- Essential physiologic parameters, such as heart rate and rhythm, respiratory rate and depth, body temperature, and mucous membrane color, should be evaluated in all patients, including exotic animal species.
- Monitoring equipment is useful but cannot replace competent hands-on monitoring techniques.
- Maintenance of normal body temperature is an important priority during general anesthesia. Young, growing animals, patients in poor body condition, and small species with a high body surface-to-mass ratio are all at particular risk for hypothermia.
- During anesthesia and recovery, strive to maintain the body temperature of every reptile patient within its preferred optimum temperature zone.
- A change in respiration is sometimes the first sign of a problem that requires intervention. Therefore, the patient’s rate, pattern, and depth of respiration should be closely monitored throughout the perianesthetic period.
- Birds tolerate apnea much less than reptiles or mammals. If breathing stops for even 10-15 seconds in the avian patient, this is often an indication to reduce anesthetic depth and assist ventilation.
- All reptiles require intermittent positive pressure ventilation at a surgical plane of anesthesia.
- This article is part of a RACE-approved Anesthetic Monitoring teaching module. Visit the articles on monitoring the degree of central nervous system depression (anesthetic depth), blood pressure, capnometry, pulse oximetry, and electrocardiography for additional information in exotic animal patients.
Even the most steadfast and seasoned veterinary anesthetist can find themselves intimidated by exotic animal patients. Monitoring anesthesia can be a nerve-wracking endeavor in and of itself, and this can be further compounded by the added intricacies of these special species. Typical canine and feline veterinary anesthesia monitors are not designed to read the extremely high (or extremely low) heart rates and respiratory rates of some exotic animals. Despite these challenges, valuable information on patient vital signs can be gathered from monitoring tools as well as hands-on techniques (Fig 1).1,23 Essential physiologic parameters, such as heart rate and rhythm, respiratory rate and depth, body temperature, and mucous membrane color should all be evaluated (Table 1).
|Table 1. Normal physiologic values for birds and small mammals|
|Beats per minute||Breaths per minute||ºC (ºF)|
|MAMMALS||Chinchilla (Chinchilla laniger)||150-250||40-80||37-38|
(Mustela putorius furo)
|Guinea pig |
|280.0 +/- 6.7||38 |
|Hedgehog, African Pygmy|
|Hedgehog, European (Erinaceus europaeus)||200-280||25-50||35 +/- 2 |
|Prairie dog, black-tailed (Cynomys ludovicianus)||83-318||40-60||35.4-39.1 |
|Rabbit, European (Oryctolagus cuniculus)||220||55||38 |
|Skunk, striped (Mephitis mephitis)||140-190||25-50||38.9 |
|Sugar glider (Petaurus breviceps)||200-300||16-40||35.8-36.6 |
|Virginia opossum (Didelphis virginiana)||90-160||12-24||35-36 |
|BIRDS||Caged birds||39-42 |
|Amazon parrot (Amazona spp.)||340-600||15-45||41.8 |
|Grey parrot (Psittacus erithacus)||340-600||25-45||41 |
|Macaw||389 (small)||20-25||42 |
|Rock dove (Columba livia)||42.2 |
|Canada goose (Branta canadensis)||133 +/- 8.4||18.7 +/- 1.3||40.2 +/- 0.58|
|Canvasback (Aythya valisineria)||158 +/- 48||21+/- 3||39.0 +/- 0.8|
|Pekin duck (Anas platyrhynchos domesticus)||281.3 |
|Pekin duck||213 +/- 8||41.3 +/- 0.2|
|Pekin duck||190 +/- 17||19 +/- 4||22|
|Pekin duck||7 (6-13)||22|
|Pekin duck||218 +/- 32||17 +/- 4||22|
|Pekin duck||15.8 +/- 1.5||22|
|Buzzards (Buteo buteo)||325.2 +/- 52.9||22|
|HR: Heart rate RR: Respiratory rate|
An important priority for every anesthetist is to maintain normothermia.31 A fall in body temperature, or hypothermia, is extremely common during anesthesia, but is of particular concern in young, growing animals, patients in poor body condition, and in small patients with a high body surface-to-mass ratio (Table 2).17,30
|Table 2. Causes of heat loss during the pre- and perianesthetic period 10|
|Evaporative||Clipping or plucking
Introduction of cold anesthetic gases
|Conductive||Placement on cold surgical tables|
|Convective||Open body cavities
Cold irrigation solutions
|Radiant||Open body cavities
Cold irrigation solutions
Exposed patient surface area
