Tusk fractures should be treated as soon as possible. Both conservative treatment and surgical treatment, using a threaded rod and dental filling materials are described. Step by step you are guided through the entire procedure. To tusk fracture By Willem Schaftenaar (DVM) with a big thank you to the dental team of the Colyer Institute in San Diego: Jim Oosterhuis (DVM), Dave Fagan (dentist, and founder of Colyer Institute), Jeff Zuba (DVM, elephant anesthesiologist ), Allison Woody ( board certified veterinary dentist), Fred Pike (DVM, board certified veterinary surgeon) Tusk fracture repair Tusk fractures are not uncommon in elephants, both in the wild as well as under captive conditions. A tusk fracture can be the result of fights, playing with "toys" (e.g. a tire hanging on a chain), digging in the soil or hitting a wall or other heavy objects (e.g. bulls into musth). Fractured tushes in female Asian elephants usually need no treatment, as the dental pulp does not pass the tusk sulcus. However, fractures in tusks are vulnerable for pulp exposure. If not treated in due time, exposed pulp may become infected and become necrotic, finally resulting in the loss of the entire tusk. Cutting the tusk too short may also result in pulp exposure. Sulcus infection after tush fracture If a tush or tusk fractured proximal to the tush sulcus, sharp pieces of the remaining part of the tush or tusk may cause wounds in the sulcus. Treatment consists of removing these sharp edges of the tusk by rasping them off. The sulcus wounds should be treated like a superficial skin wound by daily cleaning and flushing with Ringer's lactate and an antiseptic (e.g. Betadine-iodine or Chlorhexidine 1%). WARNING: if the pulp tissue is exposed, the elephant should be vaccinated against tetanus! Pulp exposure and tusk growth Exposure of the pulp tissue always results in a bacterial pulpitis. As long as sufficient healthy pulp tissue is present in the apex of the tusk, the tusk may continue to grow. However, if the pulpitis is not treated properly, the infection will finally affect the entire pulp and the tusk will become necrotic and will need to be extracted. Treatment of open tusk fractures There are 2 approaches that are being applied as treatment of a fractured tusk: A. Conservative treatment B. Surgically filling the pulp canal A. Conservative treatment of fractures with minimal pulp exposure: daily cleaning and flushing with Ringer's lactate and an antiseptic (e.g. Betadine-iodine or Chlorhexidine 1%). This is not the preferred treatment option as it will often result in a permanent fistula, as shown here on the photo (small black spot). Nevertheless, the pulp canal was closed by newly formed secondary ivory. Conservative treatment of a tusk fracture with exposed pulp tissue, treated conservatively. The pulp canal closed in 3 months, leaving a very small fistula, which luckily that did not cause any clinical troubles during the following (6+) years. B. Surgically filling of the pulp canal The difference with a conservative treatment approach is the active closing of the pulp canal by a dental surgical procedure. This procedure consists of a partial pulpectomy, followed by closure of the pulp canal and will be described below: Tusk repair procedure Until the actual repair procedure will take place, any free hanging pulp tissue should be cut off and the exposed pulp tissue should be cleaned and flushed 3-4 times per day with saline solution. End each session by spraying Betadine solution or Chlorhexidine 1% over the pulp tissue. Antibiotics are usually not required as the wound is open and under control by flushing. However, the elephant should be vaccinated against tetanus. NSAIDs are only needed if the elephant shows signs of pain. The tusk repair procedure should be performed as soon as possible after the tusk fractured. Tusk repair procdure Fractures that are more or less perpendicular to the tusk length axis have better chances to heal than oblique fractures that extend beyond the sulcus. The best chances to heal properly are fractures with a tusk remnant that allows perpendicular shortening through healthy pulp tissue. In the photo shown here the following structures can be distinguished: exposed pulp tissue, the wall of the tusk remnant and the sulcus mucosa. If the remaining pulp tissue is hanging outside the tusk remnant immediately after the fracture, it is very likely that the proximal part of the remaining pulp tissue has detached from the inner tusk wall, which will result in pulp necrosis if not treated immediately after the tusk fractured. If sufficient pulp tissue can be removed to reach healthy tissue, the prognosis of complete healing is better than in case the pulp tissue encountered after pulpectomy is still detached from the inner tusk wall. Pulp tissue that is hanging outside the pulp canal should be cut off as soon as possible. Preparation –Check the equipment list –Prepare the area where the elephant will be treated S tanding sed ation or general anesthesia? Depending on the conditions of the facility, the procedure can be done under : - Standing sedation , using detomi di ne and butorphanol (or xylazine and butorphanol if detomidine is not available). Azaperone can be used as premedication. Xylazine alone has also been used in a range country where detomidine was not available. There must be sufficient access to the working area. Best is to chain the animal to a wall on both legs on the contra-lateral side of the fractured tusk. The use of a belly belt around the abdomen is highly recommended for safety reasons in case the elephant goes down. - General anesthesia is not strictly required, but under certain circumstances it is a good alternative if standing sedation is not an option. Once the animal is secured either in standing sedation or under general anesthesia, the tusk repair can start: Step 1: create sterile workfield 1. Scrub the affected tusk thoroughly using Povidone iodine scrub. 2. Cut off the tip of the fractured tusk using a giggli wire. Keep the soft tissue out of reach of the giggli wire!!! 3. If present, remove all abnormal (black) ivory using the Dremel. 4. Clean and brush the area (tusk and face) with soap and Betadine scrub. Flush with Betadine solution and alcohol (70%). 5. Cover the surrounding, disinfected skin with a sterile surgery sheet (secure with duct tape) 6. Put on surgical gloves and suit. Step 2: filling the pulp canal 7. Cut off 20-50 mm of the pulp tissue (depending on the diameter of the open pulp canal). If any pulp tissue has been pulled out when the tusk fractured, it is assumed that it has been separated from the tusk wall and when it snapped back in, it probably pulled in bacteria. So even if the pulp looks fresh when doing the pulpectomy, i.e., bacteria could be lurking way up the wall of the canal that you can't get to. This is a challenging part of the procedure. The pulp tissue has a rubbery consistency and needs to be cut with very sharp instruments (curved scissors and scalpels). High-speed cutters used in hip replacement procedures in dogs (Acetabular Reamer) have been used . Depending on the diameter of the pulp canal, a decision needs to be made either to fill the pulp canal at this point, or to bring in a threaded rod (or pulp insert plug) to stop the bleeding. If the pulp diameter is less than 5 mm, one can decide to skip the threaded rod method (skip steps 8, 9, 13 and 14). A threaded rod should be used in any pulp canal diameter larger than 5 mm. In that case, follow the entire procedure as written below. The aim of the plug is to stimulate normal dentinal bridge formation proximal to the plug insert. 