Perioperative Supportive Care and Monitoring

1094-9194 / 00 $15.00 -t .00

SOFT-TISSUE SU RGERY

PERIOPERATIVE SUPPORTIVE CARE AND MONITORING Darryl Heard, BSc, BVMS, PhD, Dipl. ACZM

The complexity and duration of exotic animal diagnostic procedures and surgeries increase every year, which creates a need for accurate and skilled perioperative supportive care and monitoring if morbidity and mortality are to be minimized. The diversity and sm all size of exotic species encountered in practice make this difficult, however. More research is necessary to validate the accuracy, sensitivity, and reliability of monitoring equipment and to evaluate the efficacy of supportive care techniques. Fortunately, many of the principles of supportive care and monitoring can be extrapolated across species. The trend in human medicine to develop small portable or "bedside" monitoring systems (e.g., blood gas and electrolyte analyzers) and technology to measure variables not previously measured (e.g., hemoglobin saturation ) offer opportunities to improve perioperative management dram atically. The main goal of supportive care is to minimize the adverse physiologic effects of anesthesia, surgery, and preexisting disease. Monitoring is performed concurrently to detect physiologic perturbations, ensure iippropriate anesthetic depth, and assess the efficacy of supportive care.

PERIOPERATIVE PLAN

Perioperative supportive care begins with a consistently and accurately applied plan. Written protocols facilitate this and are an essential part of interpersonnel communication. A checklist is stamped or stapled

From the Wildlife and Zoological Medicine Service, Department of Small Animal Clinical Sciences, College of Veterinary Medicine, University of Florida, Gainesville, Florida

VETERINARY CLINICS OF NORTH AMERICA: EXOTIC ANIMAL PRACTICE VOLUME 3 • NUMBER 3 • SEPTEMBER 2000

587

588

HEARD

into the medical record. Completion of the checklist with a recognizable signature can be an important legal document. Each anesthetic period has a permanent record that is an extension of the checklist. Minimally, it includes pertinent physical examination and history findings, recent body weight, drugs used, start and finish of anesthesia, endotracheal tube size, and problems encountered. Ideally, this record includes systematic recording of monitored physiologic variables. Human monitoring equipment routinely includes paper and electronic data recording systems. Regular evaluation of the anesthetic machine (e.g., vaporizer setting and oxygen flow rate) and electronic recording devices are included in the monitoring protocol. A single person or "anesthetist" is assigned to each patient to manage perioperative monitoring and support; this is important in small exotic patients, in which physiologic status can change rapidly. PREOPERATIVE PREPARATION

All animals are examined physically, and their medical histories are reviewed before induction. An accurate admission weight measurement and subsequent daily weight measurements are essential patient care. Assessment of cardiopulmonary function is emphasized, and baseline values for respiration, heart rate, and temperature are determined for reference during anesthesia. Minimal diagnostic testing includes collection of blood for packed cell volume, total protein, glucose, and urea nitrogen levels (mammals). Additional diagnostic tests (e.g., hematology, plasma biochemical panel, and radiographs) are performed as indicated. Aggressive and excited animals are examined immediately postinduction. Preinduction visual examination of these animals reveals open-mouthed breathing, tachypnea, dyspnea, nasal discharge, altered mentation, abnormal mucous membrane color, cachexia, or signs of dehydration, which can alter or cancel the anesthetic plan. Ideally, all patients are physiologically stable before anesthetic induction. If sufficient time is available, abnormalities (e.g., dehydration, anemia, hypoglycemia, electrolyte abnormalities, or acid-base disturbances) are corrected. Drugs (e.g., antibiotics) and blood products that have the potential to produce anaphylactoid reactions are also administered at this time. Vascular access (discussed later) is attained in unstable animals and those undergoing prolonged procedures or procedures that are likely to produce significant hemorrhage. ANESTHETIC DEPTH

Anesthetic depth depends on the anesthetics used, drug dosage, species, presence or absence of disease, and physiologic status. Crossspecies assessment of depth is facilitated by subdividing anesthesia into its subcomponents of unconsciousness or amnesia, analgesia, and muscle

PERIOI'ERATIVE SUPI'O RTTVE CARE AN D MONrTORI NG

589

relaxation. 10 The presence of adequate muscle relaxation (i.e., immobilization) does not necessarily imply unconsciousness or analgesia, however. Conversely, unconsciousness does not imply adequate muscle relaxation or analgesia. Determination of unconsciousness is difficult. Electroencephalography is used in humans and has been evaluated in research and some domestic animals. Unfortunately, this technology is not available for routine use or has not been validated in exotic species. Clinically, an assumption of unconsciousness is based on anesthetic dosing and vaporizer setting, muscle relaxation, decreased reflex activity, and absence of iimb and body movement. Increasing depth toward a surgical plane of anesthesia is assumed when muscle tone (e.g., jaw muscle or anal sphincter) decreases, palpebral and corneal reflexes are obtunded, and respiration pattern becomes regular and even. The corneal reflex and anal tone are not lost at surgical anesthetic levels. The palpebral and corneal reflexes are not elicited in snakes and some lizard species because the eyelids are fused to form a spectacle that covers the cornea. These reflexes are also difficult to assess in rodents because of small ocular size. Changes in eye position with anesthetic level are used in mammals but are determined for each species and validated for each individual patient as anesthesia is deepened and lightened. A fixed dilated pupil, unresponsive to light, and with no corneal reflex is a cross-species indicator of excessive anesthetic depth or brainstem hypoperfusion or ischemia. Sudden erection of the feathers in an anesthetized bird is a reflection of cardiac arrest or hypoperfusion and not of awakening. Pain is the conscious perception of nocioception, the activation of specialized receptors in response to a noxious stimulus. All vertebrates are assumed to have nocioceptors and potentially the ability to perceive pain. Pain and nocioceptor response are assessed by evaluating physiologic and muscular response to a painful stimulus. The difficulty arises in providing a stimulus reflective of surgical pain that does not unintentionally traumatize tissues. This can be a toe, an ear, cloacal (reptiles) or tail pinch, a skin incision, feather plucking in birds, or visceral manipulation (during a laparotomy). In reptiles, the stimulus should be applied for a minimum of 10 seconds to allow for a delayed response. Plucking a bird's feathers is more painful than is a skin incision. A surgical plane of anesthesia is assumed when no skeletal muscle m ovement occurs and physiologic changes are minimal or absent. Sudden tachycardia, hypertension, or tachypnea in response to stimuli are indicative of inadequate anesthetic depth or analgesia. Appropriate responses include stopping the painful stimulus, graded increases in anesthetic dose (e.g., increased vaporizer setting), or parenteral administration of an analgesic, usually an opioid. Although most inhalant anesthetics produce unconsciousness, they are poor analgesics. An exception is nitrous oxide, which is a good analgesic but alone does not produce unconsciousness.