Hypothermia can adversely affect the central nervous system, as well as the cardiopulmonary, gastrointestinal, and metabolic systems, resulting in a host of physiologic effects, including acidosis, bradypnea, decreased minute ventilation and tidal volume, bradycardia, cardiac instability, and arrhythmia.10,17,32 The minimum alveolar concentration (MAC) decreases as body temperature falls, and hypothermia also adversely affects the speed and quality of recovery.2 Hypothermia can also directly influence surgical site healing.33 Severe and prolonged hypothermia can even have potentially fatal consequences.3,6,24
Body temperature is one of the simplest vital signs to monitor during anesthesia.6 A rectal thermometer can be used; however, this measurement often underestimates core temperature and the readings from a standard thermometer may not go low enough for some reptile patients.6 If a thermometer is used, monitor body temperature every 5 minutes.17 Preferably core body temperature should be measured using an esophageal or rectal/cloacal temperature probe with continuous display (Fig 2).2,6,9,24,31 If an esophageal temperature probe is used, gently advance the probe into the thoracic esophagus to avoid the cooling effects of respiratory gases. The probe should not be advanced past the level of the thoracic inlet as it could enter the stomach or crop and receive false readings from gastric contents or feed material.6
Correction of hypothermia is challenging and every effort should be made to maintain normothermia or minimize hypothermia in the exotic animal patient (Table 3).10,22
|Table 3. Forms of thermal support during anesthesia (Skorupski et al 2017)|
EXOTIC COMPANION MAMMALS
Body temperature can quickly plummet below 35ºC (95ºF) when small mammals are not provided with supplemental heat.23,27 Flecknell (2015) has described reductions in body temperature up to 10ºC in anesthetized mice in as little as 15-20 minutes. Since tiny patients can sometimes heat up just as quickly as they cool down, it is also important to carefully monitor temperature during recovery to prevent hyperthermia and/or burns.17
Marsupials normally have a lower body temperature than eutherian mammals (Table 1). To effectively measure temperature, insert the thermometer into the dorsal part of the cloaca, which communicates with the rectum.15
Birds, particularly smaller species, have higher metabolic rates and higher body temperatures when compared to mammals of similar size. Normal body temperature ranges between 39-42ºC (102.2-107.6ºF) in many avian species.4,6,17,18,22,32 This high body temperature creates a steep temperature gradient between the bird’s core temperature and the temperature of the environment, which can to lead to rapid and dramatic heat loss when a bird is placed under general anesthesia.6,17
The superior insulation provided by plumage in waterfowl and other aquatic birds can cause significant hyperthermia, particularly if the bird struggles during induction or must be captured under field conditions.12,22 Additionally, some aquatic birds can shunt blood from their periphery to preserve core body temperature, making cloacal temperatures inaccurate.12
Reptiles are poikilotherms. General metabolism and cardiovascular physiology, which affects drug clearance, oxygen demand, acid base status, as well as induction time and recovery, are highly temperature dependent.2,8,17,23,31 Hypothermia leads to a progressive decrease in cardiac output and an increase in cardiac shunting.8 This ability to selectively shunt blood flow away from pulmonary circulation is likely the cause of prolonged recovery from inhalant anesthesia in the reptile patient.31 In anesthetized garter snakes (Thamnophis sirtalis parietalis), recovery time at 21ºC (69.8ºF) was twice as long as at 31ºC (87.8ºF).2,25
Maintain body temperature within the preferred optimum temperature zone (POTZ) for your species of interest during anesthesia and recovery.2,21 Although POTZ is species specific, 25-35ºC (77-95ºF) will be appropriate for most temperate and tropical squamates during induction, anesthesia, and recovery.2 Many freshwater turtles will tolerate temperatures between 20-25ºC (68-77ºF), while sea turtles and land tortoises are better adapted to temperatures between 25-30ºC (77-86ºF).20,31 Monitor reptiles carefully during recovery to prevent overheating and/or burns.17,31 Thermal burns can occur more easily in reptiles that are dehydrated with poor peripheral perfusion.2 Although low ambient temperatures will slow or halt reptile activity, hypothermia should never be considered part of the anesthesia or sedation protocol.