8. Drill the pulp canal out to a perfectly round cylinder of the proper size, which corresponds with the diameter at the end of the drilled hole. A so-called step drill can be helpful to accomplish this task. 9. Tap threads into the wall of the tusk so that you have full threads of the plug in the tusk, PLUS, at least 2-3 cm of tusk wall above the plug. 10. Stop bleeding by compressing the pulp tissue gently with epinephrine-impregnated gauze (for several minutes). 11. Fill in the canal with Calcium Hydroxide or calcium hydroxy apatite with a push rod, which mixes some with the blood, and occasionally stops the bleeding for a short time. Other calcium sources that have been used successfully are: Calcium hydroxy-apatite paste (made at location by mixing powder with chlorhexidine or sterile water) and milled and sterilized Portland Cement. 12. Once the canal is full, clean out the calcium from the threads (usually the blood is oozing thru by then). 13. Screw in an inert plug. Plugs of different materials have been used, ranging from hardware store drain plugs made of PVC, ABS, polyoxymethylene to even brass. Sizes have ranged from 5-50mm diameter. The blood acts as a lubricant when the plug is screwed in. It also forms a nice clot next to the plug to aid in the formation of the dentin bridge. 14. The plug needs to be recessed at least 10 to 20 mm so tusk repair material can be placed over it. 15. Then flush the small remnant of the pulp canal with Chlorhexidine 1%. 16. Flush again with saline solution. 17. Let dry (if needed, use a hair dryer) 18. ETCHING: rub an etching agent on the dentin wall of the pulp canal (cotton-tip) for max. 15-30 seconds. 3M™ Scotchbond™ Universal Etchant Etching gel is a good choice; phosphoric acid (H3PO4 37,5%) or hypochlorite (NaOCl 3%, bleach) are alternatives. 19. Let dry again; use a hairdryer to reduce drying time. 20. Close the pulp canal with glass-ionomer cement. Fill the canal, but leave 5 mm for the composite. 21. Clean the cut-off side of the tusk or sand it with sandpaper. 22. Apply etching (15-30 sec), rinse again and apply bonding for composite application (e.g. Scotchbond Universal Etchant Etching gel (H3PO4 37,5%) and Scotchbond Universal Adhesive). 23. Cover the cement and surrounding area with a layer of dental composite self-curing or light curing, depending on availability of UV-light source (e.g. Tetric Evo Ceram/MIRIS/Filltek/…). 24. Cover the entire cut-off side of the tusk with epoxy glue for extra protection. 25. The tusk will then gradually wear down and the time the wear reaches the plug, the dentin bridge will have formed. At that point, the plug will usually pop out and if needed the hole can be again filled with your favorite tusk repair material. Final stage of tusk repair showing 5 different layers Photo gallery Right tusk fracture in a 5-yr old Asian elephant bull. No attempts to fill the pulp canal were done, resulting in a bacterial pulpitis. By daily cleaning and flushing the deeper part of the pulp remained healthy, resulting in continuous growth of the tusk for at least 2 years. In the end, t he conservative treatment resulted in complete necrosis of the tusk Tip of the fractured tusk and pulp of the same 5 yr-old Asian elephant bull Left tusk fracture in the same 5-yr old Asian elephant bull with the same development course as the left tusk (tusk necrosis) Tusk fracture in a 9-yr old Asian elephant bull that was successfully repaired. See also case report. Tusk fragments of the same 9 yr-old Asian elephant bull. Tip of the tusk after it was sawn off using a giggli wire. During sawing off the tusk tip, the sulcus was lifted using an elephant hook in order to prevent the giggli wire cutting into the skin. When the tusk tip was removed, a large crack filled with dirt became visible. A 'Dremel' hand tool with an extension cable was used to clean out the dirt from the crack. The diameter of the pulp canal was approximately 5 mm. Hence, no threaded rod was used to fill the pulp canal. Etching of the inner tusk wall was done by swapping hypochlorite on the inner surface. The hypochlorite was rinsed off with saline solution. After a layer of calcium hydroxy apatite was applied on top of the pulp, the pulp canal and the cleaned crack were filled with glassionomer cement, the tusk surface was sealed with dental composite. Despite the fact that it was still growing, this tusk of a 26-yr-old Asian elephant bull was extracted several years after it was fractured. These photos show the completely sealed pulp canal by a naturally forme dentin bridge. The decision to extract the tusk is questionable. List of equipment Instruments G iggli wire + handles Plyer to cut off the gigli wire Dremel + extension cable 20 ml syringes (1, 2, 5, 20 and 40 ml) Nail brush Tooth brush Surgical tool set: scissors (1 curved 1 straight) surgical clamps scalpel handle no.3 + blade no.11 sharp spoon Sterile dishes to prepare dental restauration products. Hair dryer High-speed cutter/acetabulum reamer Electric drill Drills (several diameters) Thread makers (several diameters) Threaded (nylon) rods (several diameters) . See text above) Disposables Surgical gloves Surgical suit Surgical drapes Steri le cotton tips Betadine solution (10%) Betadine scrub Ringer's lactate (or Saline solution) Chlorhexidine 1% Etching products: 3M™ Scotchbond™ Universal Etchant Etching gel or Hypochlorite (3%) Calcium hydroxy-apatite powder (to make a paste) or milled and sterilized Portland cement or calcium hydroxide. Glass-ionomer cement Bonding fluid for dental composite (light curing) Dental composite 2-component epoxy or acrylic resin Cotton tips Non-sterile cotton gauze patches (10x10) to cover the eyes Sterile cotton gauze patches (10x10) Leucoplast Duct tape Sand paper Equipment list Avenir Light is a clean and stylish font favored by designers. It's easy on the eyes and a great go-to font for titles, paragraphs & more. References: Woody A.D., Fagan D.A., Oosterhuis J.E. 2022: Large mammal dental surgery. In: Surgery of Exotic Animals, First Edition. Edited by R. Avery Bennett and Geoffrey W. Pye. © 2022 John Wiley & Sons, Inc. To page top

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

Abdominal surgery for male castration, laparoscopic ligation of the ovarian pedicles, umbilical hernia and abdominal surgery to treat repeated colics are described here. To procedures Surgery Surgery in elephants usually follows the rules that are applicable to surgery in horses. Standing sedation with or without local anesthesia is required for minor procedures, while general anesthesia (in lateral or dorsal recumbancy) is required for large procedures (such as abdominal surgery) and if standing sedation poses a risk for the surgeons. See for anesthetic procedures the anesthesia page . Abdominal surgery in elephants Access to the elephant's abdomen is restricted to a relatively small area between the last ribs on the cranio-dorsal side, the hind leg on the caudal side fusing together in the ventral midline. Indications for abdominal surgery described in the literature are limited: cesarean sections have all resulted in the death of the dam. However, a dorso-lateral approach has been used for castration of male African and Asian elephants (Fowler 1973, Byron 1985, Fourner 1994). Fourteen male African elephants (12–35 years old) were anesthetized with etorphine and supported in a sling in a modified standing position, and positive pressure ventilated with oxygen (Rubio-Martinez 2014). Anesthesia was maintained with IV etorphine. Vasectomy was performed under field conditions by bilateral, open‐approach, flank laparoscopy with the abdomen insufflated with filtered ambient air. A 4‐cm segment of each ductus deferens was excised. Behavior and incision healing were recorded for 8 months postoperatively. Successful bilateral vasectomy (surgical time, 57–125 minutes) was confirmed by histologic examination of excised tissue. Recovery was uneventful without signs of abnormal behavior. Large intestine lacerations (3 elephants; 1 full and 2 partial thickness) were sutured