinate all the dogs and cats in the community. For many years, following the recommendation for rabies vaccination in horses has been considered prudent: (inactivated!) vaccine (2 ml IM) given from the age of 6 months, to be repeated after 3-4 weeks and annually boostered. When using this vaccination schedule in elephants, antibodies against rabies could be demonstrated after 24 months (Isaza et al. 2006, Miller et al. 2009). However, this rabies vaccination strategy was evaluated in a herd of 9 African elephants, including two calves, four subadults, and three adults which lead to new conclusions about rabies vaccination strategy. Prior to 2017, elephants were vaccinated opportunistically IM. Starting in 2018, calves at least 4 months of age were administered 2 ml of a commercially available inactivated vaccine and received boosters at 1 y of age. Adults and subadults underwent annual vaccination at the same dose. After 1 year, neutralization titers in five of nine elephants were below levels considered protective in domestic animals (< 0.5 IU/ml). Therefore the dose of rabies vaccine was increased to 4 ml, which resulted in titers more consistently greater than or equal to 0.5 IU/ml for at least 6 months. Institutions with elephants under human care may consider performing rabies vaccination neutralizing titers when possible to help guide vaccination. See also: vaccination. References Aravind B., Anilkumar M., Raju S., and Saseendranath M.R. 2006. A case of rabies in an Indian elephant (Elephas maximus) . Zoo's print journal 21 (2) 2170. Browning G.R., Peters R., and Howard L.L. 2021. Rabies vaccination and antibody response in African elephants ( Loxodonta africana ) as part of a comprehensive program of veterinary care. Joint AAZV EAZWV Conference Proceedings 2021. CDC 2021: https://www.cdc.gov/rabies/diagnosis/animals-humans.html FAO 2005: Elephant care manual for mahouts and camp managers. 2005. Isaza R., Davis R.D., Moore S.M., and Briggs D.J. 2006. Results of vaccinat i on of Asian elephants (Elephas maximus) with monovalent inactivated rabies vaccine. AJVR, Vol 67 (11), 1934-1936, 2006 Miller M.A. and Olea-Popelka F. 2009. Serum antibody titers following routine rabies vaccination in African elephants. JAVMA, Vol 235 (8),978-981 2009 Mitmoonpitak C., Limusanno S., Khawplod P., Tepsumethanon V, and Wilde H. 2002. Post-exposure rabies treatment in pigs. Vaccine 20 (2002) 2019–2021. MSD, 2021: https://www.msdmanuals.com/home/brain,-spinal-cord,-and-nerve-disorders/brain-infections/rabies Nanayakkara S, Jean S. Smith, and Charles E. Rupprecht. 2003. Rabies in Sri Lanka: Splendid Isolation. Emerging Infectious Diseases • Vol. 9, No. 3, March 2003. Sharma A.K., Choudhury B, and Singh K.P. 2005. Rabies in a captive elephant . Indian Journal of Veterinary Pathology 29(2): 125-126 Wimalaratne O, and Kodikara D.S. 1999. First reported case of elephant rabies in Sri Lanka. Vet. Rec. 144 (4): 98. To page top

Esophageal spasms are rarely seen in elephants. This case reports describes this condition in an Asian elephant. Water regurgitation indicated the blockage of water, while the animal was unable to swallow any food. A home-made endoscope greatly facilitated the visualization of the esophagus and stomach wall. A standing sedation using detomidine and butorphanol was used during the treatment procedure. No mouth gag was needed to open the mouth. Continue To non-infectious diseases Case report Esophagus spasm Place: Selwo Zoo, Spain Date: 2019 Data provided by: Cecilia Sierra Arqueros, DVM History Species: Asian elephant Accommodation: Zoo Age, gender: 54 years, female For several years this female Asian elephant had episodes of rhythmic contractions in the ventral area of the neck at the entrence to the thorax (video 1). These contractions were only observed during in the cold seasons of the year. At the age of 54 years, she suddenly became unable to swallow her food and water (video 2). Video 1. Rhythmic contractions in the ventral neck area of an Asian elephant Day 1: The elephant tried to drink water. After 10-20 seconds the water came out her mouth again (regurgitation). The regurgitated water was clear and had no abnormal smell (no stomach smell). Appetite: in the morning she ate horse pellets and some roughage, but then she refused bread and apple slices (her favorites!). She tried to eat fresh gras, but after chewing on it, it came out; no smell of stomach contents. Refused to eat anymore. Regular defecation, though the fecal balls became smaller during the following day. The digestion of the fibers had not changed. Water regurgitation Day 2: No change. Oral inspection: 2 small (5 mm Ø) ulcerations on the tongue base, that were not there the day before. The animal did not cooperate as good as she did on the first day. A standing sedation was performed using detomidine and butorphanol. A 2.4 m plastic tube and flexible endoscope could be advanced into the esophagus, reaching the stomach. No mouth opener or gag was used. Gastric fluids were seen, but no obstruction in the esophagus was encountered. Video 2. Regurgitation of water. Endoscope Plastic tube with endoscope advanced into the esophagus. Treatment Rectal fluids Antibiotic + flunixin meglumine + Vit E/selenium Day 3: Standing sedation using detomidine and butorphanol. Treatment: antibiotics, dexamethasone, vitamin B complex, 240 L rectal fluids Day 4: In the morning, the elphant was able to drink water. Nevertheless another standing sedation was performed using detomidine and butorphanol. 3.5 meter tube inserted in esophangus with mini-camera. Antibiotic + dexametasone +vit- sel + complex B + Buscopan Thirty minutes after finishing the procedure, the elephant started to drink and she ate a melon. From that moment on her appetite came back and she did not regurgitate anymore. Differential diagnosis: Esophagus spasm Esophagus constriction: unlikely because this would have been confirmed by endoscopic examination. Botulism . Botulism had occured in the same environment 200 kilometers from this place, resulting in paralysis of the entire body and the death of the 4 affected elephants. Comments: Esophagus spasms in elephants have not been described before. There is only anecdotal evidence of this phenomenon in horses (van der Kolk, pers. comm. 2021). Hypocalcemia is high on the list of the differential diagnoses. Hypocalcemia has also been associated with " hiccups " in Asian elephants, occuring in the cold seasons. The total calcium concentration in this elephant was 2.7 - 2.9 mmol/l while the hematocrite was 48-50% (average normal value 35%), which is an indication that the elephant was dehydrated. The actual total serum calcium concentration in non-dehydrated condition was probably lower: 35/50 x 2.7 = 1.89 mmol/l. Conclusion: Hypocalcemia may have played a major role in this case of esophagus spasm. To page top