Changes in this vital sign can be a sensitive clue to the physiologic status of the patient (Table 4).9 There is no clinical consensus over when bradycardia or tachycardia should be treated, but conservative guidelines for intervention are approximately 20% below or 20% above normal.9 Both bradycardia and tachycardia should always be addressed when associated with evidence of poor cardiac output, reduced blood pressure, or inadequate tissue perfusion.
|Table 4. Potential causes of bradycardia and tachycardia 9|
Heart rate and rhythm can be monitored by an infant or pediatric stethoscope, an esophageal stethoscope, electrocardiogram (ECG), or a Doppler ultrasonic flow probe. The peripheral pulse rate can be palpated in birds and mammals, and an apex beat or pulse can sometimes be visualized in the reptile patient.
EXOTIC COMPANION MAMMALS
Assess the rate, rhythm, and quality of the peripheral pulse in larger species like the rabbit (Oryctolagus cuniculus). The femoral artery is most easily palpated.6 The peripheral pulse rate can also be palpated at the dorsal pedal artery, axillary artery, auricular artery, and the caudal (tail) artery. Heart rate can also be palpated directly over the thoracic cavity. Similarly, the transducer of the Doppler monitor can be placed directly over the heart or a superficial artery, like the carotid, carpal, or femoral artery, to generate real-time, continuous monitoring of heart rate.23 Use of the Doppler probe is particularly useful in tiny patients since changes can occur rapidly (Fig 3).20,31 An esophageal stethoscope can be used to monitor heart rate in larger animals, like the rabbit and ferret (Mustela putorius furo), but can induce regurgitation in guinea pigs (Cavia porcellus).14
Peripheral pulses can be found at the brachial (median or superficial ulnar), medial metatarsal, or carotid artery in the bird.10 The cutaneous ulnar or brachial artery is a popular site when the Doppler ultrasound probe is used to monitor heart rate (Fig 4, Fig 5).13 A Doppler flow probe can also be placed against the palatine artery in select species (Fig 6). The complex crop of psittacine birds and pigeons can make it difficult to pass an esophageal stethoscope into the thoracic esophagus in these species.12
The apex beat can usually be visualized along the ventral scales in snakes, approximately 25% along the snake’s length (Video 1). In many lizards, the pulse can be observed in the jugular groove.2 Doppler is considered the most reliable device for monitoring cardiac blood flow in reptiles.20,31 The Doppler probe can be placed directly over the heart (Video 2, Fig 8, Fig 9), or it can be taped over a superficial artery like the carotid artery (on the ventrolateral surface of the neck), the coccygeal artery (at the base of the tail), or the femoral artery. The reptile heart rate is highly influenced by temperature.5,31
Video 1. The wave associated with a beating heart is visible in this snake. Video credit: Katrina Lafferty, CVT, VTS.
Video 2. Doppler ultrasound probe placed over the heart of a snake. Heartbeat can be heard in the background. Note: Please enable audio to hear the heartbeat in the background. Video credit: Katrina Lafferty, CVT, VTS.
It is important to closely monitor the rate, pattern, and depth of respiration throughout the perianesthetic period.6 First, record the patient’s respiratory rate before the anesthetic event. Ideally the animal should be calm and relaxed, however many birds and small mammals will display a rapid respiratory rate in a clinical setting.6 During anesthesia, observe the frequency and range of sternal motion as well as movement of the reservoir bag. Substitute balloons for reservoir bags for easier scrutiny of bag movement in patients with small tidal volumes.6 Respiration can also be monitored by placing a sensor near the nostrils or mounted on the anesthetic breathing system close to the endotracheal tube or face mask connector. Some monitors utilize a pressor sensor, which can be triggered by chest wall movements.
