Deidre K. Fontenot, DVM
Members of the charismatic megavertebrate taxa present an enormous challenge, literally, for veterinarians but with great opportunities in medical management of these species in captivity. Intervention in medical care of these animals will likely include restraint, sedation, and anesthesia. An approach to these megavertebrate procedures must be grounded with knowledge in basic human and animal safety measures, drug pharmacology and accessibility (see pharmacy resources below), remote drug delivery systems, and comparative anatomy and physiology of the species. With this foundation, you can then build your procedural plan through species-specific medical and husbandry resources, colleague consults, and knowledge of the unique anatomical and physiological challenges that your patient will bring to the event.
Extensive species-specific handling and restraint recommendations are well described in the literature for both captive and field conditions and are listed in Appendix 1 (hard copy distribution not included in proceedings). The objectives of this lecture are focused on broad concepts focusing on the captive medical management of elephant, rhino, hippo, and giraffe species for physical and chemical restraint procedures. We will overview:
Considerations and precautions to adequately prepare for a megavertebrate procedure including human and animal safety
Comparative anatomical considerations that influence anesthesia management in these megavertebrate species in captivity
Compare/contrast the benefits and risks of physical and chemical restraint for megavertebrate species in captivity
Overview of drug selection principles/ combinations for sedation and anesthesia of megavertebrate species
Share "best practices" with current information, experiences and background in these species
General Considerations and Species Precautions
These species are high profile by nature to the guests and thus to your husbandry partners. Economic factors may play a role in decisions to intervene, as well as the implications from a public relations perspective should a negative outcome occur. Because of these complicating factors, it is not unusual to experience significant "push back" from husbandry partners on these initiatives. The decision to perform a medical procedure on these species is a partnership and dialogue that must occur between husbandry managers and veterinary teams. A benefit risk assessment must be made based on indications for procedure (elective vs. non-elective), goals of the medical procedure, ability for adequate facility and personnel support and expertise for the procedure, and acceptance of potential negative outcomes. A mutual agreement must be made that the benefits of the intervention outweigh the risks and that the decision to anesthetize is indicated with a clear expectation of the potential outcomes. In addition, it is best to make the decision for anesthetic intervention in a suitable timeline that does not complicate an already high-risk anesthetic event with a delayed intervention on a compromised animal in poor physical status.
Human safety must be the utmost priority during anesthetic events due to the size and inherent risk of handling these megavertebrate species. The veterinarian bears a large burden as the driver of the anesthetic event; however, these procedures require a team effort with delegation of responsibilities for the event. It is highly recommended to establish a safety-point person to monitor human risks during the procedure. The veterinarian must be responsible for the handling of drugs, especially concentrated narcotics, and associated tools (e.g., dart, syringes), but other human safety issues can be managed by a husbandry or other area manager. It is every procedure participant's responsibility to be aware and speak up before, during and after an anesthetic procedure to eliminate or address human safety risks that can be avoided during a megavertebrate veterinary procedure.
Appropriate facilities and availability of skilled personnel will significantly impact your anesthetic management and prevent injury to your patient and those around them. It is recommended that specialty equipment be purchased well in advance of any scheduled event (see Appendix 2). Much of anesthetic management in exotic species revolves around logistical management ahead of time. Facility infrastructure that supports physical and chemical restraint procedures are unique to each of the species and are further discussed below.
These taxa present unique challenges due to their size and anatomy that directly influence the risk assessment for anesthetic procedures. Many complications can occur with physical and chemical restraint of these species. Although some comparative anatomy and physiology can be used for developing a medical plan, each species unique characteristics must be taken into consideration as well during the preparation and implementation. Across the megavertebrate species discussed, commonalities exist.
Gastrointestinal physiology for all of these species as a ruminant (giraffe) or hindgut fermenters (elephant, rhinoceros, hippo) requires a more prolonged fast as compared to some other species. If possible, a minimum of 12–24 hours is recommended depending on influence on individual behavior, climate, and procedural goals.
Current weights may be difficult to acquire and can have significant impact on your anesthesia outcomes. Morphological data by age exist in the literature and can be a guide if current weights are not feasible. Metabolic scaling for megavertebrate drug dosing is reported in the literature. Review the information and develop your personal conclusions on the value of its use.
Large blood volume due to body mass/weight must be taken into considerations in regard to fluid therapy logistics and emergent care for dehydration or hypovolemia and direct and immediate effects will likely not occur with fluid therapy alone.