This chapter describes (standing) sedation, general anesthesia, intubation and epidural anesthesia using xylazin, ketamine, azaperone, detomidine, medetomidine, etorphin, carfentanil, gas anesthesia and lidocain. To procedures This page describes the following procedures Standing sedation General anesthesia Epidural anesthesia Anesthesia Standing sedation Sedation: In case the elephant does not cooperate voluntarily with the manipulations needed for the diagnosis or treatment the animal should be sedated (including herd mates if needed to reduce stress in the herd) Standing sedation can be performed using xylazine or (preferred) detomidine in combination with butorphanol. Medetomidine works as good as detomidine, but is more expensive. Young elephants need the higher dose range compared to older elephants. Elephants that are excited can be premedicated with azaperone (Asian elephant 0.024-0.038 IM, African elephant 0.056-0.107 IM, IV). Detomidine 0.01-0.022 mg/kg IM (can be reversed by atipamezole at 3-5 times the dose of detomidine). Young calves may need a higher dose of detomidine (0.02-0.04 mg/kg). AND Butorphanol 0.015-0.025 mg/kg given at same time as detomidine. Butorphanol can be reversed with naltrexone at 2.5-5 times the dose of butorphanol in emergency situations, but reversal is not essential and should preferably not be carried out if the calf is considered to be in pain. Alternative option for sedation (if the above mentioned drugs are not available): Xylazine : 0.04-0.08 mg/kg IM for adult Asian elephants and 0.08-0.1 mg/kg for African elephants. Juvenile Asian elephants: 0.09–0.15 xylazine mg/kg IM (Jansson 2021) If insufficient sedation is obtained by xylazine alone, an additional (low) dose of ketamine (0.03 – 0.06 mg/kg) can be given IM or IV. Xylazine can be reversed with yohimbine (0.073-0.098 mg/kg slowly IV) or atipamezole (0.1 x xylazine dose IM or 30/70 IV/IM) Another alternative option for sedation of Asian elephants: Dexmedetomidine : 2 μg/kg BM IM will provide sufficient standing sedation for approximately 70 minutes. (Buranaprim, 2022). Dexmedetomidine can be antagonized by atipamezole (10 times the dexmedetomidine dosage). If a young calf needs to be sedated, it may be necessary to sedate the dam or other adult herd mates so they are not stressed during manipulations on a calf. This can be done by the administration of: Butorphanol 0.006 mg/kg IM and detomidine 0.0026 mg/kg IM (In adult female Asian elephants, 20mg butorphanol and 10mg detomidine have been effective) Sedation can be reversed as described above but is not necessary Alternatively, xylazine (0.04–0.08 mg/kg) or other sedative agents (e.g. Azaperone at 0.024–0.038 mg/kg) can be used if detomidine is unavailable. Laubscher LL et a. 2021 described a fixed drug combination of butorphanol, azaperone and medetomidine (BAM) for African elephants. The dose is given per cm shoulder height. The composition of this anesthetic mixture is: 30 mg/ml butorphanol, 12 mg/ml azaperone, and 12 mg/ml medetomidine. The use of this combination can be recommended in captive, trained African elephants at a dose of 0.006 6 ± 0.001 ml/cm shoulder height. Oral or rectal administration of detomidine in the form of a gel (Domosedan gel, 20-50 mcg/kg) to obtain mild sedation has been described (2020, Molter). The gel must be rubbed into the oral mucosa or rectal wall. Initial, mild sedation is seen after 15-20 minutes. The maximal effect is at 30-45 minutes. A full standard sedation is characterized by the following signs: Salivation Relaxation of the trunk; the tip of the trunk will touch the ground. Relaxation of the penis and (less obvious) relaxation of the vulva. Snooring sounds. It is important to cover the eyes with gauze pads (taped to the skin with Leucoplast or ducttape) and put cotton plugs in the ears. This will deepen the sedation and reduce the risk of sudden wakening. One should always be prepared that the elephant may wake up. Safety procedures need to be discussed in advance with everyone involved in the procedure. Summary agonist - antagonists Xylazine can be reversed by atipamezole : 0.1 x xylazine dose or yohimbine : 0,05-0,13 mg/kg IV Detomidine is reversed by: atipamezole: 3-5 times the detomidine dose IM or slow IV (30/70 IV/IM) Butorphanol is reversed by naltrexone: 2.5-5 x butorphenol dose IV. Skip naltrexone if pain relieve is desirable. The naltrexone dosage provided by Laubscher LL et a. 2020 is much lower: 1 mg naltrexone per mg butorphanol. References: Buranapim, N., Kulnanan, P., Chingpathomkul, K., Angkawanish, T., Chansitthiwet, S., Langkaphin, W., Sombutputorn, P., Monchaivanakit, N., Kasemjai, K., Namwongprom, K., Boonprasert, K., Bansiddhi, P., Thitaram, N., Sharp, P., Pacharinsak, C., Thitaram, C., 2022. Dexmedetomidine Effectively Sedates Asian Elephants (Elephas maximus ). Animals 12, 2787.. doi:10.3390/ani12202787 Fowler M.E. and Mikota S.K. 2006. Chemical restraint and general anesthesia. In: Biology, medicine and surgery of elephants. Blackwell Publishing. Jansson T., Vijitha P.B., Edner A., and Fahlman A. 2021. Standing sedation with xylazine and reversal with yohimbine in juvenile Asian elephants ( Elephas maximus ). Journal of Zoo and Wildlife Medicine, 52(2) : 437-444. Liesel L. Laubscher , Silke Pfitzer , Peter S. Rogers , Lisa L. Wolfe , Michael W. Miller , Aleksandr Semjonov , Jacobus P. Raath. 2021. Evaluating the use of a butorphanol-azaperone-medetomidine fixed-dose combination for standing sedation in African elephants (Loxodonta africana). J. of Zoo and Wildlife Medicine, 52(1) :287-294 (2021). Molter C. 2020. Diagnosis and treatment of EEHV-hemorrhagic disease. Proceedings of the annual AAZV- symposium 2020. Neiffer D.L. , Miller M.A., Weber M., Stetter M., Fontenot D.K., Robbins P.K., and Pye G.W. 2005. Standing sedation in African elephants (Loxodonta africana) using detomidine–butorphanol combinations. Journal of Zoo and Wildlife Medicine 36(2): 250–256, 2005. E. Wiedner. 