extracorporeally. One elephant that was found dead at 6 weeks, had no prior abnormal signs. Skin incisions healed without complication. Laparoscopic ligation of the ovarian pedicles has been performed in free ranging African elephants (Stetter 2004). A specially designed 90 cm long operating laparoscope was used to reach for the ovaries. An umbilical hernia was diagnosed in a 2-wk-old Asian elephant (Elephas maximus) by physical and ultrasonographic examinations (Abou-Madi 2004). Umbilical herniorrhaphy was elected because the defect was large (approximately 7 cm long and 10 cm deep) and could potentially lead to incarceration of an intestinal loop. General anesthesia was induced with a combination of ketamine, xylazine, and diazepam and maintained with isoflurane in oxygen. The hernial sac was explored and contained fibrous tissue, fat, and an intestinal loop but no adhesions. The hernial sac was resected and the body wall closed using the technique of simple apposition. Following a superficial wound infection, the surgical site healed with no further complications. There is one anecdotal report on successfull abdominal surgery in a 14-month-old African elephant suffering of repeated colics ( click here for the case report). References Abou-Madi, N., Kollias G.V., Hackett R.P., Ducharme N.G., Gleed R.D., and Moakler J.P. 2004. Umbilical herniorrhaphy in a juvenile Asian elephant (Elephas maximus ). J. Zoo & Wildl. Med35(2): 221–225, 2004. Byron H.T., Olsen J., Schmidt M., Copeland J.F. and Byron L. 1987. Abdominal surgery in three adult male Asian elephants. J. Am. Vet. Ass. 187, 11. Foerner J.J., Houck R.I., Copeland J.F., Schmidt M.J., Byron H.T. and Olsen J.H. 1994.Surgical castration of the elephant (Elephas maximus and Loxodonta africana ). J. Zoo & Wildl. Med. 25 (3), pp 355-359. (Click here for summary) Fowler M .E., Hart R. 1973. Castration of an Asian elephant, using etorphine anesthesia. J. Am. Vet Ass 163, 6. Rubio- Martinez L.M. Hendrickson D.A., Stetter M., Zuba J.R. and Marais H.J. 2014. Laparoscopic Vasectomy in African Elephants (Loxodonta africana ). Veterinary Surgery 43 (2014) 507–514. Stetter M.D. 2004. Laparoscopic surgery in elephants. Int. Elephants Res. Symp. Fort worth, Texas. December 2-5, 2004 Non-abdominal surgery in elephants Surgical procedures not associated with open access to the abdomen are more common. Despite the enormous healing capacity of the elephant skin, wound healing often takes place per secundam, because it is hard to protect the sutured wound against negative mechanical and biological influences. However, even large wounds (like in vaginal vestibulotomy ) will heal completely per secundam, leaving at most a 2 mm fistula ( click here for wound healing in vaginal vestibulotomy). Trunk injuries are hard to repair because of the extreme mobility of this organ. Many attempts to suture large perfortaing trunk wounds have have failed or at best resulted in partial adhesion of the sutured sites. Repair of a perineal hernia has been described ( click here to read this case report).

Body condition assesement is a method to make an realistic estimate of the physical condition of an elephant. Two different scalings are used, one ranging from 1-10 and one from 1-5. To clinical examination Body condition assessment Monitoring the physical state of an elephant presents a critical aspect in preventive elephant healthcare. It not only allows to quantify the body condition at a certain moment, but also helps to evaluate the effect of changes in management and nutrition when determined before and after implementation of the modification. Several systems to assess the body condition of elephants have been developed. It is important to evaluate the condition of the elephant at your first visit. The scoring system used in this document is based on the extent of visibility of depressions around bone structures when an elephant is viewed laterally. Depressions around bones become prominent as an animal loses its subcutaneous fat deposits and muscles in the region concerned, thereby making bones appear more prominent. This system use scores of 1-10, where 1 represents an extremely poor (cachectic) and 10 extremely fat (obese) bodycondition. An assessment that uses a 5-points scoring system can be found here . The table below shows the characteristics for each score in this 1-10 system and the selection path to be followed to determine the score that matches with the elephant. Examples of body condition scores (1-10) in Asian elephants are shown in the photos below. The most important reference poinst are the forehead, backbone (vertebral column), scapula, ribs and pelvic bone (ilium). Instead of using photos, the body condition can also be compared with drawings. For this purpose, Schiffmann et al. drew detailed pictures from both African and Asian elephants (2): Example drawings used for body condition scoring of Asian elephants (drawings by Jeanne Peter) Example drawings used for body condition scoring of African elephants (drawings by Jeanne Peter) References: 1. Morfeld K.A., Meehan C.L., Hogan J.N., Brown J.L. (2016) Assessment of body condition in African (Loxodonta africana) and Asian (Elephas maximus) elephants in North American zoos and management practices associated with high body condition scores. PLoS ONE 11:0155146. 2. Schiffmann C., Clauss M., Fernando P., Pastorini J., Wendler P., Ertl N., Hoby S.J-MHatt. (2018) Body condition scores of European zoo elephants (Elephas maximusand Loxodonta africana): Status quo and influencing factors. Journal of Zoo and Aquarium Research 6(3) 2018 To page top

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

This page contains the links to official documents of the European Association of Zoos and Aquaria (EAZA) and the American Association of Zoos and Aquaria (AZA) with regards to elephant management and disease control: - EEHV-monitoring of calves -EEAV-treatment protocol -EEHV-antibody monitoring -Necropsy forms -Reproduction guideline -TB-recommendations -Elephant transport protocol -Vaccination advice -How to make a blowpipe -How to make a mouth opener Best Practice Guideline EAZA EEHV EEHV monitoring calves EAZA EEHV treatment protocol EAZA EEHV antibody monitoring EAZA Necropsy form EAZA Necropsy form AZA Reproduction guidelines EAZA TB recommendations USA TB recommendations EAZA Transport protocol EAZA Vaccination advice EAZA EDV Tusk fracture guideline How to make a blow pipe syringe How to make a mouth opener Urine collection for progesterone Hematology and urinalysis manual Elephant Care Manual (FAO) Musth management and Care Hand-raising manual Documents

ElephantMedicine.info provides information about infectious and non-infectious diseases in elephants, case reports, clinical and laboratory procedures and management guidelines. -Elephant Medicine -Diseases -Treatment -Anesthesia -Dental problems -Tusk repair -Laboratory tests - Hematology -elephant disease treatment -dental problems -locomotion -diagnostic procedures -physical examination -laboratory procedures -reproduction and birth procedures -anesthesia procedure -lameness -pedicure Elephant Medicine Get started A website for veterinarians taking care of elephants Sharing your clinical experience in ELEPHANT MEDICINE with elephant clinicians worldwide This website provides information about elephant diseases, procedures and recommendations. It also includes a collection of clinical case reports, submitted by veterinarians from all over the world. You will find cases that have been successfully treated, but also cases that had a sad ending. Some basic disease information is provided as well as descriptions of clinical and laboratory procedures. The information on this website is subject to changes, as our knowledge about diseases in elephants will grow by sharing our experiences. Veterinarians involved in elephant medicine are encouraged to submit reports of their experiences. Together we can create a platform for the current and next generations of veterinarians. For the benefit of an amazing, but vulnerable species: the elephant. Disclaimer: The content of this website, such as graphics, images, text and all other materials, is provided for reference and educational purposes only. The content is not meant to be complete or exhaustive or to be applicable to any specific individual's medical condition. The information is based on scientific literature, open source information and anecdotal information provided by trusted sources. You assume full responsibility for using the information on this site, and you understand and agree that the authors are not responsible or liable for any claim, loss or damage resulting from its use by you or any user. Contact