Large body mass can present a significant challenge for respiratory health during an anesthetic event. Intubation should be considered for all megavertebrate patients to improve airway access for adequate ventilation to reduce risk of ventilation-perfusion mismatch due to a variety of factors. Large body mass can also influence risk factors for myopathy and neuropathy post-procedure. Procedural time efficiency and patient manipulation during induction of anesthesia to direct patient positioning can reduce these risks.
Skin thickness in all of these species is typically 3–5 cm in target areas and can influence quality of injection, darting success and vascular access. Darting and hand injection strategies for intramuscular regimens should target the thinnest skinned areas over muscle regions. Injections should be given with needle lengths of at least 4–5 cm. Darting needles should be as long as 8–10 cm and administered as perpendicular as possible to target areas. The most common location across the megavertebrate species is the shoulder and neck region. More specific recommendations are listed below by species. Subcutaneous space is often very difficult to access for large volume administration and likely not a viable scenario for fluid replacement.
Most of these species have bulky, elongated skulls with dental patterns devoid of incisors. Although most species have a narrowed intermandibular space and dental arcade, relatively small oral cavity compared to body size, intubation can be achieved with adequate planes of anesthesia to reduce jaw tone, appropriate specialty tools for manual access and palpation of laryngeal region for tube placement.
These species have comparative gastrointestinal anatomy to that of a horse as hindgut fermenters with large gastrointestinal tract (GIT) volumes. This, as well as compromised patient positioning, could influence their risk for bloat causing diaphragm and lung compression from a bulky GIT resulting in negative ventilatory challenges. Their primary respiratory mechanism is breathing through their trunk as primary nasal breathers so use caution with trunk position when not intubated. However, positive pressure ventilation cannot be accomplished through the trunk. Elephants must be intubated to address ventilation concerns during anesthesia. Intubation is best accomplished with a manual retraction of epiglottis and blind placement of a long stylet through the fleshy larynx and then utilization of the endotracheal tube (size 35–55 cm depending on animal's weight) Murphy's eye over stylet to guide tube into trachea. Pharyngeal pouches are present that aid in vocalization and food/water storage, so be cautious to examine and clean prior to intubation. Elephant do not have a pleural space typical of other species, but possess connective tissue that secures the lung to the thoracic wall and diaphragm to prevent lung collapse and improve ventilation. They also have partitions within the lung parenchyma comprised of elastic connective tissue as well to prevent alveolar compression through decreasing pressure differentials and supporting lung weight. This can also influence ideal patient position as elephant typically should be shifted to lateral recumbency within 10 minutes of recumbency if possible but certainly for longer procedures. Vascular access via the ear veins is excellent for sample collection and catheterization, but use caution with drug administration due to higher risk of vasospasm, phlebitis and thrombosis leading potentially to segmental necrosis.
These species have comparative gastrointestinal anatomy to that of a horse as hindgut fermenters with large gastrointestinal tract (GIT) volumes. This could influence their risk for ventilatory challenges with compromised patient positioning causing diaphragm and lung compression from this bulky GIT. Best location for dart or other intramuscular injections is behind the ear in a triangular region outlined by the jugular furrow, nuchal crest (avoid), and caudal neck/shoulder musculature. Vascular access via the ear is excellent for sample collection and catheterization. Arterial sampling can be accomplished on the medial aspect of the ear versus dorsal for venous sampling. Blood gases do not appear to be influenced by sample location, so either can be used on the ear. Deep plane of anesthesia required with complementary tools to open the mouth wide enough in some species for manual intubation. Similar techniques to the elephant can be used.
These species have comparative gastrointestinal anatomy to that of a horse as hindgut fermenters, but with comparatively larger stomach capacity with larger gastrointestinal tract (GIT) volumes. This could influence their risk for ventilatory challenges with compromised patient positioning causing diaphragm and lung compression from this bulky GIT. These species are notorious for anesthesia complications, most likely related to their aquatic adaptations for dive physiology often called the "dive reflex" making them prone to apnea, hypoventilation, hypoxemia, bradycardia, and blood gas trends consistent with a transition to anaerobic metabolism. Mortality rates are reported to be high if this occurs, but can be potentially addressed through intubation, manual ventilation with high-flow ventilator and/or drug reversals. Their large size, high head to body size ratio, large volume to surface area ratio, and tendency towards obesity in captivity increase the risk of complications. Hyperthermia and airway obstruction from head/body malposition are the most common. Normal dermal secretions, often called "blood sweat," increase the rate of evaporative water loss when out of water and can increase risk for hypothermia and dehydration with longer procedures. Skin thickness, as mentioned previously, requires 6–10 cm needle size and vascular access can be difficult. Ears often do not easily permit vascular access. The triangular-shaped tail can be accessed for ventral tail vein sampling but is not good for an indirect blood pressure measurement. Nasal passages close voluntarily possibly as part of dive physiology makes nasal insufflation difficult but doable. Their large, fleshy oral cavity with a redundant soft palate makes laryngeal access challenging at best. Epiglottis often has to be pulled forward and tracheal size is surprisingly smaller to body size (25–30 mm ET tube).