2015. Proboscidea. In: Fowler's Zoo and Wild animal Medicine 8. Standing sedatin General Anesthesia General remarks: General anesthesia is required in those cases where standing sedation alone or in combination with local anesthesia does not suffice for the intervention that needs to be done. We can devide the indications in: Capture immobilization Immobilization for painful procedures Capture immobilization is mostly done in range countries. However, the escape of a captive elephant may also require capture immobilization. Elephants from this category have not been prepared for the immobilization. This means that they have been able to take food an water shortly prior to the immobilization. It aslo nmeasn that the circumstances have not (or insufficiently) been prepared for the procedure as compared to an immobilization under full captive conditions. Preparation : If possible, prepare a safe area for the people and elephant involved. Avoid an area with water and select a place that is reachable for heavy equipment. Provide shadow whenever possible. Make sure you can ccol the elephant with cold water when necessary. Heavy equipment to position the elephant in lateral recumbancy may be needed, as sternal recumbancy is highly associated with anesthetic death. If an elephant has gone down in sternal position and cannot be rolled over in lateral recumbancy, the anesthesia must be reversed immediately. Whenever possible, provide a soft bedding, preferably a deep sand layer covered by a deep layer of straw or matrasses. Straps or belts are required in case the elephant needs to be rolled over. It is important to thraw them under the elephant before the animal will go down. It helps if the elephant lays on sand and straw to get straps or a belt under the elephant's body with the help of a hooked steel wire. To protect the tusks against fractures, a car tyre can be placed under the head just before the elephant goes down. Trained elephant can be anesthetized when brought lateral recumbency. If the elephant is trained to ly down in sternal position, general anesthesia can be induced but this is very risky! Once the drugs have reached their effect, the elephant MUST be rolled over into lateral recumbency, which requires heavy equipment. Especially in trained elephants, ropes can be used to guide the elephant into lateral recumbency. Trained captive African elephant brought under general anesthesia while guided by ropes. Courtesy: Osterhaus and Fagan. For correct positioning of the elephant during general anesthesia, the use of a crane is highly recommended. First, a standing sedation is induced. After a net has been brought into position, this can be connected to the crane. This will support the elephant when the general anesthesia is induced by IV or IM injection of the narcotic drug (etorphine or ketamine). By lifting the elephant it can be positioned in the correct lateral recumbancy. Protecting cushions, matrasses and soft bedding materials should be placed underneath the head and the body. See the images of the use of a net below (Courtesy basel Zoo): Elephants should be fastened for 24-48 hours prior to anesthesia. Water should be withheld for 24 hours before the procedure. Capture immobilization is mostly done in range countries. The escape of a captive elephant may also require capture immobilization. Elephants from this category have not been prepared for the immobilization. This means that they have been able to take food an water shortly prior to the immobilization. It also means that the circumstances have not or insufficiently been prepared for the procedure as compared to an immobilization under full captive conditions. Preparation: if possible, prepare a safe area for the people and elephant involved. Avoid an area with water and select an area that is reachable for equipment. Provide shadow whenever possible. Make sure you cool the elephant with cold water when necessary. Heavy equipment to position the elephant in lateral recumbancy may be needed, as sternal recumbancy is highly associated with anesthetic death. If an elephant has gone down in sternal position and cannot be rolled over in lateral recumbancy, the anesthesia must be reversed immediately. Whenever possible, provide a soft bedding, preferably sand covered by a deep layer of straw or matrasses. If straps are required in case the elephant needs to be rolled over, it is important to thraw them under the elephant just before the animal will go down. It helps if the elephants lays on sand and straw to get straps or a belt under the elephant's body with the help of a hooked steel wire. The use of a suitable net is highly recommended as slings may slide away from the desired place of the elephant's body. Oxygen supplementation Oxygen must always be provided, even if the anesthetized elephant is not intubated. Arterial blood pressure will drop if no oxygen is provided (Heard 1986). An oxygen flow of 10-15 L/min for a juvenile up to 39-40 L/min for an adult elephant is required for maintaining arterial blood pressure at an acceptable level. Oxygen supply during general anesthesia of a 5 yr-old Asian elephant under field conditions. Due to lack of proper equipment, intubation was not possible. Oxygen was provided at a flow rate of 10 L/min via a small tube inserted in the trunk. Drugs used for general anesthesia: Captive elephants that are excited can be premedicated with azaperone (Asian elephant 0.024-0.038 IM, African elephant 0.056-0.107 IM, IV). Fast acting immobilizing drugs that are used for capture immobilization: Etorphine : 0.002-0.004 mg/kg IM (Asian elephant) and 0.0015-0.003 mg/kg IM (African elephant) OR Carfentanil : 0.002-0.004 mg/kg (Asian elephant) and 0.0013-0.0024 mg/kg IM (African elephant) These drugs can be antagonized with naltrexone 0.004 mg/kg IM (or 50/50 IV/IM) If carfentanil and etorphine are not available, xylazine (0.1 mg/kg) and ketamine (0.3-0.7 mg/kg) can be given together. The disadvantage is the large volume required for an adult elephant. For capture immobilization this combination is therefore not recommended. At the end of the procedure xylazine can be reversed with atipamezole (0.1 x dose of xylazine IM or slowly IV) or yohimbine (0.05-0.13 mg/kg IV). Under controlled conditions (if a crane is available) a standing sedation can be induced first, allowing to put a net or slings in place. When well secured, ketamine can be given i.m. (0.3-0.7 mg/kg). or i.v. using a long infusion tube for safety reasons. Once in lateral recumbancy, the elephant can be intubated and anesthesia can be maintained on isoflurane or halothane (1.5-3%). Inhalation anesthesia and intubation: Intubation in elephants is straightforward. A 30-50 mm diameter cuffed endotracheal tube can be inserted into the trachea. A rope around the lower jaw can be used to open the mouth. A gloved hand can reach the epiglottis and advance a lung tube (e.g. stocha tube for horses) into the trachea, while pushing the soft palate upward. Once in place, the endotracheal tube can be advanced into the trachea guided by the smaller tube. A special portable pressure ventilater has been developed and described by William et al. Jeff Zuba made some modifications to this design, which is now commercially available (http://www.incaseofanesthesia.com/Home_Page.html ). Schematic overview of a portable pressure ventilation device for elephants. "Zuba" ventilator used in an adult African elephant under field conditions Captive African elephant intubated for gas anesthesia using a "Zuba" ventilator. "Zuba" ventilator Under less favorable circumstances when a pressure ventilator is not available, intubation can be done in the trunk using 2 cuffed horse endotracheal tubes and 2 separate (portable) anesthetic machines (Tamas 1983). The advantages of this method are the easy intubation and the ample space in the oral cavity in the absence of the large tube. However the disadvantages are substantial: Two tubes increase the airway resistence Risk of regurgitation and aspiration of stomach contents An elephant can breath through its mouth, which will bypass the inhalation of the anethetic gas General anesthesia in a captive Asian elephant using bilateral trunk intubation (Rotterdam Zoo, 1989) Monitoring: Pulse oximetry is a reliable tool for monitoring heart frequency and venous oxygen saturation. A capnagraph is recommended to monitor the respiration. If not available, one individual should be assigned just to monitor respiratory rate and depth. ECG and arterial blood gases are recommended. As hypotension is quite common in anesthetized elephants, blood pressure measurement is also recommended. Hypotension has been treated successfully with ephedrine and dobutamine. Recovery support: Weak or debilitated animals may need help to get back on their feet during recovery. A deep sand layer is essential for the elephant to getting grip on the ground. A crane may be needed to lift the animal from the ground, using straps or belts applied around the body. References. Fowler M.E. and Mikota S.K. 2006. Chemical restraint and general anesthesia. In: Biology, medicine and surgery of elephants. Blackwell Publishing. Heard D.J., Jacobson E.R., and Brock K.A. 1986. Effects on oxygen supplementation on blood gas values in chemically restraint juvenile African elephants. J Am Vet Med Ass 189 (9)1071-1074. Tamas PM. and Geiser D.R. 1983. Etorphine analgesia supplemented by halothane anesthesia in an adult African elephant. JAVMA 183, 11 (1312-1314) . Wiedner E.. 2015. Proboscidea. In: Fowler's Zoo and Wild animal Medicine 8. Zuba J.R., Osterhaus J.E. 2012. Anesthetic complications and clinical intervention in opiod anesthetized captive elephants. In: Proceedings of the AAZV Conference, Oakland (1-6). Zuba J.R. http://www.incaseofanesthesia.com/Home_Page.html General anesthesia Always bring the elephant into LATERAL RECUMBANCY for general anesthesia Epidural anesthesia Epidural anesthesia in elephants is recommended when a vaginal vestibulotomy is performed in order to reduce tail movements of the elephant and provide additional analgesia in the perineal region. Procedure: Restrain the elephant as appropriate in a chute and sedated if necessary. Disinfect the injection site. Move the tail up and down to determine the position of the most mobile intercoccygeal space. Inject local anaesthetic (2% Lidocaine) into the skin over the injection site. Palpate the intercoccygeal space wearing a sterile glove and insert the needle (14 gauge, 3 inch) at approximately a 60 - 70 degree angle cranially. The epidural space is about 6.5 cm below the skin surface. Inject Lidocaine : 30 ml was sufficient to produce tail relaxation in a 3,000 kg elephant, and the elephant remained standing. Epidural anesthesia

Intramuscular, subcutaneous, intravenous and epidural injection techniques are described in this chapter. To procedures Injection techniques Hand injections Intramuscular injection: Subcutaneous injection: Intravenous injection: Epidural injection: An epidural injection in elephants is recommended when a vaginal vestibulotomy is performed in order to reduce tail movements of the elephant and provide additional analgesia in the perineal region. Procedure: Restrain the elephant as appropriate in a chute and sedated if necessary. Disinfect the injection site. Move the tail up and down to determine the position of the most mobile intercoccygeal space. Inject local anaesthetic (2% Lidocaine) into the skin over the injection site. Palpate the intercoccygeal space wearing a sterile glove and insert the needle (14 gauge, 3 inch) at approximately a 60 - 70 degree angle cranially. The epidural space is about 6.5 cm below the skin surface. Inject Lidocaine : 30 ml was sufficient to produce tail relaxation in a 3,000 kg elephant, and the elephant remained standing. Remote injections Make your own blow dart Blow dart injection: Jam-stick injection: Dart gun injection: To page top

Broncho-alveolar lavage (BAL) describes: gastroscopy, bronchoscopy, esophagus, trachea, mouth-opener, mouth gag. Approach is done either via the trunk or via the via, always under standing sedation. The purpose of a BAL is to collect samples for culture/PCR (tuberculosis) or histology. To physical examination Sample collection Trunk wash Broncho-alveolar lavage Trunk wash procedure Trunk wash procedure Compiled by Willem Schaftenaar Background The trunk wash procedure was developed as a method for diagnosing tuberculosis or the detection of Elephant Endotheliotropic Herpes Virus. This procedure is an active manipulation at the elephant trunk, which can be performed in free and protected contact systems in non-immobilized elephants after they are conditioned for this procedure. The principle is that a sterile 0,9% saline solution (approx. 