A special concern in tiny patients is obstruction of the endotracheal tube. This can occur as a result of bending or kinking or through blockage of the tube with respiratory secretions. Tube blockage is often indicated by a sudden change in respiratory pattern with prominent whistling, wheezing, or harsh breath sounds produced. Expiration may appear difficult, slow, or nonexistent and the reservoir bag may no longer reflect movement. An end-tidal carbon dioxide monitor may show apnea or a change in waveform.18 When blockage is suspected, quickly evaluate the endotracheal tube for a kink and correct. If the situation is not immediately improved or blockage is suspected, promptly remove the tube and replace as needed.18
Although monitoring respiration has value, this vital sign does not adequately assess ventilation.23 An end-tidal carbon dioxide monitor will provide valuable information in most intubated patients.6
A change in respiration can be the first sign of a problem that requires intervention. When apnea develops, there can be a 1-2-minute delay before blood oxygen levels fall or carbon dioxide (CO2) levels rise in mammals.6
Birds are much less tolerant of apnea when compared to mammals and reptiles. If breathing stops for even 10-15 seconds this is often an indication to reduce anesthetic depth and assist ventilation.19
Most normal reptiles exhibit an intermittent breathing pattern characterized by exhalation and inspiration, followed by an apneic period of variable length.2 All reptiles require intermittent positive pressure ventilation at a surgical plane of anesthesia. Standard guidelines recommend four to six breaths per minute; however this may need to be adjusted for each individual patient.26 Based on blood gas analysis, five breaths per minute led to respiratory alkalosis in anesthetized rattlesnakes (Crotalus durissus) and one to two breaths per minute was deemed appropriate for this species.26
A drop in arterial oxygen saturation stimulates the respiratory drive in most reptiles. This is the opposite to mammals, which respond to an increase in arterial carbon dioxide levels. 29 To stimulate spontaneous ventilation in the reptile patient during recovery, low oxygen flow rates are maintained with the use of room air and a bag valve mask (i.e. Ambu bag).
Heart rate and rhythm, respiratory rate, and body temperature, should ideally be monitored during every anesthetic procedure. These vital signs can be gathered using hands-on techniques as well as monitoring tools. A change in respiration is sometimes the first sign of a problem that requires intervention. Birds tolerate apnea to a far less degree than reptiles or mammals. If breathing stops for 10-15 seconds or more in the avian patient, this is often an indication to reduce anesthetic depth and assist ventilation. All reptiles require intermittent positive pressure ventilation at a surgical plane of anesthesia. Changes in heart rate can be a sensitive clue to the physiologic status of the patient, and it is also crucial to carefully monitor body temperature and prevent hypothermia. General metabolism and cardiovascular physiology is highly dependent on environmental temperature in the reptile patient, which should be maintained within its preferred optimum temperature zone. In all species, hypothermia can result in a host of adverse physiologic effects. Severe and prolonged hypothermia can even have potentially fatal consequences.
1. Bailey JE, Pablo LS. Anesthetic monitoring and monitoring equipment: Application in small exotic pet practice. Semin Avian Exot Pet Med 7(1):53-60, 1998.
2. Bertelsen MF. Squamates (snakes and lizards). In: West G, Heard D, Caulkett N (eds). Zoo Animal and Wildlife Immobilization and Anesthesia, 2nd ed. Ames, IA: Wiley Blackwell; 2015:657-659.
3. Boedeker NC, Carpenter JW, Mason DE. Comparison of body temperatures of pigeons (Columba livia) anesthetized by three different anesthetic delivery systems. J Avian Med Surg 19(1):1-6, 2005.
4. Dawson WR, Whittow GC. Regulation of body temperature. In: Whittow GC (ed). Sturkie’s Avian Physiology, 5th ed. San Diego: Academic Press; 2000: 343-390.
5. DeVoe RS. Reptilian cardiovascular anatomy and physiology. Proc Annu Conf Am Board Vet Pract 2011.
6. Flecknell P. Laboratory Animal Anaesthesia, 4th ed. Boston: Elsevier; 2015.
7. Flecknell PA, Thomas AA. Comparative anesthesia and analgesia of laboratory animals. In: Grimm KA, Lamont LA, Tranquilli WJ et al (eds). Veterinary Anesthesia and Analgesia: The Fifth Edition of Lumb and Jones. Ames, Iowa: Wiley Blackwell; 2015: 758.
8. Galli G, Taylor EW, Wang T. The cardiovascular responses of the freshwater turtle Trachemys scripta to warming and cooling. J Exp Bio 207 (Pt 9):1471-1478, 2004.