These species have comparative gastrointestinal anatomy to that of a cow, a true ruminant foregut fermenter with comparatively large stomach capacity, thus larger gastrointestinal tract (GIT) volumes. This could influence their risk for aspiration event during induction when significant force is inflicted on the rumen. Similar to other species as well, risk for bloat post-anesthesia exists, as well as ventilatory challenges with compromised patient positioning, causing diaphragm and lung compression from this bulky GIT. Their long neck and legs are their biggest anatomical concern during anesthesia, increasing risk for entrapment, airway obstruction, cervical and leg trauma, myopathy, and neuropathy during and post-anesthesia. Long oral cavity with narrow intermandibular space and a long meaty tongue prone to trauma/airway obstruction can make intubation challenging. Recommended tools include long laryngoscope blades with adequate lighting and stylet. Intubation techniques previously mentioned can be used for direct visualization or manual blind intubation, depending on the quality and size of your tools. Significant respiratory dead space exists due to trachea length and size. This can increase their risk for hypoventilation and atelectasis. Increasing tidal volume requirements and placing the largest ET tube possible can ameliorate these issues. Manual ventilation with a high-flow ventilator is useful. Thoracic conformation limits their chest expansion. This along with increased abdominal pressure from recumbency during anesthesia can further complicate adequate ventilation. Cardiovascular adaptations include increased cardiac size and ventricular hypertrophy and hypertension. This reinforces the need for close monitoring anesthetic event parameters that reflect cardiovascular health during the procedure to reduce risk of hypoxic injury. Finally, their dense and thick skin can be advantageous to prevent dependent fluid accumulation as a procedural complication, but can make vascular access a challenge. Aural and jugular veins can be accessed for sampling and drug administration.
Physical restraint of the megavertebrate patient can be a viable alternative or complement to your chemical options. Patient size requires additional facility components and infrastructure to permit these procedures. Foresight, typically by veterinary involvement in barn and exhibit design, is the best approach to having chute systems integrated into facilities versus retrofitting structures. Most captive megavertebrate animals can achieve basic medical behaviors to permit a limited physical exam, blood collection, and injections in a chute restraint device. Animal training plan models can be found at http://animaltraining.org. Benefits of physical restraint include elimination of many of the risks associated with immobilization, acquisition of minimum database on a patient as a preanesthetic or medical evaluation before an anesthetic event, and engagement of the husbandry team on the medical care of the animals. Experience in megavertebrate training and restraint is recommended by a husbandry manager or veterinary team member to ensure a successful program and veterinary procedure without anesthesia. The restraint devices can also support a sedative procedure to facilitate ease of sample collection and more invasive procedures such as radiographs, ultrasound, biopsy and centesis/aspirate procedures (joint, abdomen, mass). Species-specific recommendations include:
Physical restraint has many different handling approaches in these species. Free contact, modified protected contact and protected contact all allow for different levels and patient accessibility and human safety risk. These options may not require significant infrastructure needs as effective training programs can often permit basic medical evaluations, including more invasive procedures such as reproductive ultrasounds. The scope of this lecture does not permit a complete review of handling approaches and can be reviewed extensively at http://elephantcare.org/. Chute systems, typically called elephant restraint devices (ERD), can be static or hydraulic to permit anterior and posterior door closure and lift and rotating capabilities for increased patient accessibility and more controlled anesthetic inductions and patient positioning. The purpose of an ERD is to restrict the elephant's movements in an environment safe for the handlers and elephants, while allowing handlers access for routine husbandry and medical care. The AZA Elephant Standards, updated 5 May 2003, required all facilities housing elephants to have an elephant restraint device (ERD) by 2006. The resource section below outlines contact information for select companies that build ERDs. Ancillary equipment, such as belly straps integrated into the ERD, and overhead hoists around a chute structure are useful tools as well.