100 ml) is injected in each nostril of the trunk. The trunk has to be lifted actively by the elephant or passively by the keeper so that the solution is running up to the base of the trunk. The mixture of the solution and trunk mucus is collected in sterile plastic bags by active blowing of the elephant through its trunk (training required). The staff should protect themselves against spilling trunk content into their face. A full trunk wash procedure requires 3 different trunk washes performed within a period of 7 days. Each sample must be sealed and stored at 4°C. Depending on the quality of the samples, the diagnostic lab can decide to pool the samples for culture/PCR. Samples must be shipped to the TB-diagnostic lab immediately after the 3-rd sample has been taken. The maximum storage period at 4°C is 7 days. NB: follow the EU guideline for shipment of potentially hazardous biomaterials. For the (q)PCR detection of EEHV, the sample can be shipped directly or kept frozen at -20°C until shipment. Trunk wash in a non-contact situation requires a full anesthesia of the elephant and a portable fluid pump and sucking system, which allows the operation under sterile condition. The external pump and sucking system will be connected to a sterile PVC tube (1 cm diameter, with distance markers) with a length of approx. 2 meter. The amount of sterile solution and the collection bag are like described before. In non-contact situations, a bronchoalveolar lavage (BAL) under standing sedation is the preferred procedure. Training for a trunkwash procedure Broncho-alveolar lavage (BAL) and Gastroscopy Broncho-alveolar lavage Compiled by Willem Schaftenaar Procedure A bronchoalveolar lavage (BAL) may be indicated when a sample from the deeper regions of the lungs is required, such as for the diagnosis of tuberculosis (TB). Gastroscopy is a procedure that allows direct visualization of the esophagus and stomach. It enables the collection of biopsy specimens and stomach contents, and it can also be used to perform a lavage for TB diagnosis. Fluids obtained through BAL are additionally used for the isolation of mononuclear cells and for cytological evaluation (Engel, 2005). The procedure has been performed in free-ranging savanna African elephants under general anesthesia (Engel, 2005). The protocol described here outlines a workflow for the successful isolation and characterization of alveolar cells, predominantly alveolar macrophages, from BAL fluid. Differential cell counts and cellular characterization were carried out. This technique for isolating alveolar mononuclear cells provides a foundation for further investigation into the functions of respiratory immune cells. Under more controlled conditions, the procedure can also be performed under standing sedation . There are 2 ways to approach the alveoli: 1. Trunk approach An endoscope measuring 10–12 m in length is advanced through the trunk ( Hermes et al, 2017 ). The main advantage of this method is that the operator can remain outside the enclosure. However, the procedure requires an extra-long endoscope, which is not readily available. To facilitate passage of the endoscope alongside the cartilage plate in the trunk, bilateral block anesthesia of the nerve rami at the base of the trunk has been recommended (Hildebrandt et al.). Nonetheless, D. Schmidt has reported performing a similar procedure without block anesthesia (pers. comm., 2019). Once advanced, the endoscope allows visualization of the inner walls of the trachea and bronchi for bronchoalveolar lavage; biopsy samples can also be collected if required. In addition, the endoscope can be directed into the esophagus to perform gastroscopy or gastric lavage. 2. Oral Approach In this method, a 5 m long endoscope is introduced through the mouth. The primary disadvantage is that the operator must remain inside the elephant enclosure. A specialized mouth opener for elephants is required to provide access to the oral cavity. The endoscope may be guided manually into the trachea or introduced through a rigid plastic tube, which can be advanced relatively easily into either the trachea or the esophagus. Once within the trachea, the bronchial branches can be examined. Abnormalities may be assessed, and biopsy samples collected. Bronchoalveolar lavage samples are obtained by instilling 50–60 ml of 0.09% sterile saline into a bronchus, followed by aspiration of the fluid. For TB diagnosis, this procedure is repeated three times in each lung. Collection of stomach fluid and broncho-alveolar lavage in an adult African elephant by Dr. Imke Lueders ( https://www.geolifes.com/en/services/index.html) References: Engel S.C., Kerr T.J., van der Spuy G.D., Jooste T., Buss P.E., Johns J.L., Miller M.A., Kleynhans L.2025. Optimisation of bronchoalveolar lavage fluid preparation for mononuclear cell isolation and cytologic evaluation in free-ranging African elephants (Loxodonta africana ). Veterinary Immunology and Immunopathology, Volume 286 , August 2025, 110974. https://doi.org/10.1016/j.vetimm.2025.110974 Hermes R et al. 2018. Bronchoalveolar lavage for diagnosis of tuberculosis infection in elephants. Epidemiology and Infection https://doi.org/10.1017/S0950268818000122 To page top

We developed this website with the input of Elephant Veterinarians worldwide. -Willem Schaftenaar, DVM -Susan Mikota, DVM -Elephant Medicine This website is the joined effort of numerous veterinarians, who work with elephants world-wide. The name(s) of the vet(s) who contributed to a particular topic is mentioned in the left upper corner of each topic page. Some authors may wish to remain anonymous. In that case, the name has not be filled in. Some cases have been published online as open source manuscript and are incorporated in this website via an internet-link. You are encouraged to critically read the information and the clinical cases described on this website and send us your comments! Please submit your clinical cases, no matter whether they have a happy or a sad ending. By sharing our experience we can all learn from each other. Willem Schaftenaar, DVM Website moderator Worked for 30 years at Rotterdam Zoo as clinical veterinarian. Veterinary advisor to the EAZA elephant TAG Associate researcher Elephant Care International. Susan Mikota,DVM Co-founder and Director of Veterinary Programs & Research for Elephant Care International Contributions have been made both anonymously as by name. All the names are known to the moderator. Named contributors are (in alphabetic order) : Dalen Agnew DVM, PhD, DACVP, Department Chair and Associate Professor Pathobiology and Diagnostic Investigation at Michigan State University (USA) Marcus Clauss , University of Zürich (Switzerland) Thittaya Janyamethakul (Tip), DVM., MS, Patara Elephant Farm, Chiang Mai (Thailand) Christine Kaandorp , head veterinarian Rotterdam Zoo (The