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Kidney disease is quite common in elephants. The analysis of urine is important. This page describes the routine urinalysis and some specific features related to elephant urine. Protein detection can be done by refractometry and sulfosalicylic acid (SSA)precipitation. Comparing osmolarity in serum and urine gives an indication for the presence of kidney failure. To lab diagnosis Urinalysis The routine urinalysis is a quick and relatively inexpensive test which can be readily performed in a field laboratory. Ideally urine should always be collected at the same time as blood for hematology and clinical chemistry (if possible) and before any treatment is administered. Urine should be examined as soon as possible after collection, because artifacts will occur in the urine over time (cells lyse, crystals form in vitro). If a delay is anticipated before analysis, the urine should be refrigerated. Refrigerated urine should always be brought to room temperature before testing. A complete urinalysis consists of 1) evaluation of physical characteristics, 2) measurement of specific gravity, 3) dipstick analysis and 4) examination of sediment. Equipment and Materials Microscope Refractometer Slides Urinalysis dipsticks Urine sediment stain Physical Characteristics of Urine 1. Evaluate and record urine color Normal mammalian urine is yellow to amber. Urine volume and concentration affect the depth of the color. Some examples of various urine colors and corresponding common possible causes: 2. Evaluate and record urine turbidity Fresh urine is clear to very slightly cloudy. Urine may become more cloudy if left standing or if refrigerated. The turbidity of the urine sample is reported as: clear, slightly cloudy, cloudy, opaque, or flocculent. Excess turbidity results from the presence of suspended particles in the urine. The cause can usually be determined based on the results of the microscopic urine sediment examination. Common causes of abnormal turbidity include: increased cells (RBC, WBC) numerous crystals bacteria lipiduria (lipids often rise to the surface) mucus (especially in horses) semen fecal contamination 3. Odor Ammonia is formed from urea by bacterial action. An ammonia smell may be prominent in retained or old urine samples. An acetone small may suggest ketosis. Some drugs may impart a characteristic odor. 4. Volume Urine volume may be estimated from urine specific gravity. In general, volume and specific gravity (S.G.) are inversely related in health and in most diseases. Exceptions include: Diabetes mellitus. Polyuria and high S.G. coexist because of glucosuria. Acute and chronic renal disease. Oliguria (↓ urine volume) may be accompanied by a lack of renal concentrating ability. 5. Measure specific gravity (S.G.) Specific gravity is the ratio of the refractive index of urine compared to water. Refractometry is the easiest method to measure urine S.G.Dipsticks are not as accurate. Glucose and protein may falsely ↑ urine S.G. Knowledge of the hydration status is necessary to interpret the S.G. Urine S.G. can vary from 1.001 to 1.065 in most healthy animals but this range also includes values associated with renal abnormalities. S.G. has been inadequately studied in elephants and obtaining a baseline during health and evaluating sequential samples during illness is recommended. Isosthenuria (fixed S.G.) is the constant maintenance of urine S.G in the range of the glomerular filtrate (1.008-1.012). Isosthenuria means the kidney is neither concentrating nor diluting urine. Hyposthenuria is a S.G. < 1.008. In hyposthenuria the kidneys still have some water-balance function. 6. Perform dipstick analysis and record results. Mix the urine Dip the dipstick; remove extra urine Start timing Compare to chart on bottle in good light The dipstick method is not reliable for protein detection in elephants. Using the SSA-method is more reliable (see 6.1) 6.1 Urine protein detection. "PROT-SSA" represents the reaction observed on the sulfosalicylic acid (SSA)precipitation test. The SSA reagent is added to a small volume of urine. Acidification causes precipitation of protein in the sample (seen as increasing turbidity), which is subjectively graded as trace, 1+, 2+, 3+ or 4+. Unlike the "PROT-STIX" test, the SSA reaction will detect albumin andglobulins (although it is more sensitive to albumin). In addition, the SSA detects Bence-Jones proteins, although it often underestimates them. In alkaline urine, the SSA reaction is a more accurate measure of urine protein content than the dipstick. The most accurate measurement of urine protein output is measurement of urine protein excretion over 24-hours. False positives Contrast media Antibiotics in high concentration, e.g. penicillin and cephalosporin derivatives Uncentrifuged turbid urines can look positive. Therefore, SSA should always be performed on urine supernatant. False negatives Highly buffered alkaline urine. The urine may require acidification to a pH of 7.0 before performing the SSA test. Dilute urine Turbid urine - may mask a positive reaction. https://www.klimud.org/public/atlas/idrar/web/www.diaglab.vet.cornell.edu/clinpath/modules/ua-rout/protssa.htm 7. Urine sediment examination Centrifuge a fresh urine sample for 5 minutes at about 1500 RPM Remove supernatant Add 1-2 drops of stain to sediment Tap the bottom of the tube to mix Transfer one drop to a slide and place a coverslip Examine the entire coverslip at 10X and 40X * Low maginification (10x): casts, large crystals, debris, parasitic ova are visible * High magnification (40x): leukocytes, erythrocytes, epithelial cells, fat droplets, small crystals, sperm, debris and bacteria are visible To page top

This case report describes a colic episode in an adult Asian elephant. Salmonella sp. was identified in a fecal sample taken during this period of colic. - colic -Salmonella To colic general information CAse report Colic and Salmonellosis in an adult Asian elephant Date: Place: Data provided by: History This adult multiparous female had been on GnRH-vaccine for over 4 years. Because of chronic joint disease, the elephant had been on phenylbutazone for over 1 year in combination with omeprazol. Sudden onset of apathy, anorexia and hardly drinking water. Normal feces. Occasionally the elephant goes into a sitting position or lateral recumbancy, showing moderate straining activities. After this labor-like behaviour, herd mates investigate the perineal area of the elephant with their trunk. Differential diagnose: Labour, colics (intestinal, uterine or urinary) Treatment The administration of phenylbutazone was discontinued No specific treatment was given on the first day. Treatment results During the night the elephant became more active and the symptoms decreased. The next morning, the animal behaved normal. Diagnostic notes Salmonella sp. was cultured from the feces on the day it showed the above mentioned symptoms To page top