Training programs for these species may also permit basic medical evaluations without chute system infrastructure including more invasive procedures such as reproductive ultrasounds. Hydraulic and static chute systems with underlying scales are great tools to assist with training programs, standing sedation and anesthesia induction in more controlled settings. Equipment such as belly straps and overhead hoists around a chute structure are useful tools as well. Chute designs and development processes can be reviewed at www.rhinoresourcecenter.com/pdf_files/117/1175856518.pdf and http://www.rhinoresourcecenter.com/pdf_files/117/1175858638.pdf
Training programs for these species may also permit basic medical evaluations without chute system infrastructure. Hydraulic and static chute systems with underlying scales are great tools to assist with training programs and anesthesia induction in a more controlled setting. Chute designs for hippo are difficult to find but are developed very similar to rhino chutes. Examples can be found at http://www.faunaresearch.com/hippo_rhino.htm. Equipment such as belly straps and overhead hoists around a chute structure are useful tools as well.
Training programs for these species will permit basic medical evaluations (visual exam +/- small blood volumes from ear or facial vein) in a basic stall setting; but, typically are most successful within chute system infrastructure. This allows for better control and patient accessibility during medical husbandry procedures (large volume blood collection from jugular vein, more extensive physical exam, hoof work) without sedation or anesthesia including more invasive procedures such as reproductive ultrasounds. Numerous chute structure designs exist in zoos but all have similar static structure with manual or hydraulic front and rear doors, usually manual squeeze mechanism side door, a "break away" side door for removal of anesthetized giraffe or in an emergent "down giraffe" situation, and side panel configurations to facilitate multiple access zones for sampling, injections, and examination. Underlying scales are great tools to assist with training programs and accurate anesthetic dosing. Equipment such as belly straps, break away halters and overhead hoists around a chute structure are useful tools as well.
Equipment and Procedure Considerations
A list of recommended supplies is included (Appendix 2). Each species present unique needs but many tools can be utilized across species and do not require specialty production, just adequate preparation and timely acquisition.
Along with knowing your species anatomy and physiology considerations mentioned above, you must know your individual and restraint conditions to adequately plan. Assessment of individual patient temperament is extremely important during the preanesthetic evaluation. In most cases, it provides the basis for subsequent handling, drug selection, personnel and equipment requirements. Time permitting, any preanesthetic desensitization that can occur with the patient can increase patient comfort level with the procedure events, including comfort with medical behaviors, head and neck manipulation, contact with ropes, straps, and halters, and normal chute exposure routine. Scheduling of the anesthetic event should avoid ambient temperature complications (hot or cold) and minimize change in individual animal's routine to reduce external factors on quality of anesthesia.
It is important to know the limitations of your facilities, team, and equipment and gather adequate information, consultants, and specialist to maximize your success, as well as diagnostic and therapeutic outcomes. Communications about the specifics of the event should make expectations and role clarifications clear for the team involvement and should also include non-procedural partners, including public relations, executive level management, facilities/maintenance equipment operators, and pathology. If you are utilizing concentrated opiates, consider notification of your local first responder ahead of time so they are adequately prepared for a call.
Preprocedural walk-through with all team members involved should include facility evaluation with evaluation of any potential facility hazards that may impact safety or logistics. Examples include filling large drinkers with sandbags or stuffed repurposed feedbags, application of soft, non-slip substrate for induction and recovery, and padding for poles or bollards that may be a hazard during induction or recovery during a stall or paddock immobilizations. Avoiding water hazards especially in hippo species is critical to planning. It is best to have all necessary equipment secured and present during preprocedural walk-through unless heavy machinery such as hoist, forklift, cranes and vehicles must be rented. Immobilization space must take into consideration the staffing and equipment capacity that is like to consume the space, as well as potential need for overhead access for support beams, eye bolts or crane and/or forklift access.
Chemical Restraint and Anesthesia
Preparation of a solid anesthetic plan can increase success and prevent most complications. This should include clinical and non-clinical considerations and factors that will contribute to the successes or challenges of the procedure. Your anesthesia equipment should be minimally different than any other anesthetic event with the exception of portability and size. Monitors should be easy to bring into the field and provide maximum physiologic information.
The use of premedications may make anesthetic induction smoother, both from a handling point of view as well as decreasing the amount of drug needed to induce anesthesia.