Netherlands) Arne Lawrenz DVM, director of Wuppertal Zoo (Germany) Susan Mikota DVM, Co-founder and Director of Veterinary Programs & Research for Elephant Care International (USA) Fieke Molenaar DVM, senior veterinary officer Zoological Society of London Zoo (UK) Joost Philippa DVM, zoo veterinarian at Rhenen Zoo (The Netherlands) Tina Risch DVM, Thüringer Zoopark Erfurt, Am Zoopark 1 D-99087 Erfurt (Germany) Ann-Kathrin Oerke , PhD, researcher at Göttingen Primate Center (Germany) and Research advisor to the EAZA elephant TAG Vijitha Perera DVM, Senior veterinarian Elephant Transit Home, Sri Lanka Van Thin Pham DVM, head veterinarian at Dak Lak Elephant Conservation Center (Vietnam) Christian Schiffmann DVM, veterinarian (Germany), research advisor to the EAZA elephant TAG Linda Schiffmann , zoo keeper, TBZ - Tierbegegnungszentrum, Hochrhein (Germany). Linda van Sonsbeek, DVM, head veterinarian Rotterdam Zoo (The Netherlands) Taweepoke Angkawanish PhD,DVM, Lampang Elephant Conservation Center (Thailand) Francis Vercammen DVM, zoo veterinarian at Antwerp and Planckendael zoo (Belgium) Jürg Völlm DVM, Basel Zoo (Switzerland, † 2021) Christian Wenker DVM, Basel Zoo (Switzerland) Kasper Willebrands, elephant headkeeper Rotterdam Zoo (The Netherlands) About Us

Bone fractures are not uncommon in elephants. Radiography is needed for a proper diagnosis. A case of mandibular fracture is described in this chapter. To non-infectious diseases Bone fractures Mandibular fracture Mandibular fracture Mandibular fracture Mandibular fracture Mandibular fracture Mandibular fracture Bone fractures Mandibular fracture

Dirt, foreign bodies, a short tusk remnant after a tusk fracture and parasites (Cobboldia sp) can cause a purulent sulcus infection in elephants. To parasitology To dentistry case report Tusk sulcus infection Place: Dak Lak elephant Conservation Center Vietnam Date: 2017 Data provided by: Van Thinh Pham, DVM History Purulent discharge from the dental sulcus in an adult Asian elephant bull since a few days. The area around the sulcus was itching, demonstrated by the bull by frequent rubbing the tusk base against trees. Diagnosis Frequent blowing sand in the sulcus area may also be the cause of this problem. in this case, no sand or dirt was present in the sulcus. An infection with larvae of the stomach bot ( Cobboldia sp. ) was suspected. Treatment The dental sulcus area was cleaned with cotton wool and flushed with Betadine and the bull was treated with ivermectin SQ, 0,2 mg/kg BW Treatment results The sulcus lesion healed completely within 7 days. Cobboldia (stomach bot) larvae To page top

Edema in elephants is not uncommon. The 2 most frequently seen forms are edema in the neck, head and upper parts of the front legs (EEHV-HD) and ventral edema (general edema). Figure 1. (a) A focal moderate ventral edema. Note the smooth skin structure with reduced wrinkles in the edematous region. (0.1 African elephant, 33yrs.) (b) Focal moderate ventral edema in lateral view. (0.1 African elephant, 22yrs., 2.5 months before giving birth) (c) A moderate ventral edema extending to the external genital region. (0.1 African elephant, 46yrs., advanced pregnancy >18 months). Figure 2. Ventral edema in a 7 yr-old Asian elephant bull suffering of Salmonellosis (Photo: Willem Schaftenaar). Click here to read this case report. Figure 3. A 45 yr-old female African elephant with ventral edema showing signs of irritation (left) and sloughing of skin (right, arrow). Differential diagnosis of ventral edema In the young elephant a swelling around the umbilicus can be an indication of an umbilical hernia , sometimes accompanied by local edema. A blunt trauma of the abdominal wall can result in an abdominal hernia. Intestines can be visualized using ultrasound examination. Figure 4. Asian elephant (>35yrs) with traumatic ventral hernia. Movement of the intestines and fecal balls in the subcutaneous space could be visualized during transcutaneous ultrasound examination. Photo: Willem Schaftenaar Pathogenesis In general The body always tries to maintain the balance between intravascular and interstitial fluid, driven by four different pressures acting in the capillary bed (Fig. 5). In addition, the capacity of the lymphatic system is critical for the physiologic reabsorption of interstitial fluid and its return transport into the blood circulation (Fig. 5). Beside an increased permeability of the capillary wall, any alteration in each of these five factors can cause edema. In particular an increase in the capillary hydrostatic pressure and a decrease in the plasma oncotic pressure (= osmotic pressure induced by the plasma proteins) lead to an increased shift of fluid towards the interstitial space. If this fluid load exceeds the lymphatic capacity, fluid will accumulate and edema will develop. The aforementioned parameters do vary across different body regions, leading to a locally varying susceptibility to edema development. Therefore, edema can occur both multifocal (e.g. EEHV-HD) or focal (e.g. ventral edema) with respect to the predisposition of certain body regions and the underlying cause. The latter can be systemic or focal. The localization of edema is also determined by gravity forces and species-specific anatomical characteristics. Extracellular fluid will have the tendency to migrate downwards due to gravity. Extracellular spaces that are surrounded by tightly fitting, non-elastic tissue, are not prone to show edema, even if they are at the lowest point of the body: in humans edema can easily develop in the feet, while in elephants edema has never been reported in the distal parts of the limbs. Figure 5. Four critical parameters are determining fluid shift in the capillary bed through the semipermeable capillary wall. An increase in capillary hydrostatic pressure and interstitial oncotic pressure leads to an increased fluid shift towards the interstitial space, as well as a decrease in plasma oncotic pressure and interstitial hydrostatic pressure. The lymphatic vessels are running in parallel to the blood vessels and are collecting the interstitial fluid according to their transport capacity. In elephants According to Mikota (2006), no single underlying etiology for ventral edema in elephants has been identified so far. More likely, it presents a non-specific response to a variety of physiological stressors (Mikota 2006). In our opinion these stressors or pathological alterations can be categorized based on the general pathogenesis of an