This page describes the normal birth process in elephants, including progesterone and ultrasound monitoring, preparation, list of requirements for veterinary assistance, transrectal massage to induce the Ferguson reflex and post-partem care. To reproduction Normal birth process This chapter is based on the EAZA Guidelines for the reproduction-related management of female elephants . Taken care at the birth process means that the pregnancy has been confirmed. More detailed information is provided in the chapter: pregnancy confirmation. After a pregnancy period of 87- 95 weeks (610- 670 days, 20-22 months), parturition is sometimes announced by subtile changes in the behaviour of the elephant. Twentyfour hours observation including the use of a (time-lapse) video recorder starting in week 85 may add to information about relevant events prior to parturition: night pacing, kneeling down, climbing, short periods of separation from the group, beating the vulva with the tail, frequent production of small-sized feces and small quantities of urine, loss of mucous plug, playing with mucous plug, rupture of the allantois sac, labour waves. Preceding any recommendation in this chapter, the following remarks need to be made: It is a myth to think that a multiparous elephant does not need to be monitored and assisted during parturition according to this protocol. Too many calves have been born dead or very weak because of the fact that parturition had started unnoticed and stopped unnoticed. In our view, this protocol should be followed in ALL cases of elephant parturition as much as the elephant management allows. It is a also myth that oxytocin is a harmless drug to be used in elephant parturitions. In at least 3 cases the administration of oxytocin has been associated with the occurrence of an uterus rupture because of insufficient cervix dilatation. It should never be used without ultrasonographic examination of the cervix uteri. The visible presence of the allantois sac as a subcutaneous bulging mass under the tail is not a guarantee for a sufficiently dilated cervix. Many elephants, including very experienced multiparous elephants may just stop the parturition process, often unnoticed. Hypocalcemia is one of the main causes for this phenomenon to happen.This will compromise the health of the calf. Don’t relay only on what you see from the outside! For a proper judgment about the presence of labor activities or the progress of parturition, your professional judgment should be based on the results of progesterone tests and ultrasonographic examinations. If you do not believe in the above mentioned statements, you may find yourself confronted with a dead elephant calf or a dead elephant mother or both. So, discuss this item with your staff and make your decision before you start breeding your elephants. If the management of elephants in your zoo does not allow blood sampling or ultrasonographic examinations, you must be aware of the risks associated with a silently interrupted birth process, no matter the reproductive experience of the animal involved. Safety for the personnel should never be challenged by unacceptable risks. Predicting the time of parturition, measures and observations: Daily blood sampling from week 89: every other a day progesterone assay, until week 91 (637 days). From week 91 (637 days): daily assays and as soon as progesterone starts to decrease: sample twice daily. Daily monitoring of progesterone is only possible if you have a nearby facility that runs these assays on a daily basis. Find out from your nearby hospital in the early stage of pregnancy! Many veterinary labs do not have tests that are sensitive for levels of progesterone (P4) in elephants 1-3 days prior to delivery. Also check the availability during the weekend. NOTE: a decrease of progesterone to baseline level is not always occurring. A 23-yrs-old Asian elephant at the Rotterdam zoo (with a baseline progesterone level < 0.6 nmol/l) delivered a healthy calf while her progesterone remained above 0.65 nmol/l. Progesterone plasma concentration during pregnancy. After a gradual decrease in the last months of gestation, progesterone drops to baseline level a few days before parturition. Note that the absolute height of the progesterone concentration depends on the assay used! Different assays measure different metabolites = different levels! Signs of an approaching parturition may include: ·Loss of mucous plug (not seen in many facilities) ·Pre-and post-parturition ventral edema may be noticed. ·Group members may react differently (vocalizations, restlessness) ·In most cases (75%) rupture of allantois sac and loss of allantois fluid (looks like urine) is seen within 2 hours prior to birth. ·The size of fecal balls may get smaller towards the end of pregnancy. ·Frequency of urination may increase around parturition, resulting in more “watery” consistency of the urine, resembling more like allantois fluid . ·Development of mammary gland and production of milk shortly before birth is often seen, however this may also occur in a much earlier stage of pregnancy . Milk accumulation can be visualized using transcutaneous ultrasonography several hours prior to parturition. ·Softening of the pelvic ligaments (due to estrogen surge) may result in slightly abnormal locomotion of the hind legs. Parturition/Preparations for calving: ·Training and/or desensitizing of the pregnant elephant for veterinary intervention, like blood sampling, injections, IV-infusions, milking and rectal manipulations ·If possible, store some colostrum (freezer) or store plasma obtained from the dam in weeks prior to parturition. ·Have artificial milk available (Salvana GmbH, Germany; hand raising has been done at Emmen Zoo and Berlin Zoo) ·Check restraint chains and fixation points for the legs and one extra fixation point between the hind legs for pulling devices. Soft ropes for pulling the calf away if needed should be available. The use of a calf harness has been described. ·Check the stable and place bars where a calf could possibly escape. Block all possible escaping routes for the calf (not for staff!!). ·Take out all obstacles. ·Be prepared for closing the elephant house for the public (sign post, etc.) ·Make sure there is a good stock of commercial cat litter or saw dust to be used on a concrete floor as soon as the calf is born. This will absorb much of the allantois fluids and prevent the animals from slipping on the wet floor. ·2 or 3 pairs of keeper-gloves (soccer) to get a better grip on the wet, slippery calf when needed ·Plastic hose pipe (with pump, if necessary) for rectal cleaning with lukewarm water ·3 birth-chains with proper handles (2 for the legs, 1 for trunk or tail); find a way to avoid back sliding when manual extraction (vaginal vestibulotomy) is required. ·Drugs to be kept in store: ·Ca-borogluconate for I.V. infusion ·Estradiol gel (EstroGel® 0.06%) ·Oxytocin ·Lidocain ·Xylazine, detomidine or medetomidine ·Butorphanol ·Azaperone ·Atipamezole ·Doxapram ·Oxygen ·(Betadine®-)iodine solution for navel disinfection (umbilical infection is a major cause of perinatal complications) ·Lubricant (many liters). J-lube®, a concentrated lubrication powder, has proven to be very useful. The normal calving process should take place within 2 hours after rupture of membranes (release of fetal fluids). If the plasma progesterone level decreases to below base line concentration, this is a signal that calving should take place within 48 hours. Prolonged intervals have been reported (as long as 14 days) still resulting in the birth of a living calf, but it is very likely that this is due to a disturbance of the normal birth process that should be corrected before the health of the calf is jeopardized. There is one report of a parturition taking place without a complete drop to base line level. The calving process is a natural process. Elephants should give birth in their own