590

HEARD

Inhalant anesthesia (usually isoflurane) is the basis for most small exotic animal anesthetic regimens. Minimum anesthetic concentration (MAC), or dose (MAD), in birds 1' 19 is a measure of inhalant anesthetic potency. MAC is defined as the anesthetic concentration (volume%) that produces immobility in 50% of an anesthetized population subjected to a noxious stirnulus. 1K 2 ' Knowledge of the MAC value in one species is used to approximate vaporizer settings for surgical anesthesia in other species. MAC is similar within and across animal classes. 25 Maintenance surgical anesthesia vaporizer settings are approximately 25% highe r than MAC. For example, the measured MAC of isoflurane usually lies between 1.5% and 2.0%, whereas maintenance vaporizer settings range from 2.0% to 2.5%. Several factors must be accounted for when one is using MAC as a guide to vaporizer setting. Hypothermia decreases MAC. 25 In poikilotherms (i.e., reptiles and amphibians), patients with a body temperature lower than their preferred optimum require a lower maintenance anesthetic setting. Although less dramatic, similar effects are observed in hypothermic birds and mammals. Premedication with opioids and other premedicants also decreases MAC. For example, butorphanol, 1 mg / kg, reduced MAD in cockatoos and African grey parrots by 25% and 11 %, respectively but had no significant effect in blue-fronted Amazon parrots.3• 4 MAC is not used to predict induction and recovery times because it is measured when inhalant anesthetic levels in the lower respiratory tract have equilibrated with those in the CNS. Induction and recovery times are primarily determined by inhalant anesthetic tissue and blood solubilities, ventilatory efficiency, and circulation time. Hence, isoflurane inductions are more rapid than are those with halothane because of the lower blood and tissue solubilities of isoflurane. Reptiles with inefficient respiratory systems and slow circulation times take longer to induce with an inhalant anesthetic than do mammals and birds. Another complicating factor in reptiles (particular aquatic and semiaquatic species) and amphibians is the presence of cardiac shunts. These shunts bypass part or occasionally all perfused pulmonary tissue, with consequent delays in inhalant anesthetic induction or recovery. During inhalant anesthetic induction, snakes relax from the head to tail and conversely recover in the opposite direction. Hence, the absence of a response to a tail pinch is suggestive of an adequate surgical plane of anesthesia. Similarly, the toe pinch response is used to monitor anesthetic depth in lizards, crocodilians, tortoises, and turtles. The palpebral or corneal response is used in crocodilians, tortoises, turtles, and most lizards. A classification system for assessing anesthetic depth in avian patients has been described. 1· u Anesthetic depth in birds and small mammals is assessed using the pinch response, palpebral and corneal reflexes, eye position, jaw tone, respiration rate and depth, and heart rate response.

PERIOPERATTVE SUPPORTIVE CA RE AN D ivlOr\lTORING

591

CARDIOVASCULAR MONITORING AND SUPPORTIVE CARE Anatomy and Physiology

Amphibian and most reptilian hearts are composed of three chambers: two atria and one ventricle. 33 The single ventricle functionally acts as two ventricles, with a variable amount of right-to-left shunting. Increased shunting bypasses the lungs with decreased systemic arterial oxygenation. Crocodilian, avian, and mammalian hearts are composed of four chambers: two atria and two ventricles; however, crocodilians also can shunt blood from right to left through a vascular foramen. 33 Vertebrate hearts are located on or slightly to the side of midline in the cranial third of the body cavity. The ventricle(s) are directed between ventral and caudal, and between midline and left lateral. In lizards, cardiac location varies from between the forelimbs (e.g., iguanas, chameleons, skinks, and water dragons) to almost the middle of the body (e.g., monitor lizards, tegus).22 In snakes, cardiac position is influenced by the predominant lifestyle; the distance from heart to the head is ranked arboreal < terrestrial < aquatic. 17 Cardiac location is determined by turning the snake and visualizing the movement of the ventral scales over the heart associated with cardiac contraction. This is facilitated by light reflecting off the scales. The avian heart is enveloped by the liver lobes and protected ventrally by the wide flat sternum. The mammalian heart (left ventricle) extends cranially from the point where the left elbow lies over the costochondral junction. Reptilian, avian, and mammalian heart rates are primarily determined by temperature, size, metabolism, respiratory state, and the presence or absence of painful stimuli. 33 Hypothermia is most important in reptiles because of the wide range of body temperatures encountered. 1--Ieart rate is inversely related to body size.27 The resting heart rate (min - 1 ) for m ammals is calculated from the allometric equation 241 X Mb- cm

where M b = bodyweight (kg).27 The equation for birds is 155.8 X M b - 0 23 • A heart rate 20% above or below the calculated rate for an individual patient is considered to be either tachycardic or bradycardic, respectively.30

Principles of Cardiovascular Monitoring

The main function of the cardiovascular system is oxygen (0 2 ) and nutrient delivery to tissue cells, and carbon dioxide (CO 2 ) and waste (e. g., lactate) removal. Adequate cardiovascular function implies suffiient capillary blood flow to fulfill these functions. Unfortunately, no ·eliable, accurate, and cost-effective way to measure tissue capillary flow :·xists. Although cardiac output can be measured, it is not feasible for