9. Haskins SC. Monitoring anesthetized patients. In: Grimm KA, Lamont LA, Tranquilli WJ et al (eds). Veterinary Anesthesia and Analgesia: The Fifth Edition of Lumb and Jones. Ames, Iowa: Wiley Blackwell; 2015: 86-113.
10. Hawkins MG, Zehnder AM, Pascoe PJ. Cagebirds. In: West G, Heard D, Caulkett N (eds). Zoo Animal and Wildlife Immobilization and Anesthesia, 2nd ed. Ames, IA: Wiley Blackwell; 2015:770-772.
11. Hawkins MG, Pascoe PJ. Anesthesia, analgesia, and sedation of small mammals. In: Quesenberry KE, Carpenter JW (eds.) Ferrets, Rabbits, and Rodents: Clinical Medicine and Surgery, 3rd ed. St. Louis: Elsevier Saunders; 2012: 429-451.
12. Heard D. Anesthesia. In: Speer BL (ed). Current Therapy in Avian Medicine and Surgery. St. Louis: Elsevier; 2016: 611-615.
13. Heard D. Birds: Miscellaneous. In: West G, Heard D, Caulkett N (eds). Zoo Animal and Wildlife Immobilization and Anesthesia, 2nd ed. Ames, IA: Wiley Blackwell; 2015: 932.
14. Heard D. Rodents. In: West G, Heard D, Caulkett N (eds). Zoo Animal and Wildlife Immobilization and Anesthesia, 2nd ed. Ames, IA: Wiley Blackwell; 2015:1576-1579.
15. Holz P. Marsupials. In: West G, Heard D, Caulkett N (eds). Zoo Animal and Wildlife Immobilization and Anesthesia, 2nd ed. Ames, IA: Wiley Blackwell; 2015: 942-943.
16. Kik MJL, Mitchell MA. Reptile cardiology: A review of anatomy and physiology, diagnostic approaches, and clinical disease. Semin Avian Exot Pet Med 14(1):52-60, 2005.
17. Ko JC, Krimins RA. Thermoregulation. In: West G, Heard D, Caulkett N (eds). Zoo Animal and Wildlife Immobilization and Anesthesia, 2nd ed. Ames, IA: Wiley Blackwell; 2015:188-197.
18. Lierz M, Korbel R. Anesthesia and analgesia in birds. J Exotic Pet Med 21(1):47-48, 50, 54, 57-58, 2012.
19. Ludders JW. Comparative anesthesia and analgesia of birds. In: Grimm KA, Lamont LA, Tranquilli WJ et al (eds). Veterinary Anesthesia and Analgesia: The Fifth Edition of Lumb and Jones. Ames, Iowa: Wiley Blackwell; 2015: 808, 812.
20. McArthur S, Meyer J, Innis C. Anatomy and physiology. In: McArthur S, Wilkinson R, Meyer J (eds). Medicine and Surgery of the Tortoises and Turtle, 1st ed. Oxford: Blackwell Publishing; 2004: 35-72.
21. Mosley CI, Mosely CA. Comparative anesthesia and analgesia of reptiles, amphibians, and fishes. In: Grimm KA, Lamont LA, Tranquilli WJ et al (eds). Veterinary Anesthesia and Analgesia: The Fifth Edition of Lumb and Jones. Ames, Iowa: Wiley Blackwell; 2015: 784-787.
22. Mulcahy DM. Free-living waterfowl and shorebirds. In: West G, Heard D, Caulkett N (eds). Zoo Animal and Wildlife Immobilization and Anesthesia, 2nd ed. Ames, IA: Wiley Blackwell; 2015:880-886.
23. Ozeki L, Caulkett N. Monitoring. In: West G, Heard D, Caulkett N (eds). Zoo Animal and Wildlife Immobilization and Anesthesia, 2nd ed. Ames, IA: Wiley Blackwell; 2015: 149-165.
24. Phalen D, Mitchell ME, Cavazos-Martinez MI. Evaluation of three heat sources for their ability to maintain core body temperature in the anesthetized avian patient. J Avian Med Surg 10(3):174-178, 1996.
25. Preston DL, Mosley CAE, Mason RT. Sources of variability in recovery time from methohexitol sodium anesthesia in snakes. Copeia 105(2):496-501, 2010.
