Peregrine L. Wolff, DVM
Small ruminant species commonly kept outside of a zoological institution include: 1) Cervids: - North American species such as elk (Cervus elaphus sp) white tail deer (Odocoileus virginianus) and reindeer (Rangifer tarandus) or Eurasian species such as sika deer (Cervus nippon), fallow deer (Dama dama) axis deer (Axis axis) red deer (Cervus elaphus) and muntjac (Muntiacus muntjak); 2) Antelope: - scimitar-horned oryx (Oryx dammah), addax (Addax nasomaculatus) blackbuck (Antilope cervicapra) and 3) domestic and exotic sheep and goat crosses - moufflon (ovis musiman) and moufflon crosses (Corsican, Texas dall, black Hawaiian, painted desert, etc.) aoudad (Ammotragis lervia), and goats (angora, Spanish or Catalina) and Ibex (Capra ibex).
Physiology and Physical Assessment
Remember the Rumen
Ruminants eat plant fiber and cellulose. However not all ruminants choose the same type or part of a plant. Depending on feeding strategy, ruminants are generally classified as grazers, browsers or intermediates. The majority of small ruminants are intermediate feeders meaning that they successfully digest either grasses (high fiber and starch, low digestibility cell walls) or browse (low fiber and starch, high digestibility, cell soluble). Feeding strategy is not always related to size or family relatedness. Goats and white tail deer are intermediates whereas mule deer are browsers and sheep are grazers. Feeding strategy influences digestive morphology and physiology and will influence the type and amounts of the diet fed to captive small ruminants. Grazers have a larger rumen and abomasum, slower fermentation times and are primarily foregut fermenters whereas browsers have a smaller rumen and abomasum, faster fermentation times with a greater emphasis on hind gut fermentation. Depending on life stage, grazers may maintain on high quality grass hay as the primary fiber source whereas a browser will require alfalfa. Regardless of feeding strategy, sudden changes in diet are a problem for ruminants, especially increasing the amount or type of concentrate. Dietary changes should always be made gradually, usually over a 1 - 2 week period. Questions concerning diet fed and any changes made, is an important part of history taking for a sick small ruminant.
All ruminants are born without a developed fore stomach system, and can be treated as monogastrics up to approximately 3 weeks of age.
Evaluation of rumen function is part of the physical exam. Listen or feel in left paralumbar fossa for strong primary contractions, normally occurring every 2 or 3 minutes, with two or three secondary contractions between. Slow, weak, or absent rumen contractions suggest weakness, dietary problems, or severe metabolic disease. Gassy or bubbly borborygmi are not normal. Also ballot the rumen with your fingers or fist. The normal rumen has a solid, doughy consistency. Splashy contents are bad.
Temperature, Pulse and Respiration
The normal temperature, pulse and respiration for domestic small ruminants are: Temperature (102-104°F), pulse (goats 70-95 bpm / sheep 70-80 bpm), and respiration (15 - 30 breaths per minute).
Oral Exams and Teeth
All ruminants lack upper incisors. The lower canine has moved forward to become incisiform and thus there are 4 lower teeth on each side. In some deer species the upper canine remains and in muntjac these canines may protrude past the upper lip and are sharp. Extensive wear of the incisor teeth is common in older animals. Depending upon the species, the incisors are, slightly loose as compared to other animals. Ruminants have narrow mouths that do not open very far, thus hampering visual exam of the molar teeth. Careful palpation through the cheeks or use of a mouth speculum (Harp speculum) will facilitate an exam. Abnormal wear, periodontal disease, sharp points, missing teeth or lesions on the tongue or buccal cavity may be seen.
Restraint and Handling
The approach to restraint and anesthesia for these species will be governed by the level of habituation to handling and human contact, the handling facilities (holding pens, raceways and squeeze chutes) available and the animal handling expertise of the caretakers. All ruminants are prey species and flight is likely their first response to a threat. Many species also come with weapons (antlers, horns, canine teeth and sharp hooves) and will vigorously fight back.
The more intensively reared or tame the animals, the easier they will be to work and decreasing the flight zone will make the capture or immobilization procedure less unpleasant and stressful for all involved.