592

HEARD

routine monitoring in veterinary practice. Systemic arterial pressure measurement provides an indirect assessment of cardiac output, and, by extension, tissue perfusion. Increased peripheral resistance (e.g., increased arteriolar tone in response to epinephrine) results in apparently adequate systemic arterial blood pressure (i .e., in mammals a mean arterial pressure 2: 60 mm Hg), but decreased or absent capillary blood flow to one or more tissue beds (e.g., kidneys or gastrointestinal tract). Other indirect indicators of peripheral perfusion include capillary perfusion time, mucous membrane color, urine production, and blood gas analysis. Heart rate is one determinant of cardiac output, and marked tachycardias, bradycardias, and arrhythmias (e.g., ventricular tachycardia, fibrillation, third-degree heart block) can therefore decrease flow. Auscultation

Auscultation is used to determine heart rate and rhythm and to detect and assess the presence of cardiac murmurs. Heart sounds usually are not auscultable in amphibians and reptiles. In birds, the large pectoral musculature and broad sternum obtund the cardiac sounds. Avian and mammalian hearts are auscultated on either side of the chest and at the thoracic inlet. An esophageal stethoscope is used during all anesthetic procedures in which the patient is intubated. (Alternatively, a high-quality pediatric stethoscope is usually available for immediate use.) The esophageal stethoscope is an inexpensive, simple monitor of heart and lung sounds. It consists of a catheter fitted with openings in the distal 2 cm to 3 cm, which are covered by a rubber cuff. More sophisticated versions incorporate ECG electrodes. Esophageal stethoscope placement is difficult in birds with complex crops (e.g., parrots and pigeons). The lubricated tube is passed down the esophagus on the right side of the neck until it reaches the crop. The tip is then palpated through the crop wall and directed to the midline to pass through the thoracic inlet into the thoracic esophagus. The esophageal stethoscope is best placed soon after anesthetic induction, rather than during an emergency. It is impractical to use an esophageal stethoscope in small rodents, and it can induce regurgitation in guinea pigs. Capillary Refill Time

Capillary refill time is an indirect measure of peripheral tissue perfusion. Concurrent assessment of mucous membrane and skin color can indicate anemia (pallor), endotoxemia (hyperemia then pallor), and hypoxemia (blue or purple). Interpretation of mucous membrane color must allow for the diversity of normal pigmentation in exotic patients. The tongue tip of many iguanid lizards is a dark red.

PERIOPERATIVE SUPPORTIVE CARE AN D MONITORING

593

Anesthetic drugs affect interpretation of color and refill time; cx-2-adrenergic agonists (e.g., xylazine, detomidine, and medetomidine) produce intense peripheral vasoconstriction and pale mucous membranes. Similarly, hypothermia and hypovolemia or hypotension produce vasoconstriction. Inhalant anesthetics can produce an initial vasodilation and apparent congestion of mucous membranes and skin (e.g., reddening of the facial patches in macaws). Although normally pale, reptile mucous membranes become congested when the head is placed lower than the body or the neck is compressed by constrictive bandages. To determine capillary refill time, the mucous membrane of the oral cavity is digitally compressed until blanched, and the time it takes to return to its original color is d etermined. This time should be 2 seconds or less in birds and mammals; a prolonged capillary refill time is indicative of poor peripheral tissue perfusion. Electrocardiography

ECG monitoring is indicated for routine monitoring and w henever an abnormal pulse or arrhy thmia is detected or when the history suggests the possibility of an arrhythmia (e.g., syncope). The ECG should have a multichannel oscilloscope with nonfade tracing and freeze capabilities. Also, it must be able to record at speeds of 100 mm / s and amplify the signal to at least 1 m V equal to 1 cm. The ECGs of mammals, birds, and reptiles resemble each other in general form, with clearly defined P, QRS, and T components. In reptiles, an SV wave can be observed preceding the P wave. 34 Standard lead positions described for cats and dogs are used for small mammals, birds, and reptiles. Traditional positioning does not provide adequate wave deflections in reptiles w ith low signal voltages, however. Electrodes are placed in the cervical region in lizards with hearts located at the pectoral girdle. 22 In snakes, the electrodes are placed two heart lengths cranial and caudal to the heart. 22 In tortoises and turtles, the cranial leads are placed on the skin between the neck and the forelimbs. 22 To improve signal detection, stainless steel suture is passed through the skin and attached to the electrodes. In birds, electrodes are attached to the prepatagial regions of the wings and on the medial thigh regions. 26

Doppler Flow Detection

Doppler flow detection, used for audible monitoring of blood flow in unconscious or cardiovascularly unstable animals, is based on the Doppler principle, which states that the frequency of transmitted sound w aves is altered when reflected off moving red blood cells. 2 The magnitude and direction of the frequency shift is related to the velocity and direction of the cells and is converted into an audible sound. The

594

HEA RD

Doppler flow detector uses a probe placed as close as possible to blood flow in an artery or the heart. The probe contains the high-frequency sound transmitter and the receiver. Three types of probe exist: (1) human adult and (2) pediatric and (3) pencil. The pencil probe has a small probe surface at the end of an elongated holder and is valuable for assessing blood flow in small patients. Decreases in volume indicate a decrease in blood flow or displacement of the probe. The animal always should be checked first when the Doppler does not seem to be working. The Doppler is used wherever major arteries are close to the skin. In small mammals and reptiles, contact sites include the ventral aspect of the tail base; the carotid, femoral, and auricular (rabbit, Fig. 2) arteries; and directly over the heart. In birds, contact sites include the ulnar (Fig. 1), metatarsal, and carotid arteries. In chelonians (i.e., turtles and tortoises), the pencil probe is placed at the thoracic inlet to detect cardiac flow. An alternative site is the eye, to detect flow in the optic artery. Arterial Blood Pressure Measurement

Arterial blood pressure measurement usually is not performed in exotic animals because of small patient size, level of expertise required, and paucity of comparative data validating available equipment. The mean arterial blood pressure is a better indicator of tissue perfusion

Figure 1. Placement of the probe of a Doppler flow detector over the ulnar artery of a cockatiel. Note that the probe has been secured with a splint made from two tongue depressors .