edema (Fig. 5). With this approach, each condition reported to be associated with edema in elephants so far, can be ascertained to one of the four defined etiologic categories (Fig. 6). Fowler & Mikota (2006) consider the ventral distribution of an edema in elephants caused by the gravitation of fluids into this area. But if gravitation alone would present the critical parameter for the characteristic ventral occurrence of an edema in the elephant, one would expect the swelling to occur primarily in the distal limb regions. The very thigh skin surrounding the legs with minimal elasticity may prevent this pattern. Apart from this, we assume the anatomy and physiology of the lymphatic system to explain the specific distribution pattern of ventral edema in elephants (Fig. 7). Unfortunately, anatomical knowledge on the lymphatic system in elephants is limited to one incomplete description in a fetal Asian elephant (Mariappa 1986). In this individual, a peculiarity was reported with the Cisterna chyli located in the thoracic cavity (Mariappa 1986). In humans, the horse, dogs & cats the Cisterna chyli is located in the abdominal cavity (Berens von Rautenfeld 2000, Herpertz 2013, Salomon et al. 2008). We do rather question the validity of the report for the fetal Asian elephant, than expect a significant peculiarity in the anatomy of the lymphatic system in the elephant. Figure 6. Four defined etiologic categories for edema in elephants, each with examples reported in the existing literature. Note that underlying alterations may vary extremely but result in the same clinical sign of accumulated interstitial fluid. Therefore, due to the lack of solid evidence, our line of arguments is largely based on the anatomy of the lymphatic system in horses and extrapolated to the elephant (Berens von Rautenfeld 2000, Salomon et al. 2008; Fig. 7). Assuming that the lymphatic watersheds in the elephant are running similar to the situation in the horse, it becomes obvious that the characteristic location of a ventral edema presents the region between the major transversal and horizontal watershed (Fig. 7). In this proximal part of the lymphatic territory VII, the lymphatic vessels drain towards the deep abdominal lymphatic centers and have no connection to a relevant superficial lymphocentrum. Therefore, it seems reasonable that increased abdominal pressure (e.g. during late pregnancy) may reduce the drainage of this territory. At the same time, interstitial hydrostatic pressure in the subcutaneous tissue of this body region may be low compared to the limb or the thoracic wall where bony and muscular structures are supporting the lymphatic capacity. These factors together with gravitation can serve as an explanation for the specific distribution of ventral edema in elephants. In contrast, the limbs may rarely be affected by edema, because the relatively rigid skin in combination with the underlying musculoskeletal apparatus will result in kind of a physiologic compression bandage as reported for the horse (Aurenz 2020). Figure 7. Hypothesized lymphatic territories in the elephant. The seven distinct lymphatic territories with their specific drainage areas were extrapolated from the situation in the horse (Berens von Rautenfeld et al. 2000) and numbered accordingly. The blue lines indicate the lymphatic watersheds. Note the proximal part of area VII is lacking a connection to a relevant superficial lymphocentrum. Treatment of ventral edema Given the wide diversity of underlying causes (Fig. 6) no general treatment protocol can be defined. In the literature, hot and cold pressure bandages (du Toit 2001) and increasing protein in the diet (Fowler & Mikota 2006) have been recommended. The administration of Furosemide (1mg/kg i.m.) has been unsuccessful (Martelli 2006). Considering the different etiological pathways leading to an edema, we strongly encourage the treatment of the underlying cause. To do so, an underlying cause needs to be determined or at least a classification according to Figure 6 should be strived for. The latter seems realistic by a thorough anamnesis and clinical examination. For example in cases of heart failure, positive inotropic agents may reduce the capillary hydrostatic pressure and simultaneously support the lymphatic capacity, as shown in humans (Scallan et al. 2016). In less severe cases of assumed cardiorespiratory insufficiency, which has been observed to repeatedly cause ventral edema in geriatric Asian females during hot summer days, herbal medicine can present a helpful approach (Crataegus Dilution vet.®, DHU-Arzneimittel GmbH & Co. KG, Karlsruhe, Germany; three times a day, 6.0-8.0ml orally) (personal observation in four cases). In addition to the treatment of the underlying cause, or in cases where only a symptomatic treatment is realizable, the following methods may facilitate the reabsorption of an edema. Moderate walking will centrally activate the lymphatic flow and subsequently increase the lymphatic capacity. Hence locomotion is considered a critical part of edema therapy in horses (Aurenz 2020). A sufficient amount of satisfying recumbent rest will also support the reabsorption of interstitial fluid by reducing the negative impact of gravitation. Moderate pressure washing may have a positive effect similar to manual lymph drainage in horses (Aurenz 2020). Under the assumption of a similar anatomy of the lymphatic system, adhering to the protocols established in equine lymph drainage seems a reasonable approach (Berens von Rautenfeld 2000). Given that the selectively applied pressure for manual lymph drainage could be applied by a water jet, even treating from a safe distance might become an option. Further research is needed to base such an approach on scientific findings and formulate a detailed practical guidance. Additional note With respect to our very limited knowledge on the anatomy of the lymphatic system in elephants and the corresponding physiological pathways, a major part of this compilation is very hypothetical. Although we based these hypotheses on evidence from other mammalian species, they remain to a certain amount speculative and should be interpreted with caution. References Aurenz S (2020). Manuelle Lymphdrainage beim Pferd. Hands on 2:25-31. Berens von Rautenfeld D, Rötting A, Rothe K, Lüdemann W, Boos A, Schubert T, Hertsch B (2000). Manuelle Lymphdrainage beim Pferd zur Behandlung der Beckengliedmaße - Teil 1: Anatomische Grundlagen und Behandlungsstrategien. Pferdeheilkunde 