Sedative and Induction Regimens
The scope of this lecture does not allow for a complete review of the drug selection. With each drug combination, each species poses its own special risks and considerations as discussed above. The best option is to invest in a few good reference books and start with published doses and modify them to suit your own needs (see Appendix 1 for bibliographic references for anesthetic regimens). Table 1 provides limited recommendations based on author's institutional experiences. This table provides standing sedative and immobilizing options for induction and maintenance via manual continuous infusion [MCI] or continuous rate infusion [CRI] by species.
Analgesics should be routinely used whenever pain is present or anticipated. Nonsteroidal antiinflammatories such as flunixin meglumine, phenylbutazone, and carprofen at horse and cattle doses are often utilized.
Table 1. Megavertebrate chemical restraint recommendations based on author's institutional experiences
This table provides standing sedative and immobilizing options for induction and maintenance via manual continuous infusion [MCI] or continuous rate infusion [CRI] by species.
African elephant - standing sedation
Detomidine 20 µg/kg(D) + / butorphanol (B) 20 µg/kg IM (mild to moderate sedation)
African elephant - immobilization, anesthesia, field and captive dosing
Detomidine 20 µg/kg + butorphanol 20 µg/kg IM premed, then etorphine 1 µg/kg + azaperone 10–15 µg/kg IM or etorphine 2 µg/kg, azaperone 20 µg/kg IM for immobilization
0.5– 0.6 µg/kg/h Etorphine CRI
0.08–0.1 µg/kg/10–12 min Etorphine MCI IV in absence of mechanical pump
White rhino - standing sedation
Azaperone 30–60 µg/kg IM (mild to moderate sedation)
or Detomidine 20 µg/kg + butorphanol 20 µg/kg IM (moderate to heavy sedation)
or Detomidine 5–6 µg/kg + azaperone 60 µg/kg IM (moderate to heavy sedation)
White rhino - immobilization anesthesia (fair muscle relaxation with M99, consider additional azaperone or alpha-2 IV/IM)
Etorphine 1–2 µg/kg+ azaperone 25 µg/kg IM
or Etorphine 1µg/kg + detomidine 2–3 µg/kg IM
or Butorphanol 60 µg/kg + azaperone 40 µg/kg + medetomidine 20 µg/kg +/- ketamine 0.5–1mg/kg IM
Etorphine 0.5 µg/kg/h MCI IV every 15–20 min in absence of mechanical pump
or Guaifenesin (G)/ketamine (K) 1–2 ml/kg/h IV [Mix G 5% (50 mg/ml) + K (0.5–1 mg/ml), titrate dosing for depth/physiologic effects, may require loading dose of ketamine 0.1–0.2 mg/kg IV prior to MCI/CRI start]
Black rhino - standing sedation
Etorphine 0.2–0.3 µg/kg + detomidine 5 µg/kg IM (moderate to heavy sedation)
Black rhino - immobilization anesthesia
Etorphine 2.7 µg/kg + azaperone 60 µg/kg IM
Butorphanol 60 µg/kg, azaperone 40 µg/kg, medetomidine 25 µg/kg IM
Guaifenesin (G)/ketamine (K) 0.25–1 ml/kg/h [Mix G 5% (50 mg/ml) + K (0.5–1 mg/ml), titrate dosing for depth/physiologic effects, may require loading dose of ketamine 0.1–0.2 mg/kg IV prior to MCI/CRI start]
Hippo - standing sedation
Detomidine 30–60 µg/kg + butorphanol 80–150 µg/kg (moderate to heavy sedation)
Hippo - immobilization anesthesia
Etorphine 1.5–2 µg/kg + xylazine 50–60 µg/kg or medetomidine 40–50 µg/kg IM
or Butorphanol 100 µg/kg + azaperone 70 µg/kg + medetomidine 40 µg/kg IM
Typically utilize ketamine MCI at 0.5–1 mg/kg as needed
or Isoflurane via nasal insufflations or ET tube (allow 15 min of oxygen prior to reversal)
Giraffe - standing sedation
Azaperone 0.2–0.3 mg/kg IM (mild to moderate sedation)
Detomidine 30–40 µg/kg + butorphanol 30–40 µg/kg IM (moderate to heavy sedation)
Giraffe - immobilization anesthesia
Detomidine 35 µg/kg + butorphanol 35 µg/kg IM premed, then thiafentanil 10 µg/kg + ketamine 0.75–1 mg/kg IV for immobilization
Guaifenesin (G)/ketamine (K) 0.5–1 ml/kg/h - [Mix G 5% (50 mg/ml) + K (0.5–1 mg/ml), titrate dosing for depth/physiologic effects]
or Thiafentanil CRI IV 5 µg/kg/h, may require loading dose of thiafentanil 3–4 µg/kg IV prior to CRI/MCI
1Reversal agents across species are dosed as per below:
Carfentanil/etorphine/thiafentanil: naltrexone at 100x concentrated narcotic dose and 2x butorphanol dose IM or IV
Medetomidine/detomidine/xylazine: yohimbine at 0.1mg/kg IM or atipamezole at 5x alpha-2 agonist dose (author prefers IM)
Anesthetic monitoring in wildlife anesthesia is often underutilized. Megavertebrate anesthesia and all the ventilatory challenges that can occur reinforce the need for extensive monitoring in these species.