A properly set up chute and squeeze system is the most efficient and safest method for handling exotic small ruminants. A standard squeeze chute (with or without a head catch) is utilized for larger animals such as elk. A drop floor chute (where after entering the chute, the floor is dropped leaving the animal suspended and thus decreasing struggling and minimizing stress) is appropriate for deer and antelope species. The width of the drop floor chute should be adjustable to accommodate a variety of species and allow access to the front and rear and the back and legs. Once in a chute the animal should be blind folded to reduce visual stimulation. A halter can be applied and the head tied. Local anesthetic, sedation or immobilization agents can then be administered.
Extremely tame animals may allow administration of anesthetic agents by hand syringe either IM or IV. Pole syringes or jab sticks can be utilized on animals that can be confined to small pens or stalls. Remote delivery systems involving darts delivered by blow pipe, dart pistol or rifle are available.
It is beyond the scope of this paper to list specific drugs and dosages for exotic small ruminants. However there are excellent references that provide such information for all species.1,2,4 Alpha-2 agonists (xylazine, medetomidine, detomidine) alone or combined with a cyclo-hexamine (ketamine or Telazol [tiletamine and zolazepam]) are the most common non-narcotic mixtures used for immobilizing small ruminants. Combining ketamine or Telazol with an alpha-2 agonist balances out the excitement, convulsions and muscle hypertonicity that may be seen when using only a dissociative. The addition of the alpha-2 agonist also decreases the dosage of ketamine or Telazol required, increases the reversibility of the combination and enhances sedation and analgesia. A reversing agent (atipamezole, tolazoline, yohimbine) should be on hand whenever an alpha 2 agonist is used. Reversing the sedative properties of an alpha 2 agonist also reverses the analgesic properties.
As a rule of thumb, the alpha-2 should not be reversed sooner than 30 and ideally 60 minutes after induction if the combination contained ketamine or Telazol. This usually allows enough of the cyclo-hexamine to be metabolized before the alpha-2 is reversed. The opioid agonist - antagonist butorphanol or the butyrophenone tranquilizer azaperone are often added into these cocktails to enhance sedation. Butorphanol can be antagonized with the opioid antagonist naltrexone, azaperone has no antagonist and is often used to smooth the recovery. A new combination that has been used effectively in cervid species is butorphanol, medetomidine and azaperone or BAM (ZooPharm, Inc. P.O. Box 2023, Fort Collins, CO 80522-2023, USA, (www.zoopharm.net). The advantage of BAM is that the butorphanol and medetomidine are fully reversible greatly shortening recovery times over cyclo-hexamine based combinations.
The butyrophenone tranquilizer haloperidol along with the long acting neuroleptics (LANs) such as zuclopenthixol and perphenazine are used to calm extremely flighty species before transport, group introductions or game auctions.
Prolonged procedures or those requiring the animal to be positioned in dorsal recumbency should utilize general anesthesia with intubation. Adult animals should be fasted for 12-24 hours and water withheld for 8-12 hours to lessen the likelihood of regurgitation. Neonatal animals should not be fasted do to the risk of hypoglycemia. Correct positioning of the anesthetized animal is essential. Ruminants salivate profusely and continue to do so under anesthesia. The "head up, nose down" rule should be followed to allow for drainage of saliva. Rolled towels or sand bags can be placed under the throat area to raise the head and allow the nose to point down. If the animal is maintained only on a mask then saliva should be periodically drained from the mask.
Ruminants produce gas during anesthesia yet are unable to eructate. Positioning the animal with the thorax higher than the abdomen will help decrease pressure from the viscera on the lungs. Animals should be monitored for bloat; if excessive pass a stomach tube. During recovery place animals in sternal recumbency as soon as possible to allow for eructation.
Intubation is slightly more challenging because ruminants cannot open their mouths very far and have a narrow jaw. The base of their tongue is thick and the laryngeal opening can be difficult to visualize. The head and neck should be extended and the upper and lower jaws opened using roll gauze or string. The use of a 12" or longer Miller type laryngoscope blade will facilitate visualization of the larynx. The use of a polypropylene dog urinary catheter as a stylet may aid in placement of the endotracheal tube. O2 flow rates of 5.0 ml/lb/min (11.0 ml/kg/min) should be used.