social environment, excluding any external disturbing factors (e.g. noisy building in the neighborhood, visits of unknown individuals, etc.). With a well trained animal, blood sampling and ultrasonographic examinations can be done while the animal is temporarily separated from the group. Immediately afterwards, the animal should return to its group. All efforts should be made to make sure that the calf is born in the group while the mother is NOT chained! This will stimulate the acceptance of the calf by the mother and group members and is an investment for future breeding successes for the entire elephant group. To determine the right moment when calving starts, 2 parameters are essential: - The progesterone blood level - The relaxation of the cervix, monitored by ultrasonographic examination. Progesterone: the sensitivity of the equipment and the time needed to run the assay are the bottleneck for using the progesterone concentration as a reliable tool. Today many human hospitals use advanced equipment with a very low detection level that can provide results in less than 2 hours. Make sure that you have made arrangements with a lab long before you expect the parturition. Ultrasonography: to use this technique as a reliable tool, it is indispensable for the veterinarian to gain experience long before the elephant birth is expected. This will enable the veterinarian to distinguish a normal cervix from the relaxed cervix (figure 4a + b) from the normal cervix. Preferably a 3.5 MHz probe should be used transrectally. Longitudinal, transrectal ultrasonographic image of the vagina and the closed cervix of a pregnant e lephant. The same image as with the vagina and cervix indicated with white lines. Transversal, transrectal ultrasonographic vagina and cervix of a pregnant elephant. Allantois sac with cloudy fetal fluid in the (partly) dilated cervix of an Asian elephant 12 hours prior to delivery (transverse section, white arrows: allantois sac, open arrow: pelvic bone) The same image with the vagina and the closed cervix indicated by white lines. Note the folds in the cervix uteri. Allantois sac with cloudy fetal fluid in the (partly) dilated cervix of an Asian elephant 12 hours prior to delivery (longitudinal section, white arrows: allantois sac, open arrow: pelvic bone) During the last 2 weeks of gestation, the mucous that is present in the vagina during gestation will be discharged gradually. This is a clear indication for a pending parturition. Recognizable onset of parturition occurs normally within 24-48 hours after progesterone has dropped to below base-line level. In these guidelines the absence of visible signs of parturition in the same time frame is considered an abnormal condition; this status requires veterinary intervention. At this point 2 situations may occur: the parturition process has started but has been interrupted without or with spontaneous rupturing of the allantois sac. Interpretation of findings and action to be taken 1. No rupture of allantois sac noticed If the calf is not born by natural way 24 hours after blood progesterone has dropped to baseline level, rectal palpation and ultrasonographic examination of the cervix is highly recommended. This will demonstrate the rate of relaxation of the cervix, the presence of the allantois sac or parts of the fetus in the cervix or vagina and should be repeated at least every 8 hours. A blood sample should be taken to measure the calcium level. If below 2.5 Mmol/l, calcium should be administered as an IV-infusion (NB: when given in an ear vein, it should be given strictly IV in order to avoid damage to the vein) or orally (suggestion: calcium syrup concentrate for human use, enveloped in the carton core of toilet paper, covered and sealed with fresh tamarind paste has worked well; most elephants will eat it, including the carton material). The effect of the calcium administration should be confirmed by rectal palpation (increase of contractibility of the uterus) and determination of the blood calcium level. Store an EDTA and heparine sample for herpes virus diagnostic purpose (both cells and plasma in freezer after separation). Transrectal ultrasonographic examination at 48 hours: A. No relaxation of the cervix at 48 hours: search for calf movements and nail position of the fetus (palpation and ultrasound) and blood flow in fetal vessels (ultrasound). Transcutaneous ultrasonographic examination (both flanks have a small ”window” where visualization of fetal movements may be seen) may help to determine the status of the fetus. Apply estrogens rectally. Good results have been obtained by the rectal and transdermal (perineum) application of an estradiol containing estrogen gel (Estrogel 0,06%, total dose 700-800 mg estradiol). The effect on the cervix dilatation should be monitored closely by transrectal ultrasonography! At this time, at least 1 hour after the local application of estradiol, rectal massage should be applied to test and stimulate the contractibility of the uterus. Technique: remove feces from rectum, flush out the rectum, use abundant lubrification, keep both gloved hands (NB: the rectal mucosa is vulnerable due to estrogens) with the fists joined in a firm grip and press with the wrists or the dorsal sides of the hands against the pelvic ring to stimulate the pelvic receptors until strong labor waves appear or at least 10 minutes. When labor waves occur, continue this massage for 3 hours (if needed change operator). Check regularly by means of ultrasound the condition of the cervix. If there is still no cervix relaxation, continue monitoring the viability of the calf. If no fetal parts can be detected, consider the presence of pseudopregnancy (ovarian tumor, dysfunction of the pituitary gland, etc.). The application of estradiol gel (total dose: 400-500 mg 17-β-estradiol) as described above may be repeated 3-4 hours after the first treatment if the cervix dilatation is still incomplete. B. Partial or complete relaxation of the cervix at 48 hours or later as a result of the situation described under 1A: Apply rectal massage to test contractibility of the uterus. If limited or no reaction, the administration of oxytocin is contra-indicated. In this case, the administration of calcium is recommended (even when blood calcium level is within normal ranges). After 2 hours the use of estradiol as described under 1.A is recommended. Only if uterus contraction can be provoked by the rectal massage, the use of oxytocin may be considered using the following dosage: 25-50 IU oxytocin s.c. or i.m. (if needed use a blow dart). Oxytocin should be used with care, as it may dramatically exhaust the contractibility of the uterus muscles as well as the general condition of the female. There might also be the risk of reduced blood circulation in the umbilical chord, due to the spasms in the myometrium. Prostaglandine E (dinoproston) has been used on a few occasions for cervix dilatation (after the administration of estradiol) and to stimulate uterus contractions. As there is still limited knowledge of its efficacy after transrectal administration and the risk of stormy uterus contractions, it should only be used when any obstructions or abnormalities of the calf can be ruled out. Depending on the progress obtained, rectal massage and the administration of oxytocin are the 2 major treatments to follow from this point. In between these treatment events, the animal should be exercised to relieve the pain and stimulate position changes of the calf and preferably it should be kept in the group. Only when the animal cannot be separated whenever required, the cow should be kept separated from the group, but with as much physical contact as possible. Oxytocin should be given in intervals of at least 2 hours for a maximum of 12-24 hours under the guidance of ultrasound to evaluate tAhe progress. Continue this approach