The key is to not allow devices to remove your attention from the patient. Delegate instrumentation so that you can use your eyes, ears and stethoscope to monitor what is actually going on with your patient. Monitoring depth of anesthesia can be accomplished through assessment of palpebral, corneal, and anal reflexes. Changes in eye position, muscle relaxation, jaw tone and tongue and ear movements are the first signs and animal is too light or too deep. Respiration, heart rate, circulation, response to stimuli and body temperature should be evaluated every 5–10 minutes and recorded. This will allow you to act accordingly and modify your plan. Resting heart rate is correlated with relative size and thus megavertebrate species tend to have ranges of 25–40 bpm. See species considerations below
Depth of anesthesia and sedation can be gauged by trunk and penile tone, palpebral reflex, ear movement, stance, and recumbency. Opioids have been reported in the literature to cause hypertension and subsequent "pink foam syndrome" from the trunk and ET tube likely due to increased pulmonary hydrostatic pressure. Azaperone has been used for its vasodilatory effects with the potent opiates. Ear arteries are excellent for pulse rate.
Depth of anesthesia and sedation can be gauged by head droop or pressing, stance, muscle relaxation, and recumbency. White rhino are notoriously sensitive to concentrated opioids with respiratory depression, muscle rigidity, and often incomplete anesthesia. Facial artery along the medial aspect of the mandible can be a viable site for pulse rate monitoring.
Depth of anesthesia and sedation can be gauged by head droop or pressing, stance, muscle relaxation, recumbency. Hippos can be difficult to monitor with auscultation without amplified stethoscope and peripheral pulses are difficult to find.
Depth of anesthesia and sedation can be gauged by head droop, stance, muscle relaxation, recumbency. Auscultation and facial artery can be utilized for heart/pulse rate monitoring.
Arterial Blood Gas Analysis
This is the gold standard for monitoring oxygen, carbon dioxide and pH. There are multiple options available for this type of equipment; however, to be effective it must be portable for field operations. The author uses an ISTAT (Abbott Point of Care, Inc., Princeton, NJ) system, which is a small handheld unit that has a selection of cartridges that plug into the machine and can test for different variables including blood chemistries. This system works very well but has limited temperature parameters in which to work (60°F–85°F). Venous and arterial access is previously discussed. A recent study in rhinoceros showed no significant difference in auricular arterial and venous sampling. Average blood gas and pH for ungulates: pH = 7.40 (range 7.35–7.45); PaO2 = 95 mm Hg (range 80–110 mm Hg), hypoxemia PaO2 = 70 mm Hg; PaCO2 = 40 mm Hg (range 35–45 mm Hg), hypercarbia PaCO2 > 50 mmHg.
This is the noninvasive measurement of oxygen bound to hemoglobin. It is measured via a light-emitting probe and provides real time, continuous and immediate information regarding blood oxygen levels. Developed for human use, this technology must be adapted for megavertebrate patients. The probe may be used on an ear, lip, scrotum, labia and even an eyelid. In species with dark skin, this technology may not work as the melanocytes interfere with the probe's light scores. In some species like hippo and rhino, if you scrape the ear with a scalpel blade and then apply the pulse oximetry probe, it will often pick up a reading. Care must also be taken to realize that the oxyhemoglobin dissociation curve drops off fast and a reading of 90 and 80% SaO2 may have magnitude difference in oxygen levels. Normal levels should be above 95% saturation. Pulse oximetry levels will be elevated with the addition of oxygen, but this must be balanced with CO2 levels to get a true picture.