Pulse oximetry probes can be placed on the tongue, ear, vulva or teats. When anesthetized, sheep and goats are likely to develop hypoxemia caused by hypoventilation, venous admixture, and recumbency. Supplemental oxygen at a rate to keep the SpO2 above 95% should be provided if the animal is not intubated. Due to low tidal volume during spontaneous respirations, end tidal C02 may not be reliable. Small ruminants should be "sighed" every 5 minutes. Mean arterial pressure is 75-100 mm Hg (systolic 80-120 and diastolic 60-80 mmHg). The palpebral reflex is lost under light anesthesia and thus this reflex is not useful for monitoring depth of anesthesia. Monitoring of the heart rate, pulse pressure, respiratory rate, capillary refill time, and muscle relaxation will aid in the assessment of anesthetic depth. Preventing hypothermia, padding pressure points and administration of a balanced electrolyte IV fluid at a rate of 5 ml/lb/hr (11 ml/kg/hr), are all required for a successful anesthetic procedure in the small ruminant.
Analgesia and Pain Management
Many procedures can be completed with sedation and local anesthesia. If the dose of lidocaine is calculated correctly, then it does not appear to be any more toxic to small ruminants than to other animals. The recommended limit, 6 mg/kg of lidocaine is conservative. Six mg/kg comes to 12 ml of 2% lidocaine in a 40 kg (88lb) goat but only 0.9 ml in a 3kg (7lbs) kid. When working with smaller volumes in neonates or smaller species, lidocaine is diluted to 1% or ½% to allow for infusion of larger volumes. Dilution does not reduce efficacy, but may shorten the duration of anesthesia. Bupivacaine may also be used at 2 mg/kg or mixed 50:50 with lidocaine for procedures where prolonged analgesia may be beneficial. Lidocaine as manufactured has a very low pH, which makes it painful on injection. Adding one part 8.4% bicarbonate to 9 parts lidocaine just before injection will create a more neutral pH and a much less painful injection. The solution will become cloudy, but it seems to function just fine in the tissues.
Non-steroidal anti inflammatory drugs (NSAIDS) are reasonably well absorbed by oral administration in ruminants and appear to be useful in pain management. Flunixin meglumine (1-2 mg/kg, IV, IM, SQ, SID), carprofen (2-4 mg/kg, PO, IV, SID), ketoprofen (2.2 - 3 mg/kg, IV, SQ, SID) and meloxicam (0.05 - 0.5 mg/kg, IV, PO, SQ, SID) have all been used.
Vaccination programs should at a minimum include an annual combination, clostridial vaccine which includes tetanus. These vaccines may cause lumps or sterile abscesses at injection sites. Utilizing sterile technique and massaging the injection site may help. Other vaccines for caseous lymphadenitis, orf, abortion diseases, footrot, and respiratory disease are available, but should be used with caution and only when a need is established based on diagnosis and history.
Hoof trimming may be required and frequency is dependent on climate, activity level, and substrate.
Fecal exams should be quantitative; either centrifuged sugar flotation or modified McMaster technique. In order to effectively control parasites while maintaining efficacy of anthelmintic, the following rules apply. Parasites should be monitored (80% of the worms are in 20-30% of the animals) and aggressively treated during periods of peak transmission, i.e. warm humid seasons. Periods of dry, very hot, or very cold weather, decrease transmission of gastrointestinal parasites. It is advisable to treat animals just prior to the onset of adverse weather. If possible avoid use of injectable or pour on dewormers in small ruminants, concentrating instead on orally administered products. Injectables have been shown to increase the rate at which parasites develop resistance to a product. Due to increasing reports of parasite resistance to anthelmintics a holistic program should be incorporated into the management program (manure and pasture management, reduced stocking densities, environmental barriers and optimal nutrition) where internal parasites are a major cause of morbidity. An excellent source for information regarding parasites in small ruminants is The Southern Consortium for Small Ruminant Parasite Control (www.scsrpc.org).
Trichostrongyle species especially Haemonchus contortus are a significant parasites in adult small ruminants. Recently a lectin staining technique which differentiates Haemonchus from other trichostrongyles has been made commercially available at a number of diagnostic laboratories (oregonstate.edu/vetmed/diagnostic/tests/haemonchus-contortus-identification).