26. Schumacher J, Mans C. Anesthesia. In: Mader DR (ed). Current Therapy in Reptile Medicine and Surgery, 3rd ed. St. Louis: Elsevier; 2014: 303, 310-313.
27. Sikoski P, Young R, Lockard M. Comparison of heating devices for maintaining body temperature in anesthetized laboratory rabbits (Oryctolagus cuniculus). J Am Assoc Lab Anim Sci 46(3):61-63, 2007.
28. Skorupski, Anna M ; Zhang, Jingyi, et al. Quantification of induced hypothermia from aseptic scrub applications during rodent surgery preparation. J Am Assoc Lab Anim Sci 56(5): 562-569, 2017.
29. Taylor EW, Leite CA, McKenzie DJ, Wang T. Control of respiration in fish, amphibians and reptiles. Braz J Med Biol Res 43(5):409-424, 2010.
30. Taylor DK. Study of two devices used to maintain normothermia in rats and mice during general anesthesia. J AM Assoc Lab Anim Sci 46(5);37-41, 2007.
31. Vigani A. Chelonians (Tortoises, turtles, and terrapins). In: West G, Heard D, Caulkett N (eds). Zoo Animal and Wildlife Immobilization and Anesthesia, 2nd ed. Ames, IA: Wiley Blackwell; 2015: 681-702.
32. Zehnder AM, Hawkins MG, Pascoe PJ. Avian anatomy and physiology. In: West G, Heard D, Caulkett N (eds). Zoo Animal and Wildlife Immobilization and Anesthesia, 2nd ed. Ames, IA: Wiley Blackwell; 2015:723-739.
33. Ballard B, Cheek R (eds). Exotic Animal Medicine for the Veterinary Technician, 3rd ed. Ames, IA: Wiley Blackwell; 2017.
34. Cinar A, Bagci C, Belge F, Uzun M. The electrocardiogram of the Pekin duck. Avian Dis 40(4):919-923, 1996.
35. Clark-Price S. Inadvertent perianesthetic hypothermia in small animal patients. Vet Clin North Am Small Anim Pract 45(5):983-994, 2015.
36. Espino L, Saurez ML, Lopez-Beceiro A, Santamarina G. Electrocardiogram reference values for the buzzard in Spain. J Wildl Dis 37(4):680-685, 2001.
37. Heard D. Insectivores (Hedgehogs, moles, and tenrecs). In: West G, Heard D, Caulkett N (eds). Zoo Animal and Wildlife Immobilization and Anesthesia, 2nd Ames, IA: Wiley Blackwell; 2015: 956.
38. Yahav S. Regulation of body temperature: Strategies and Mechanisms. In: Whittow CG (ed). Sturkie’s Avian Physiology, 5th San Diego: Academic Press, Inc; 2000: 882.
Becker DE, Casabianca AB. Respiratory monitoring: physiological and technical considerations. Anesth Prog 56(1):14-22, 2009.
Churgin SM, Sladky KK, Smith LJ. Anesthetic induction and recovery parameters in bearded dragons (Pogona vitticeps): Comparison of isoflurane delivered in 100% oxygen versus 21% oxygen. J Zoo Wildl Med 46(3):534-539, 2015.
Hacker SO, White CE, Black IH. A comparison of target-controlled infusion versus volatile inhalant anesthesia for heart rate, respiratory rate, and recovery time in a rat model. Contemp Top Lab Anim Sci 44(5):7-12, 2005.
Heard D. Galliformes and columbiformes. In: West G, Heard D, Caulkett N (eds). Zoo Animal and Wildlife Immobilization and Anesthesia, 2nd ed. Ames, IA: Wiley Blackwell; 2015:871.
Longley L. Anaesthesia of Exotic Pets. London: Elsevier; 2008.
Krosniunas EH, Hicks JW. Cardiac output and shunt during voluntary activity at different temperatures in the turtle, Trachemys scripta. Physiol Biochem Zool 76(5):679-694, 2003.
Nevarez JG. Monitoring during avian and exotic pet anesthesia, Semin Avian Exot Pet Med 14(4):277-283, 2005.
Lafferty K, Pollock CG. Monitoring vital signs in exotic animal species. May 17, 2018. LafeberVet Web site. Available at https://lafeber.com/vet/monitoring-vital-signs-in-exotic-animal-species/