PER IOPERATf VE SUPPORT I VE CARE A ND MONITORING

595

Figure 2. Rabbit ear demonstrating the central position of the auricular artery. This artery is used for direct arterial blood pressure measurement or placement of a Doppler flow probe. The vessels around the edges are marginal ear veins.

than systolic or diastolic pressure. The two forms of systemic arterial blood pressure measurement are indirect and direct. Indirect measurement is least accurate and most appropriate for physiologically stable patients. The three main techniques used are oscillometric, automatic, and Doppler.2· 13 The oscillometric technique uses a manometer within the occlusion cuff to d etect oscillations of the encircled artery. As cuff pressure increases to more than systolic pressure, a rterial blood flow ceases. As cuff pressure slowly decreases to less than systolic pressure, intermittent blood flow is detected as needle oscillations on the manometer roughly corresponding to systolic pressure. Recorded needle oscillations on the manome ter reach maximal amplitude at a pressure corresponding to the mean arterial pressure. The oscillations then decrease in amplitude below the mean pressure. Diastolic pressure is read at the point where needle oscillations no longer d ecrease in magnitude and is the least accurate of the three

596

HEARD

measurements. The automatic technique uses the same methodology as the oscillometric technique, except that the cuff is inflated and deflated automatically. With the Doppler technique, as cuff pressure slowly decreases and pulsatile arterial blood flow begins, an audible Doppler signal is detected corresponding to systolic arterial blood pressure. Diastolic arterial blood pressure is indicated by an abrupt muffling or cessation of sound as the vibrations of the arterial wall cease. Sites for indirect blood pressure measurement include the legs, forearms, tail, and ears (in rabbits). Indirect measurement is least accurate at low systemic pressures and when small arteries are used. One must have the appropriate cuff size; its width should be approximately 40% of the circumference of the limb. 2 Narrow and wide cuff widths provide erroneously high and low pressure measurements, respectively. 2 Direct arterial blood pressure measurement is indicated in hemodynamically unstable (shock) patients, patients receiving vasoactive drugs, and in critical or cardiac patients undergoing anesthesia or surgery. It is used less frequently than indirect blood pressure measurement because it is technically difficult and requires an expensive monitor. It is most easily performed in rabbits because of the easily accessible auricular artery in the center of the pinna. In birds weighing 1 kg or more, arterial catheters are placed percutaneously in the ulnar or the dorsal metatarsal artery. The advantages of direct blood pressure measurement are that it gives a constant reading and the catheter can be used as a source of arterial blood for gas analysis.

Central Venous Pressure

The central venous pressure (CVP) reflects intravascular volume, cardiac function, and venous compliance. Its measurement is indicated in patients with marginal cardiac function, with decreased glomerular filtration rate, or those receiving large volumes of fluids. Although not commonly used, it can be determined whenever a jugular cathe ter tip extends close to the junction be tween the cranial vena cava and the right atrium. 13 A three-way stopcock is attached to the jugular catheter, to the administration set of the animal's intravenous fluids, and to a vertically positioned manometer. The manometer is zeroed to the level of the right shoulder (sternal recumbency) or center of the thoracic inlet (lateral recumbency). When the manometer is filled from the fluid bag and the stopcock is opened to the jugular catheter, the fluid in the manometer slowly drops and levels out at CVP. This procedure is repeated at least three times and the average recorded in cm H 20. Single CVP m easurements do not reflect an animal's hemodynamic status; changes with time are most important. Normal mammalian CVPs range from 0 to 5 cm H 20.

PERJOPERAT!VE SUPPORTI VE CA RE AND MONITO RI NG

597

Dehydration

Determination of relative hydration involves integration of data from several sources, including history, physical examination, and blood work. Although skin tenting is emphasized in domestic animals, it is difficult to interpret in exotic patients, given the wide range of skin types. Gently grasping a fold of skin and rolling it between fingers is a more reliable technique. Normally hydrated skin moves easily, whereas in dehydrated animals, a "sticky" feeling exists. Dehydration also produces dry mucous membranes, sunken eyes, and decreased tear production. The packed cell volume (PCV), total protein level, plasma osmolality, and electrolyte concentrations often are increased. The urine specific gravity usually is increased, and urine production usually is decreased. Dehydration is graded as mild (2% to 4%), moderate (6% to 8% ), or severe (10% to 12% ). Fluid Therapy

Vascular access is established in physiologically unstable patients and patients likely to decompensate from hemorrhage, endotoxemia, or other complications during the perianesthetic period. Few published studies have evaluated the most appropriate fluid and minimal and maximal fluid rates in anesthetized small exotic patients. 20 Small patient size makes venous and intraosseous catheterization difficult, but practice and attention to technique enable attainment of these essential skills for small exotic animal practice. Catheterization sites are described in Table 1 and illustrated in Figures 3 to 8. The author prefers 20- to 24-gauge over-the-needle catheters for venous catheterization. Penetration of the skin with a hypodermic needle or scalpel blade before placement is done to prevent catheter buckling. Catheterization requires steady, directed movement in small, incremental steps. A change in resistance is often the first indication of correct placement before blood is observed in the catheter hub. The catheter and stylet are advanced 0.5 mm to 2.0 mm into the vessel before threading. In thin-skinned animals (i.e., birds and some small mammals, particularly amelanistic breeds) and where a cut-down is performed, the catheter is visualized inside the vessel before advancement off the stylet. Intraosseous catheterization is performed in lizards, birds, and small mammals (Table 2, Figs. 9 and 10). Spinal needles are recommended in all but the smallest patients to prevent blockage of the needle bore with a bone plug. Needle length is approximated b y measuring half of the length of the bone. All intraosseous catheter sites are cleaned and aseptically prepared for introduction of the needle. The bone selected for catheterization is grasped firmly in one hand, and the needle is inserted with a twisting of the wrist using the landmarks described here. Successful placement is indicated by a sudden change in resistance and rapid advancement of the needle, a "grating" feeling as the needle passes

598

HEARD

Table 1. INTRAVENOUS CATHETERIZATION SITES FOR SMALL EXOTIC PATIENTS Class

Order

Site

Amphibia

An uria (frogs and tuads)

Ventral abdominal: Located on ventral midline. Percu taneou s catheterization or a small mid line skin incision to visualize vessel.