In the eastern and mid-western United States, the presence of Parelaphostrongylus tenuis the brain worm of white tailed deer has caused neurological disease in other free-ranging cervid species (moose) as well as captive cervids and other small ruminant species housed in areas with high numbers of white tail deer as well as an environment which supports the gastropod intermediate host. Ivermectin has been used as a monthly preventative treatment often resulting in resistance problems with other parasites present in the population.
Any oral medication must first pass through the rumen and few antibiotics, for example, make it through the rumen intact. Oral trimethoprim sulfa combinations, commonly used in horses, are not efficacious for ruminants.
Because almost all drugs used in small ruminants are used in an extra label manner; and because sheep, goats and deer are considered to be food animals; veterinarians who work with them should be familiar with and follow AMDUCA regulations concerning food animals. Establishment of reasonable withdrawal time should always be considered before a drug is given to a potential food animal (www.farad.org).
In addition to Procaine penicillin G / benzathine penicillin G combinations, and ox tetracycline in a depot form, there are a number of newer broad spectrum, long acting, antibiotics that have been used in domestic and exotic small ruminants. The value of these antibiotics is their prolonged dosing interval: Naylor® (Florence: 2-3 days) (Internet, Sheering-Plough Animal Health, Millsboro, Delaware19966, USA), Excede® (ceftiofur crystalline free acid: 5-7 days) and Draxxin® (tulathromycin: 10-12 days) (Pfizer Animal Health, Kalamazoo, Michigan, 49001, USA).
A number of infectious diseases have been transmitted between exotic small ruminants and domestic livestock , are of regulatory concern or have zoonotic potential.
Johne's disease (mycobacterium paratuberculosis); has been documented in many species. In small ruminants, diarrhea may only be apparent as a terminal event and few blood tests have been validated for species other than cattle. Fecal culture methods remain the accepted diagnostic test for most non-bovine species. Further information can be found on The Johne's Information Center website (www.johnes.org).
Tuberculosis (mycobacterium bovis); all ruminants are susceptible and transmission has occurred between domestic species and wildlife (free-ranging and captive). Specific skin testing protocols are required for testing cervid species.
Caseous Lymphadenitis (Corynebacterium pseudotuberculosis); most commonly causes external lymph node associated abscesses. However internal abscesses can occur primarily in the neck and thorax.
Epizootic hemorrhagic disease (serotypes EHV1 and EHV2) and Blue tongue Virus (BTV serotypes 2,10,11, 13 and 17). Clinical disease with these orbiviruses is indistinguishable between EHDV and BTV. The incidence is seasonal due to vector transmission via biting midges of the Culicoides sp. Morbidity and mortality rates are variable between species. More information on EHD and BTV can be found at the Southern Cooperative for Wildlife Disease Study website (www.SCWDS.org).
Malignant Catarrhal Fever (MCF); this subgroup of ruminant gamma herpesvirus include not only the alcelaphine herpes viruses but also sheep associated OvHV-2, Caprine associated CpHV-2 and an unidentified gamma herpesvirus that has caused MCF like disease in white-tail deer. MCF is commonly spread between small ruminant species. A number of outbreaks have occurred in zoos where infected sheep and goats within the petting zoo have caused high morbidity and mortality in other exotic ruminant species (cervids appear to be extremely sensitive).
Urolithiasis is seen in all male small ruminants and is extremely common in castrated goats. When suspected, this condition is considered a medical emergency. "Is the animal urinating a normal stream?" is the most important question to ask your client with a sick male small ruminant. Diagnosis, surgical and medical management and prevention are described in detail.3
1. Kreeger, T. J. and J. M. Arnemo. 2007. Handbook of Wildlife Chemical Immobilization 3rd ed. T. J. Kreeger, Wheatland, Wyoming.
2. Kreeger, T.J., J.M. Arnemo and J.P. Raath. 2002. Handbook of Wildlife Chemical Immobilization Intl ed. 2002. Wildlife Pharmaceuticals, Inc. Fort Collins, Colorado.
3. Van Metre, D. C, House, J. K, Smith, B. P, George, L. W, et al. 1996. Treatment of urolithiasis in ruminants: Surgical management and prevention. Comp Cont Ed Pract Vet 18: S275-289.
4. West, G., D. Heard and N. Caulkett. 2007. Zoo Animal & Wildlife: Immobilization and Anesthesia. Blackwell Publishing, Ames, Iowa.