of treatment until parts of the calf have entered the pelvic cavity. If the efforts remain unrewarded and no access to fetal parts is possible, not much can be done. Continue monitoring the viability of the calf. If the calf has died, while the membranes are still intact, the risk of intoxication is limited, but immunosuppresion could be a complication for the cow. To date, no proper data are available. The dosage of oxytocin may only be increased to 100 I.U. after parts of the calf have entered the pelvic area and progress is clearly observed. At this time, a bulge containing parts of the body under the tail of the dam should be visible. Progress of parturition must be monitored strictly at this stage. If this increased dose of oxytocin does not result in parturition a vaginal vestibulotomy should be performed soon to get better access to the calf. See next chapter. Expulsion of the calf should follow soon after the bulge appears under the tail of the dam. The allantois sac usually ruptures during the (induced) passage through the pelvic canal. One complication described at this stage, is reduced passage space as a result of edema in the urogenital canal resulting in a “catching effect” of the head and/or shoulder of the calf inside the soft part of the distal (vertical part) genital tract. Suffocation of the calf is a realistic complication. This condition has been observed several times in primiparous elephants of more advanced age. Elephants in this category should be prepared by the local application of J-Lube in the distal part of the vaginal vestibulum and massage of oestrogel/creme in the skin between anus and vulva. If this condition occurs, quick intervention is required by applying firm manual pressure from the outside on the calf in the sliding direction of the calf. Be aware of the risk of kicking by the mother. 2. Ruptured allantois sac A significant event in the parturition process is the rupturing of the allantois sac, which – when intact - acts as a hydraulic dilatator for the cervix, a natural lubrication for the dam and a pressure protection for the calf. NB: The amniotic sac that covers the body directly, usually remains intact during the expulsion of the calf and ruptures during the final passage through the birth canal and is actively removed by the dam. NB: a chained dam, may not be in the position to remove these membranes, possibly resulting in suffocation of the calf. Differentiation between urine and fetal fluids is extremely difficult; smell, creatinin test strips and possibly protein concentration could be helpful. If no progress in parturition is observed, major complications should be considered, like a dead calf, malposition of the calf (which is often dead), oversized calf, malformation and twin pregnancy. Because of the urgency of this situation, the calf should be born within 2 hours after rupture of the allantois sac and loss of allantois fluid. If not so, veterinary intervention has to take place. Two situations may occur: No fetal parts positioned in the pelvic area: treatment should aim on the urgent relaxation of the cervix. Calcium status should be determined and treated accordingly (see above). The further approach is according to 1A, however the situation is more critical for both the dam and the calf. Fetal parts have entered the pelvic area: Calcium status should be determined and treated accordingly (see above). Ultrasound is essential to determine which fetal parts have entered the birth canal, determine the position of the calf (visualization of the nails, posterior or anterior position, number of nails, trunk) and viability. Malposition (e.g. only one leg in birth canal, no head while in anterior position) is an indication for vaginal vestibulotomy or fetotomy. If no abnormalities are found during ultrasound, 50-100 I.U. of oxytocin should be given i.v. or i.m. and rectal massage should be practiced. Birth should be completed within 1 hour. Other drugs used Uterine laxants have been used in elephants on rare occasions. There are some anecdotal reports about the use of denaverinehydrochloride (Sensiblex®, Veyx) at a dose of 0,04 – 0,05 mg/kg BW (i.m.). Isoxsuprinelactate (Duphaspasmin. Fort Dodge Animal Health, 6290AA Vaals, The Netherlands) was used in a fetotomy case at 0,15 mg/kg BM (i.m.). Carbetocine (Depotocin®, Veyx), a long acting oxytocine has been used in 3 occasions dosage (0,09 µg/kg BW) i.m. Presentation of the calf A study in 46 elephants demonstrated that the overall ratio between anterior and posterior presentation was 12:34. In dystochia cases the situation was quite different: 6:3. In normal birth procedures the ratio was 6: 29. Anterior presentation has a higher risk of dystochia than posterior presentation of the calf. (Ilic D. et al. 2021. The incidence of anterior and posterior presentation at birth in Asian (Elephas maximus ) and African elephants (Loxodonta africana ): A Review Study. Indian Journal of Animal Research.DOI: 10.18805/IJAR.B-1319) It has been observed that the calf may rotate during the final phase of the expulsion, similar to what happens in horses and cattle (see video). This may be important in case of a dystocia, if the calf has remained in complete 'horizontal position' and a vaginal vestibulotomy is indicated. Note that the calf has rotated during the final phase of the expulsion. Transrectal massage: Transrectal massage is a very effective method to stimulate uterine contractions if the uterus is prepared for it . Certain conditions (especially hypocalcemia) may impair the contractions. This should be solved first. Stimulation of the pelvic wall by rectal massage is known as the Ferguson reflex . Transrectal massage of the pelvic wall is best performed by firmly pressing the closed fost against the roof and sides of the pelvic cavity and on the dorsal side of the vagina making the movements as shown in this video. Post-partum care Disinfection of the navel with Betadine® iodine is strongly recommended (if the mother allows its application). The afterbirth usually comes off within 12 hours. There are a few reports on retained (parts of) placenta for several weeks, without major complications for the dam. Hygienic measures should be applied to reduce the infectious burden for the calf. Be aware of the fact that a second calf can still be present in the dam. There are reports that second calves were born between several hours up to 3 months (the prevalence of twins in elephants is 1:3000). The calf should drink (colostrum) as soon as possible, at least within 24 hours. If not, or when the calf makes a weak impression, the banked serum (or freshly taken serum) should be given to it orally. Try to find out the reason why the calf is not drinking successfully: e.g. too small, weak, painful mammary glands, malbehaviour of the dam. If for any reason the mother is rejecting the calf, lactating herd member can take care of the calf. There is evidence that the mother takes over from this surrogate mother during the first 72 hours. If no lactating elephant is present in the herd, training for bottle feeding should start after 12 hours and continued for 72 hours. If the mother still rejects her calf, the best option for the calf is to move it to another herd with a lactating elephant. If introduction to this herd fails too, bottle feeding is the last option. To date very few bottle-raised elephants have reached the age of puberty. To page top