Reptilia

Squa mata (snakes)

Palatine: Loca ted paralle l and medial to the palatine teeth in the roof of the mouth (Fig. 3). Care must be taken to prevent injury to fingers. Medium to large snakes. Jugular: Right larger than the left. A cutdown incision is made 4 to 7 scutes cran ial to the heart at the junction of the ventral scutes and the right latera l body scales. The vein is identified by blunt dissection just medial to the ti ps of the ribs." Heart: Ind icated for short procedures or emergencies. Not recommend ed by the author.

Squamata (li zards)

Cephalic: Located on the dorsal (anterior) su rface of the distal foreleg. A skin incision from the elbow distal and medial over the dorsal forearm to allow visualiza tion of vein _12 Ventral abdominal: Located on ventral midline (Fig. 3). Can be entered percutaneously or following a small skin incision in the midline to visualize vessel.

Chelonia (tortoises and turtles)

Crocodilia

Jugular: Located on la teral neck, more dorsal than would be expected in mammal. Requires a long itudinal incision and blunt dissection to visualize vessel. PPV can assist distension of vessel for cathcterization. Jugular: Located on the lateral surface o f neck at the level of the ear (Fig. 4). In some anima ls can be percutaneously catheterized . In h ypovolemic / h ypotcnsi ve an imals a longitudin al skin incision is required for visualization. PPV can assist distension of vessel for catheterization. Jugular: Located on ventrolateral surface of neck. Requires lon gitudinal incision and blunt dissection to visua lize.

Table continued on opposite page

PERJOPERATIVE SLPrORTIVE CA RE AND MO NITOR l.\iG

599

Table 1. INTRAVENOUS CATHETERIZATION SITES FOR SMALL

EXOTIC PATIENTS (Continued). Class

Order

Site

Basilic: Located on the medial surface of the e lbow. Used temporarily during anesthesia.

Aves

Jugular: Righ t jugular vein larger than left (Fig. 6). Located right dorsolateral of surface in area absent of feath ers. Percutaneously catheterized. Dorsal and superficial plantar metatarsal: Located on dorsa I surface of tarsometata rsus as it runs media lly across the hock and the inside of the leg (Fig. 7). Usually requires plucking some feathers. Use butte rfly catheter or small needle (25- 27 gauge) in bi rds < 500 g, over-the-needle ca the ter (24-22 gauge) in birds > 500. Cranial and caudal tibial: Located on the cranial and caudal tibiotarsus of large long-legged birds.

Mammalia Lagomorpha (rabbits), Carnivora (ferrets)

Jugular: Sarne position and technique as in cats and dogs. May require a cutdown to identify vessel.

Lagomorpha (rabbits), Rodentia (guinea pigs), Carnivora (ferrets, skunks), Primates

Cephalic: Passes medial to lateral over the dorsa l (cranial) surface of the forearm (Fig. 8).

Primates, Lagomorpha (rabbits), Rodentia (guinea pigs)

Lagomorpha (rabbits) Rodentia (rats), Ma rsupia li a (opossum), Primates

Saphenous: Particularly useful in small primates, where it runs along the caudal aspect of the leg. Aural: Located on the edge of the ea rs (Fig. 2). Used in large rabbits. Potential for ea r ischemia if thrombosis occurs. Coccygeal: Lateral surfaces of tail. Can require b utterfl y cathe te r or smallgauge catheter.

600

HEARD

Figure 3. Jugular catheterization in a yellow-footed tortoise. A 22-gauge , over-the-needle catheter was placed percutaneously, attached to an injection cap, and sutured in place. A 19-gauge butterfly catheter is being used to infuse fluids. Note the relatively dorsal position of the jugular vein on the lateral neck.

through medullary bone, and ease of infusion of a test injection of heparinized saline. Intraosseous catheters are maintained for up to 3 days before removal. A technique has been described in tortoises using the bony bridge between the carapace and the plastron. 12 After investigating this technique in living animals and with dissection of cadavers, however, the author believes that the catheters merely are infusing fluid into the coelomic cavity. Subcutaneous administration of fluids is an inappropriate route for correction of deficits or replacement from hemorrhage in the perianesthetic period. Fluid absorption is minimal because subcutaneous tissues are poorly vascularized, and peripheral vasoconstriction is the usual response to dehydration and hypotension. Hypertonic dextrose solutions administered subcutaneously can exacerbate dehydration and fluid deficits. 20 Balanced electrolyte solutions (e.g., 0.9% sodium chloride or lactated Ringer's solution) are used for routine fluid administration. An argument has been made not to use lactated Ringer's solution in reptiles because of the prolonged half-life of lactate. 24 It is suggested that a 50:50 combination of 5% dextrose and a nonlactated multiple electrolyte solution, such as Normosol-R or Plasma Lyte, be used .24 In mammals and birds, blood glucose levels of 60 and 200 mg / dL, respectively, are indicative of hypoglycemia. Insulinomas are common in ferrets, and the

PERIOPERATIVE SUPPORTIVE CARE AND MONITORING

601

Figure 4. Ventral abdominal vein catheterization in a red tegu. A small midline skin incision was made to visualize the vein and a 22-gauge, over-the-needle catheter introduced and secured to the body wall.

Figure 5. Palatine vein catheterization in medium to large snakes. A 19-gauge butterfly catheter has been placed in the large palatine vein of an amelanistic Burmese python. Note the location of the vein just medial to the palatine teeth.

602

HEARD

Figure 6. Metatarsal vein catheter in a sulphur crested cockatoo.