Capnography (End-tidal CO2)
Capnography measures the amount CO2 being removed during exhalation. Most units show a capnogram, which characterizes the length and depth of the inspiratory phase. This is helpful in characterizing how the animal is breathing. Some capnography units are handheld and are available with a built-in pulse oximeter. These units are very helpful in that they are often more predictive of respiratory problems than pulse oximetry alone. SaO2 levels will be increased with oxygen supplementation, but CO2 levels can also remain high leading to a respiratory acidosis despite good oxygen supply.
Blood lactate is a measure of the transition from aerobic to anaerobic metabolic activity. Overexertion and inadequate oxygen or perfusion of blood will lead to an elevated lactate level. Lactate levels < 2.5 mmol/L are considered normal. Lactic acidosis or hyperlactatemia causes a decrease in blood pH, which is a precursor to capture myopathy. With the advent of small portable and inexpensive lactate analyzers for the human athlete field, this measurement can be easily taken with a single drop of blood. A good rule of thumb is to take a measurement at the start, middle and end of the procedure to see if a trend is developing. High lactate at the start of the procedure may indicate overexertion and should slowly decrease over the length of the procedure. If lactate is critically high > 15 mmol/L, supportive measures such as IV fluid with bicarbonate should be undertaken.
Blood pressure, which is commonly monitored during anesthesia, tends to decrease with increasing anesthetic dose in most species. Unless an arterial line can be placed, indirect blood pressure measurement is most often accomplished via the tail. Use caution with interpretation of absolute values and primarily look to trends for anesthesia monitoring. Be sure to run measurements with the tail as level with the heart as possible.
Anesthetic Complications, Emergency and Interventional Considerations
To start, improper dosing is a common cause of induction or anesthetic complications. These issues can be due to poor weight estimations, incomplete injection, new regimen in new species, and uncontrollable environmental or individual responses to drugs. Accurate weights, proper needle choice, dart placement and projectile pressure can alleviate some of these issues. Walk through and talk through "worst case scenario" situations and what scenario responses would be balanced with goals of procedure and prognosis of outcomes.
The most common anesthetic complication in megavertebrate species is hypoxemia and hypercapnia. Breath holding, such as the dive response in the hippo, will likely require intervention from diagnostic and therapeutic standpoints. Blood gas analysis will help determine the level of concern and intervention from nasal insufflation or intubation for passive oxygen administration or demand positive pressure ventilation to drug reversal and procedural abortion if non-responsive to supplementary measures. Respiratory stimulants, such as doxapram at 1–3 mg/kg IV, can be used, but only after oxygen supplementation has been addressed.
Other issues such as bloat, aspiration, hyperthermia, hypothermia, myopathy, and neuropathy should be addressed as you would with your smaller patients with prevention as the primary key. Airway protection with intubation is your greatest tool for many of these complications. Controlled inductions and proactive strategies for patient positioning, especially in giraffe, can be key in prevention of regurgitation with increased abdominal pressure during an induction fall and myopathy and neuropathy from poor patient position.
The author's practice has developed a Microsoft excel-based pharmaceutical reference sheet with calculation macros based on body weight and is produced for each anesthetic procedure for emergent issues. Quick dosage/dose references for triage, resuscitative, or commonly used therapeutics keeps our support team "ahead of the game" when drugs are requested during a procedure.
Reversal, Recovery and Post-procedure Considerations
The patient response can often be unpredictable during the reversal and recovery phase, but environmental parameters surrounding that event can be controlled and carefully communicated prior to reversal. Enforce the need for a quiet and calm environment during reversal, with human safety a primary priority, then followed by animal safety. Patient position during induction phase likely sets the stage for reversal position, except maybe in juvenile animals that can be manipulated, and must be taken into consideration when planning positioning for the procedure. Most of the megavertebrate patients utilize their head for standing as a lever system and should be taken into consideration from a human safety perspective. The breakaway halter in the giraffe works well to maintain some head control during recovery and can be removed as animal stands.
Recovery times are highly variable depending on a variety of factors of the individual, procedural time, and reversal methods (IV, IM). Appropriate substrate and good feet positioning are the key the effective attempts to stand within the first attempts. Appropriate use of electric cattle prod can encourage effective standing attempts. Monitor patient closely for 24–72 hours post-reversal for renarcotization with concentrated opiate combinations. Hippos should be restricted from water hazards for at least 4–5 hours post-reversal with sprayers to encourage appropriate thermoregulation.