The most important zoonotic disease in elephants is tuberculosis. Other examples of zoonotic pathogens are cowpox, Salmonalla, Anthrax, and Pasteurella. Several microbes are opporunistic pathogens, like rabies, leptospirosis, foot and mouth disease, and organisms involved in local processes (abscess, feces, urine, exudate). Zoonoses Zoonotic diseases in elephants. Zoonotic diseases are defined as infectious disease of humans caused by a pathogen (an infectious agent, such as a bacterium, virus, parasite or prion) that can jump from a non-human (usually a vertebrate) to a human and vice versa. Some of these pathogens can be considered as opportunistic, others as primary infections. Relevant primary pathogens in elephants are : Bacterial diseases Mycobacterium tuberculosis complex (MTBC). The prevalence of MTBC in captive elephants in European zoos is relatively high. A study of the post-mortem reports between 1985 and 2024 showed that 20/301 Asian elephants and 12/196 African elephants had died of MTBC (Data from EAZA elephant TAG, WS). From 1997 through 2011, the median point of prevalence within the Asian elephant population in USA-zoos was 5.1%, with a range from 0.3% to 6.7%. In contrast, the annual point prevalence during the same time period within the African elephant population was 0. Although exact data about the prevalence of MTBC in range countries are not known, there are many reports of MTBC in captive and to (a lesser degree) wild Asian elephants. Data on MTBC in African elephants in range countries are limited. Click here to read more about tuberculosis in elephants. Non-tuberculous mycobacteriosis: Mycobacterium elephantis ; only found in humans, never in elephants. However, the strain is genetically related to Mycobacterium confluentis and M. smegmatis cultured form lung lesions in an elephant (Lacasse, 2007) . Bacillus anthrax: Click here to read more about anthrax in elephants. Pasteurella multocida: Click here to read more about Pasteurellosis in elephants. Salmonella spp .: Click here to read more about salmonellosis in elephants. Leptospira interrogans found in urine of captive elephants in Asia. This is a potential risk for humans in close contact with these elephants (Athapattu 2019). Click here to read more about leptospirosis in elephants. Viral diseases: Cowpox virus (Orthopoxvirus bovis) . Asian elephants are very sensitive to a pox virus infection, African elephants to a lesser degree. The fluid that fill the pox vesicles are full of virus. Once the fluid is exposed, humans can become infected. Click here to read more about pox virus infections in elephants. Foot and Mouth disease virus (FMD-virus): Asian elephants are very sensitive to FMD. There is only one report of FMD in an African elephant that was experimentally infected. Click here to read more about FMD in elephants. Rabies: transmission of rabies virus from elephants to humans have never been reported, but saliva of diseased rabid elephant is a potential risk for humans. Click here to read more about rabies in elephants. Opportunistic pathogens: Opportunistic pathogens can be found in the environment, but when concentrated in a pathological condition in an animal (abscess, feces, urine, exudate), they can cause disease in humans: Escherichia coli Pseudomonas Bacteroides spp. Staphylococcus aureus Streptococcus spp. Klebsiella spp. Mycobacterium avium Fungi: there are no reports on fungus infections in humans acquired through contact with elephants. Serological responses detected in elephants without evidence of causing disease: Elephants may be (temporary) silent carriers of several potentially pathogenic microbes. The historical contact is expressed by the presence of antibodies in the blood of the elephant. Examples of these conditions are: African horse sickness virus : the presence of antibodies in African elephants has been described (Barnard, 1995). Humans are usually not affected. However, severe disease has been reported in lab workers who were producing a AHSV-vaccine (van der Meyden, 1991; Reid,1991). Influenza type A : 1 serologically positive elephant reported (Schröder 1992). Eastern equine encephalitis: 1 serologically positive elephant reported (Christy, 2009) Bluetongue : antibodies were detected in 7 out of 109 serum samples of captive Asian el ephants in India (Bhat, 1998). Canine distemper: antibodies were detected in 25 out of 144 serum samples of captive Asian elephants in Thailand (Ono, 2006). Yersinia pestis (plague); In one study in wild African elephants 0.3% of the cohort animals were found seropositive for antibodies against Yersinia pestis (Gordon, 1979). No transmission of plague from elephants to humans has been reported. Toxoplasma gondii : 35% of captive elephants in a study in Thailand was serologically positive (Udonsom 2022). As the elephant is not an end-host for toxoplasmosis, transmission of toxoplasmosis from elephants to humans is unlikely to occur. Cryptosporidium spp .: found in African elephants at a European zoo (Gracena, 2002). No transmission to humans has been reported. References Athapattu TPJ, Fernando BR, Koizumi N, Gamage CD. Detection of pathogenic leptospires in the urine of domesticated elephants in Sri Lanka. Acta Trop. 2019 Jul;195:78-82. doi: 10.1016/j.actatropica.2019.04.029. Epub 2019 Apr 29. PMID: 31047864. Barnard BJH, Bengi RG, Keet DF, Dekker EH, Verwoerd DW. 1995. Epidemiology of African horsesickness: antibodies in free-living elephants (Loxodonta africana) and their response to experimental infection. Onderstepoort journal of Vet. Res. 62, 1995. Bhat N, Manickam R, Arunp W.1998. Detection of bluetongue antibody and antigen in Indian elephants, spotted deer and blackbucks. Indian Journal of Animal Sciences 68 (2) : 135, February 1998 Christy L. Rettenmund CL, Terrell SP, Miller M. 2009 Eastern Equine Encephalitis Virus (EEEV) Titers in African Elephants (Loxodonta africana) At Disney’s Animal Kingdom. American association of Zoo Veterinarians Conference 2009 Feldman M, Isaza R, Prins C, Hernandez J. 2013. Point prevalence and incidence of Mycobacterium tuberculosis complex in captive elephants in the United States of America, Veterinary Quarterly, 33:1, 25-29. Gordon DH, Isaacson M, Taylor P. 1979. Plague Antibody in Large African Mammals. Infection and Immunity, Nov. 1979, p. 767-769 Gracenea M, Gómez M., Torres J, Carné E, Fernández-Morán J. 2002. Transmission dynamics of Cryptosporidium in primates and herbivores at the Barcelona zoo: a long-term study. Veterinary Parasitology, 104(1), 19–26. doi:10.1016/s0304-4017(01)00611-2. Lacasse C, Terio K, Kinsel MJ, Farina LL, Travis DA, Greenwald R, Lyashchenko MDKP, Miller M, Gamble KC. 2007. Two cases of atypical mycobacteriosis caused by Mycobacterium szulgai associated with mortality in captive african elephants (Loxodonta africana). Journal of Zoo and Wildlife Medicine 38(1): 101–107, 2007. Oni O, Wajjwalku W, Boodde O, Chumsing W. 2013. Canine distemper virus antibodies in the Asian elephant (Elephas maximus). The Veterinary Record, September 23, 2006. Reid, R, van der Meyden, CH, Erasmus, BJ, Meyer, H and Hamilton, AMP. 1991. Encephalitis and chorioretinitis associ[1]ated with neurotropic African horsesickness virus infection in laboratory workers. Part II. Ophthalmological findings. S Afr Med J 81:454–458. Schröder, H.D., Fischer, M. and Ippen, R. 1992. Contribution to the occurrence of infection of zoo mammals with influenzavirus type A. Erkrankungen der Zootiere. Verhandlungsbericht des 34. Internationalen Symposiums uber die Erkrankungen der Zoo- und Wildtiere, Santander-Spain, pp. 119–125. Udonsom R, Nishikawa Y, Fereig RM, Topisit T, Kulkaweewut N, Chanamrung S, Jirapattharasate C.2022. Exposure to Toxoplasma gondii in Asian Elephants (Elephas maximus indicus) in Thailand. Pathogens 2022, 11, 2. van der Meyden, CH, Erasmus, BJ, Swanepoel, R. and Prozesky, OW. 1991. Encephalitis and chorioretinitis associated with neurotropic African horsesickness virus infection in laboratory workers. Part I. Clinical and neurological observa[1]tions. S Afr Med J 81:451–454. Zachariah A, Pandiyan J, Madhavilatha G, Mundayoor S, Chandramohan B, Sajesh P, et al. Mycobacterium tuberculosis in Wild Asian Elephants, Southern India. Emerg Infect Dis. 2017;23(3):504-506. https://doi.org/10.3201/eid2303.161741 To page top