Figure 7. Catherization of the right jugular vein in a cockatiel. A 24-gauge, over-the-needle catheter was placed percutaneously and secured temporarily with tape.

l'ERIOPERATJVE SU PPORTIVE CARE AND MONTTORlNG

603

Figure 8. Catheterization of the cephalic vein in a guinea pig. A 24-gauge, over-the-needle catheter was placed percutaneously before surgery. Note the wooden splint to support the catheterized leg.

blood glucose level should be checked routinely preoperatively. Blood glucose levels in reptiles range from 30 to 100 mg/ dL or more, depending on species. Isotonic dextrose solution (5%) or combinations are indicated when hypoglycemia is present or expected. Selection of the appropriate fluid to infuse in hemorrhaging patients depends on the severity and duration of hemorrhage, the initial hematocrit, the species, and the presence of underlying cardiopulmonary disease. The hematocrit and total protein do not reflect hemorrhage severity acutely. A general guideline to fluid selection is for 5% to 10% of blood volume lost: balanced electrolyte solution at three times the volume of the estimated blood loss should be used. At 10% to 20% loss, plasma expanders (e.g., hetastarch or plasma) should be used; and for 20% to 30% loss, whole blood transfusion should be used. Preferably, blood is collected before anesthetic induction into heparinized syringes or acid citrate phosphate dextrose anticoagulant. Transfusions are confined to the same species. Ideally, a cross-match is performed, but a first transfusion is usually safe in small exotics of the same species. Small-volume infusors are commercially available (e.g., Medfusion 3 2010i, Medexing, Duluth, GA) but are expensive. They are essential for accurate fluid infusion in small anesthetized patients, however. Some infusors can be preprogrammed to flow rates for emergency and other drugs, so that all that is necessary for administration is to enter the body weight of the animal. They also allow for a continuous infusion, which is preferable to bolus injection. The rate of fluid infusion during anesthesia depends on hydration status, daily fluid requirement, severity of hemorrhage, type of fluid to be infused, and the presence or absence of underlying renal or cardiac

604

HEARD

Table 2. TECHNIQUES AND SITES FOR INTRAOSSEOUS CATHETERIZATION IN SMALL EXOTIC PATIENTS Animal Class

Order

Amphibi a Reptilia

Aves

Site

Distal femur Sguamata (lizards)

Distal femur: Flex stifle, curve in dis tal femur usually allow s catheter to be introduced proximal to the joint (Fig. 9). Proximal tibia: Differentiate from lateral fibula. Pass ca theter through tibial crest and pass need le to med ial surface of leg as it is passed in to the bone. Proximal humerus

Chelonia (tortoises)

Carapace/plastron bridge? Probably catheter is placed in to coe lomic ca vity.

Chelonia (sea turtles)

Distal humerus 16 ? Animal placed in sternal recumbency. From the front grasp left or right forefl ip per, insert need le in distal 1/ , of the medial aspect of the humerus at an angle of approx. 30°-45° from parallel. Needle inserted as far d ista lly as possible w ithout en tering joint capsule. Tu rtle bone is very dense and d ifficult to introduce ca theter. Distal ulna: Lateral surface of bone (Fig. JO). Use condylar rid ge as entry point. Placement is confirmed by test dose of heparinized saline and observing the basilic vein clear. Proximal ulna: Count three to four fli gh t feathers in from the elbow. Introduce catheter at a steep angle to start, then change to acu te angle as it is inserted. Proximal tibiotarsus: Flex stifle, introduce catheter through tibia l crest

Mammalia

Proximal femur: Adduct femur, pass catheter into the trochanteric fossa. Proximal tibia: Flex stifle, introduce catheter through tibial crest. Proximal humerus: Flex shoulder, introduce catheter through the greater tubercle.

l'ERTOPERATIV E SUl'PORTIVE CARE AND MONITORlNG

605

Figure 9. lntraosseous catheterization of the femur in a bearded dragon. The curve of the distal femur allows for needle placement without entering the stifle.

disease. During most anesthetic episodes, the main goal of fluid therapy is to maintain vascular access and replace fluids lost to dry anesthetic gases. The author usually infuses a balanced electrolyte solution at 5 to 10 mL/kg/h.

Drugs

Pharmacologic manipulation of cardiovascular function occasionally is indicated in anesthetized patients. The main difficulties are in early detection of instability and subsequent monitoring of drug effect. Positive inotropes, chronotropes, and vasoconstrictors (e.g., dopamine, dobutamine, ephedrine, epinephrine, and norepinephrine) can be used. Without arterial blood pressure measurement to adjust infusion rates, however, these drugs can cause detrimental hypertension or arrhythmias. The use of small-volume infusors described earlier is essential for accurate administration of these drugs in small patients.

606

HEARD

Figure 10. lntraosseous catherization of the distal ulna in the skeleton of a pigeon. Note the alignment of the needle with the long axis of the ulna. The entry point for the needle is medial to the dorsal condylar ridge.

Cardiopulmonary Resuscitation

Cardiopulmonary resuscitation in small exotic patients follows the same priorities as other veterinary patients (i.e., airway, breathing, circulation, and drugs). The p eriod of ischemia or hypoxia that the brain can tolerate before irreversible damage is incurred is indirectly related to metabolic rate and body temperature. Consequently, small mammals and birds have a shorter window of time in which resuscitation is effective than do cats and dogs because of their higher metabolic rates. To improve response time, emergency drug dosages are precalculated or drawn into labeled syringes in preparation for use. Because of their lower metabolic rates and body temperatures, reptiles and amphibians are much more tolerant of ischemia or hypoxia. Hence, resuscitation attempts can be successful after prolonged cardiac arrest (e.g., > 20 min in green iguanas and ball pythons).