Finally, whether immediately post-procedure or in a meeting later, a forum-type debrief of the procedure is highly recommended to discuss what worked and what could be improved at the next procedure. A safety check-in is recommended to be a part of that conversation.
Appendix 2. Supply List Example for Megavertebrate Anesthesia Event
Hay bales/feed bags for body support (rhino, hippo)
Padding, mattresses, water/ air beds for body support (elephant, giraffe)
Several towels (eye cover)
Ropes (lariat and soft of various sizes)
Equine break away halter (giraffe)
Various sized looped-ended straps - tow straps (to open mouth, lift head, move animal)
Come-a-long & chain set or block and tackle
Filled grain bags for eliminating fall/trip hazards in drinkers or other structures
Large plastic "rescue sled" (image in presentation)
12–14" inner tubes
Block wedges (elephant, rhino, giraffe - hard wood such as teak)
Rubber mallet (for mouth wedges)
10 x18" block with hole in center (hippo - hard wood such as teak, hole large enough to reach through for manual intubation)
Hydraulic/ pneumatic wedge to open mouth (elephant, rhino)
Bags of ice
Hammer, drill, pliers
Metal rakes (for I.V. poles)
Padded A-frame ladder or padded neck support board (approx. 8ft long) (giraffe)
3/4–1"plywood to fit a stall (this needs to surround the inside of a stall to keep rhino from going down with its head between the bollards)
Ear plugs (washcloths with tape strings separate or together)
Hay bags to place around poles/ bollards procedure/reversal area
Work lights, flashlights
5 gal buckets, brushes
Video camera, still camera
Cattle prods (consideration)
"Kill rifle" (consideration)
Remote drug delivery systems
Concentrated wildlife drugs (opiates, alpha-2 agonists, benzodiazepines, BAM cocktail)
Elephant medical management including AZA handling guidelines
Contact information for companies that build ERDs http://www.elephanttag.org
Frank Couch, Couch's Steel Construction, P.O. Box 249, Gulf Hammock, FL 32639, 352-486-4068
Basset, Inc. (formerly Bill Cummings & Sons), 2202 West Jones Avenue, Garden City, KS 67846, (620) 275-0300
Todd Ricketts, phone 915-479-5700, Fax 915-845-6916, email : email@example.com
Custom Venturi ventilator for megavertebrates and other ungulates (great field application) ~$800 http://mercurymed.com
Large animal ventilators (less wieldy for field procedures)
http://mallardmedical.net (custom ventilators for field application in large exotic species)
High volume fluid pump by Cole- Palmer
I-stat point of care blood gas analyzer
The author would like to thank Dr. Jeff Zuba for allowing me to share his comprehensive bibliographic compilation for this lecture. I feel we also must also acknowledge our zoo veterinary community for trail blazing the art and science of megavertebrate anesthesia so that we may provide excellence in animal care to the level of other species in our collections.
1. Citino S, Bush M. Giraffidae. In: West G, Heard D, Caulkett N. Zoo Animal and Wildlife Immobilization and Anesthesia. Blackwell Publishing;2007:595–612.
2. Fowler M, Mikota S. Chemical restraint and general anesthesia. In: Fowler M, Mikota S. Elephant Biology, Medicine, and Surgery. Ames, IA: Blackwell Publishing;2006:91–118.
3. Horne W, Loomis M. Elephants and hyrax. In: West G, Heard D, Caulkett N. Zoo Animal and Wildlife Immobilization and Anesthesia. Ames, IA: Blackwell Publishing;2007:507–521.
4. Mama K. Anesthesia in large exotic mammals. Small Animal Proceedings, American College of Veterinary Surgeons. 2009 Symposium.
5. Miller M. Hippopotami. In: West G, Heard D, Caulkett N. Zoo Animal and Wildlife Immobilization and Anesthesia. Ames, IA: Blackwell Publishing;2007:579–593.
6. Portas T. A review of drugs and techniques used for sedation and anaesthesia in captive rhino species. Aus Vet J. 2004;82(9):542–549.
7. Radcliffe R, Morkel P. Rhinoceros. In: West G, Heard D, Caulkett N. Zoo Animal and Wildlife Immobilization and Anesthesia. Ames, IA: Blackwell Publishing;2007:507–521
8. Zuba J, Citino S. Megavertebrate anesthesia. Am Assoc Zoo Vet Wetlab. 2011.