16:30-36. Caple IW, Jainudeen MR, Buick TD, Song CY (1978). Someclinico-pathologicfindings in elephants (Elephas maximus) infectedwithFasciolajacksoni. Journal of Wildlife Diseases 14:110-115. Chandrasekharan K (2002). Specific diseases of Asian elephants. J Indian Vet Assoc Kerala 7:31-34. du Toit J (2001) Veterinary care of African elephants. South Africa, South African Veterinary Foundation and Novartis. Emanuelson K, Agnew DW (2002). Wasting syndrome in a bull African elephant (Loxodonta africana). IAAAM Joint Conf, New Orleans, Louisiana. Emanuelson K, Kinzley C (2000). Salmonellosis and subsequent abortion in two African elephants. IAAAM Joint Conference New Orleans, Louisiana. Fowler ME, Mikota SK (2006). Biology, Medicine, and Surgery of Elephants. Iowa, USA, Blackwell Publishing. Fuery A, Pursell T, Tan J, Peng R, Burbelo PD, Hayward GS, Ling PD (2020). Lethal hemorrhagic disease and clinical illness associated with the elephant EEHV1 virus are caused by primary infection: Implications for the detection of diagnostic proteins. Journal of Virology 94:1-14. Heard DJ, Kollias GV, Merritt AM, Jacobson ER (1988). Idiopathic chronic diarrhea and malabsorption in a juvenile African elephant (Loxodonta africana). The Journal of Zoo Animal Medicine 19:132-136. Herpertz U (2013). Ödeme und Lymphdrainage. Stuttgart, Schattauer Verlag. Howard L, Schaftenaar W (2019). Elephant endotheliotropic herpesvirus. Fowler´s zoo and wild animal medicine: current therapy. E. Miller, N. Lamberski and P. Calle. St. Louis, Elsevier:672-679. Jensen J (1986). Paralumbar kidney biopsy in a juvenile African elephant. Proc Amer Assoc Zoo Vet, Chicago, Illinois. Lueders I, Young D, Maree L, van der Horst G, Luther I, Botha S, Tindall B, Fosgate G, Ganswindt A, Bertschinger H (2017). Effects of GnRH vaccination in wild and captive African elephant bulls (Loxodonta africana) on reproductive organs and semen quality. PLoS ONE 12:e0178270. Mariappa D (1986). Anatomy and histology of the Indian elephant. Michigan, USA, Indira Publishing House, Michigan, USA. Martelli P (2006). Veterinary problems of geographical concern - Section III Indochina and Bangladesh. Biology, Medicine, and Surgery of Elephants. M. E. Fowler and S. K. Mikota. Ames, Iowa 50014, USA, Blackwell Publishing: p. 452. Mikota SK (2006). Chapter 18 - Integument System. Biology, Medicine, and Surgery of Elephants. M. E. Fowler and S. K. Mikota. Ames, Iowa 50014, USA, Blackwell Publishing: pp. 253-261. Morris P, Held J, Jensen J (1987). Clinical pathologic features of chronic renal failure in an African elephant (Loxodonta africana). 1st Intl Conf Zool Avian Med, Turtle Bay, Hawaii. Murray S, Bush M, Tell L (1996). Medical management of postpartum problems in an Asian elephant (Elephas maximus) cow and calf. J Zoo Wildl Med 27:255-258. Perrin KL, Kristensen AT, Bertelsen MF, Denk D (2021). Retrospective review of 27 European cases of fatal elephant endotheliotropic herpesvirus-haemorrhagic disease reveals evidence of disseminated intravascular coagulation. Scientific Reports 11(1):14173 Pinto M, Jainudeen MR, Panabokke R (1973). Tuberculosis in a domesticated Asiatic elephant (Elephas maximus). VetRec 93:662-664. Salomon F-V, Geyer H, Gille U (2008). Anatomie für die Tiermedizin. Stuttgart, Enke Verlag. Scallan J, Zawieja S, Castorena-Gonzalez J, Davis M (2016). Lymphaticpumping: mechanics, mechanisms and malfunction. J Physiol 594.20:5749-5768. Seneviratna P, Wettimuny S, Seneviratna D (1966). Fatal tuberculosis pneumonia in an elephant. Vet Med Small Anim Clin 60:129-132. Windsor RS, Scott WA (1976). Fascioliasis and salmonellosis in African elephants in captivity. British Veterinary Journal 132:313-317. Edema by Christian & Linda Schiffmann Definition A local or general swelling due to excessive accumulation of fluid in the interstitial space of tissues. This condition can be caused by various underlying alterations. Depending on the composition of the fluid (in particular the protein content), an edema can be further categorized. Relevance of edema in elephants In elephants the occurrence of the so-called ventral edema is a well-known and quite common clinical symptom (Mikota 1994) (Fig. 1 and 2). Ventral edema is defined as edematous swelling in the ventral abdominal wall and tissues surrounding the external genitalia (Mikota 2006). Although the clinical impact of ventral edema is often not visible, the underlying mechanism indicates a disturbance of the internal fluid balance. In addition, edema in the submandibular region and multifocal has been described in cases of hemorrhagic disease due to herpes virus infection (EEHV-HD) (Fuery et al. 2020, Howard & Schaftenaar 2019). Clinical signs The characteristic swelling in edema may develop immediately or over the course of several days, depending on the underlying cause. Edemas caused by a local inflammatory response may be warm and painful upon palpation. The swelling in case of ventral edema without any underlying inflammatory process may feel slightly cooler compared to other body regions. Palpation is not painful and moderate pressure with the thumb may result in a dent. Such dent may also be produced if the edema is the result of an inflammatory process, in which case the pressure will provoke a pain reaction. Compared to non-edematous areas, the skin will look smoother with reduced wrinkles (Fig. 1a). If ventral edema extends from the umbilical to the genital area (Fig. 1c), the skin may become irritated through the repeated contact with the medial hind legs while walking. In severe cases, this irritation may lead to pressure necrosis and sloughing (Mikota 2006). In cases without such complications, edema may resolve without treatment within months (Mikota 1994), although this will heavily depend on the underlying cause, which should be treated accordingly. Prevention Depending on the underlying cause, the occurrence of ventral edema in elephants can be prevented. A continuous health monitoring program with focus on individuals at peculiar risk such as geriatric elephants, pregnant females or individuals suffering from cardio-respiratory or renal insufficiency will enable early supportive and/or curative treatment. Is ventral edema bad? Although ventral edema as such may not necessarily present a serious condition in an elephant, it is often associated with serious health issues and bears the risk for complications. Therefore this symptom should be investigated thoroughly and its development monitored closely.