PERJOPERATIVE SU PPORTI VE CARE AN D MON ITORIN G

607

RESPIRATORY MONITORING AND SUPPORTIVE CARE Anatomy and Physiology

The respiratory system shows the greatest morphologic and physiologic diversity among animal classes. Most vertebrate lungs, however, are paired with the gas-diffusing surfaces in the dorsocranial half of the body cavity. Reptile lungs are saclike, with variable degrees of compartmentalization to increase surface area for gas diffusion. 32 The left lung of snakes (except boas and pythons) is small or absent. 32 Although it is an elongated sac that extends for 50% or more of the coelomic cavity, it is only the cranial portion that participates in gas exchange. This diffusing surface is located cranial to the heart in arboreal snakes, an adaptation that is thought to reduce pulmonary edema in a vertical position.17 The avian respiratory system, which is the most efficient of any vertebrate group,7 is characterized by paired lungs located in the craniodorsal coelomic cavity, against the thoracic vertebrae and ribs, and is connected to multiple air sacs that act as reservoirs and participate in air movement. Gas diffusion occurs in the small air capillaries connected to the terminal bronchi or parabronchi. Birds do not possess a diaphragm.7 The avian air sacs do not preclude the use of positive-pressure ventilation (PPV); air sac membranes would not likely be ruptured by all but high pressures. Mammalian lung structure is familiar to veterinary practitioners and is not discussed here. In mammals, inspiration is active and expiration is passive at normal ventilation volumes. In contrast, reptiles and birds have active expiration and inspiration.7· 34 In all reptiles, the airflow during the ventilatory cycle is biphasic, beginning with exhalation followed by inhalation. 34 Although reptile ventilatory patterns are diverse, they can be characterized as arrhythmic, with significant nonventilatory periods, or apnea. 32 The two basic patterns are single breaths separated by periods of breath holding (terrestrial species), and episodes of consecutive lung ventilations followed by long nonventilatory periods, lasting from a few minutes to more than an hour (aquatic species). In birds, most inspired air completes a full circuit through the lungs and air sacs in two breaths. 7 Respiration rate (RR) is inversely related to body weight. Resting RR (min - 1) for mammals is calculated from the allometric equation 53.5 X Mb - 0 ·26 where Mb

body weight (kg). 27 The equation for birds 27 is 17.2

X

Mb- 03 1

An RR 20% more or less than the calculated rate for an individual patient is considered tachypneic or bradypneic, respectively. 30 Respiration in reptiles under anesthesia is slow, ranging from 1 to 5 breaths / min to 1 breath/ 3-5 min. No studies to evaluate the advantages of spontaneous \'Crsus assisted ventilation in reptiles have been performed.

608

HE A RD

Auscultation

As with cardiac auscultation, evaluation of the respiratory system in small patients requires a good-quality stethoscope with appropriate head and length of tubing and an experienced ear. The esophageal stethoscope can be used to evaluate respiratory noise; however, artifactual noise must be distinguished from true respiratory noise. Blood Gas Analysis

Arterial blood gas analysis assesses patient oxygenation, acid-base status, and adequacy of ventilation. Venous blood gas samples are more easily obtained than are arterial samples, but interpretation is difficult. They reflect arterial CO 2 and 0 2 levels only crudely, and are affected by local tissue metabolic activity and low blood flow. Cardiocentesis blood gas samples in turtles 14 and other reptiles are not an accurate reflection of systemic (i.e., carotid) arterial values because of intracardiac shunting. Arterial blood gas analysis is used periodically to assess the accuracy of pulse oximetry or capnography during surgery. The principles of blood gas analysis are well described by Muir and deMorais.21 Blood gas values are corrected to the patient's body temperature or analyzed at a standardized temperature. 21 Arterial blood gas samples are obtained from any palpable artery. Collection sites include the femoral, metatarsal, ulnar (birds), and auricular (rabbit) arteries. Infiltration of the periarterial area with 1% lidocaine without epinephrine can prevent reflex vasoconstriction. Samples are collected into heparinized 1-mL or 3-mL syringes through a 25-gauge or 27-gauge needle. After collection, all air is evacuated from the syringe, a cork placed on the needle, and the sample processed as soon as possible. The sample syringe is placed in crushed ice if the time between collection and analysis is prolonged. Avian and reptile red blood cells have a high metabolic activity, so that immediate sample processing is recommended. 34 The blood gas analyzer i-STAT (Heska, Waukesha, WI) is a portable, cost-effective alternative to traditional analyzers and requires relatively small blood volumes for analysis. The main problems with this machine are that it works only in a narrow environmental temperature range and it has not been critically evaluated for use in exotic species. Pulse Oximetry

Pulse oximetry is indicated for monitoring blood oxygenation and controlling 0 2 administration. The latter allows for administration of the lowest concentration of inspired 0 2 compatible with safe levels of arterial oxygenation. Commercial pulse oximeters also measure pulse rate. Pulse oximeters estimate arterial hemoglobin 0 2 saturation (Sa0 2 ) by measur-

PERIOPERATIVE SUPPORTIVE CARE AND MONITORING

609

ing pulsatile signals across (transmission) or by reflectance (reflection) from perfused tissue at two discrete wavelengths (660 nm, red; 940 nm, infrared) using the constant component of absorption (i.e., that caused by everything except arterial blood) at each wavelength to normalize the signals. 28· 31 A ratio (R) between these two normalized signals is calculated and related to arterial 0 2 saturation using an empiric algorithm. This R value varies from 0.4 to 3.4 over the saturation range of 0% to 100%. All commercially available pulse oximeters calculate hemoglobin saturations from algorithms developed by measuring the R in human volunteers and simultaneously sampling arterial blood for in vitro saturation measurements.31 An R of 1.0 corresponds to a pulse oximeter saturation reading (Sp0 2 ) of approximately 85%. 3 1 Potential sites for placement of transmission pulse oximeter sensors include the ear, tongue, buccal mucosa, paw (Fig. 11), vulva, prepuce, and tail. A reflectance pulse oximeter sensor is used in the esophagus, or the rectum or cloaca. Surprisingly, the best site in rabbits is the base of the tail rather than the ears, which could be because of excessive compression of the aural vasculature. Pulse oximeters require adequate plethysmographic pulsations to allow them to distinguish arterial light absorption. They are, therefore, inaccurate in the presence of decreased blood pressure, decreased pulse pressure, and vasoconstriction. The oximeter measures a pulse but not peripheral perfusion. 31 The presence or absence of a pulse is quickly detected, but the presence of a pulse does not ensure adequate blood flow. Pulse oximeter calibration assumes only two hemoglobin types in the bloodstream: (1) oxyhemoglobin and (2) hemoglobin. If methemoglobin or carboxyhemoglobin is present, it is interpreted as oxyhemoglobin, hemoglobin, or some combination of the two. 31 High methemoglobin

Figure 11. Pulse oximetry in a ferret. The sensor has been placed over the front paw.

610

HEARD

levels force Sp02 toward 85% regardless of Pa0 2 and Sa02 values. Methemoglobin levels in reptiles can be high in otherwise healthy reptiles .34 In turtles, m ethemoglobin levels of 15% or more are routine, and values of 90% or more are not uncommon. High methemoglobin levels also are found in lizards and snakes. Schmitt et aP evaluated two pulse oximeters, the Pulsox 7 and the Oxycount Mini (Andos, Hamburg, Germany) in pigeons, macaws, and African gray parrots. Sp02 values were compared with Sa0 2 values calculated from ulnar Pa0 2 using a published pigeon 0 2 hemoglobin dissociation curve. The probes were positioned over the gastrocnemius and tibialis cranialis muscles. The study demonstrated a different photom etric behavior for avian and human hemoglobin, which would be expected to result in an underestimation of saturation values in birds. Saturation values recorded by pulse oximetry did not correlate well (R = 0.81) with calculated Sa02 values. The investigators concluded that pulse oximetry is unsatisfactory for routine use in avian practice. Additional research is necessary to develop an avian calibration curve for pulse oximetry. Diethelm et aP evaluated pulse oximetry in green iguanas. Oxyhemoglobin and deoxyhemoglobin absorbencies at 660 nm and 940 nm were determined in vitro to be similar to human hemoglobin. Although iguana hemoglobin seemed to absorb more light at 660 nm than did human hemoglobin, it was thought to be a preparation artifact. Six anesthetized iguanas were administered inspired 0 2 concentrations of 7%, 10%, 20%, and 100%. A reflectance probe was placed in the esophagus and the measured (SDI Vet/Ox 4402, Sensor Devices Inc, Waukesha, WI) Sp0 2 values were compared with Sa02 values calculated from Pa0 2 values of blood samples obtained from the abdominal aorta. No significant differences were found between Sp02 and Sa02 values, and the investigators concluded that pulse oximetry is an excellent tool for monitoring pulse rate and oxygen saturation in iguanas. The study abstract5 does not mention how the Sa02 values were calculated and does not address methemoglobin concentrations. Additional research is necessary to evaluate pulse oximetry in other orders of reptile, as well as the effect of methemoglobinemia. 34

Capnography

Capnometry is the measurement of CO 2 concentrations in expired gases; 8 capnography refers to the display of these concentrations on an oscilloscope screen or recording chart, usually as a function of time. CO 2 is monitored by direct in-line measurement at the sample site by a flowthrough device or by aspiration of the gas sample into a separate monitor. In-line sensors are fragile, expensive, cumbersome, and have a relatively large dead space. 8 Analyzers that continually withdraw gas samples are plagued by excess moisture and require that the small-

PERIOPERATIVE SUPPORTIVE CARE AND MONITORING

611

diameter sampling tube be as short as possible to improve response time.8 End-tidal CO 2 tension (PETC02 ) is used to provide a noninvasive estimate of PaC0 2 • PETC02 is usually less than PaC02 . Lung disease increases this difference, as can sampling errors. The most obvious sampling error is a system leak. Another sampling problem is related to the sampling rate and the minute ventilation of the patient. When used in small patients, high sampling rates (i.e., 250 mL / min) can result in entrainment of room air and lower PETC02 values. In human pediatric patients, the sampling rate is set at 50 mL / min. This rate is still too high in many small exotic patients. Ventilation

General anesthetic agents usually produce a dose-dependent ventilatory depression. This effect can be additive or synergistic, with underlying disease resulting in marked hypercapnia or ventilatory arrest. Studies in birds6 and reptiles 29 sl1i.ow that high inspired 0 2 concentrations decrease ventilatory drive. Ventilation also is affected by body position and compression of the respiratory exchange tissues by distended viscera or obesity. Dorsal recumbency in awake chickens results in a 40% to 50% decrease in tidal volume and a 20% to 50% increase in respiratory rate, with an overall 10% to 60% decrease in minute ventilation. 15 Inadvertent compression of the chest by surgeons, common in small exotic patients, can be prevented by developing good hand position techniques, having an attentive anesthetist and using clear plastic drapes. Because of their very compliant chest walls, ferrets seem dependent on diaphragmatic movement for ventilation. Rabbits have small chest cavities relative to body size and have a high prevalence of respiratory disease. Also, they often develop tachypnea under anesthesia, with a normal to decreased alveolar ventilation. Guinea pigs have relatively small-diameter tracheas and are prone to airway obstruction because of regurgitation and profuse salivary secretions. The relatively long size of commercially available endotracheal tubes makes one-lung ventilation a common complication of endotracheal intubation of small mammals. Similarly, primates have short tracheas, facilitating endobronchial intubation. Prevention of endobronchial intubation requires that endotracheal tubes be premeasured to extend to just inside the thoracic inlet. Correct placement also can be determined by ventilating the animal and auscultating for lung sounds on both sides of the chest or obtaining thoracic radiographs. Adequacy of ventilation is assessed most accurately using PaC02 • Capnography provides an indirect estimate of PaC02 but is too inaccurate (see discussion earlier) in most small exotic patients to be used for anything other than validation of successful endotracheal intubation. Pneumotachography for measurement of tidal volume usually is not performed in exotic animal practice because of the large mechanical

612

HEARD

dead space and resistance, species variation, and cost. Visualization of chest wall movement is a deceptive guide to adequacy of ventilation. For example, several species of birds have been shown to develop mild to marked hypoventilation under inhalant anesthesia despite apparently normal chest wall movement. 18• 19 Consequently, the author recommends assisted or controlled ventilation of intubated patients under general anesthesia. Doxopram is not recommended for use in h ypoventilating patients unless intubation and mechanical ventilation is impossible or contraindicated. Routine endotracheal intubation is feasible in reptiles of 50 g or more, birds of 80 g or more, rabbits, ferrets, and primates. Tortoises and turtles can be ventilated by flexing and extending their forelimbs in and out of the shell. Small birds are ventilated in the absence of endotracheal intubation by rapidly flexing and extending their wings. The unique respiratory anatomy of birds also allows for intubation of an air sac when tracheal intubation is contraindicated (i.e., a partial obstruction) or a hindrance to surgery or diagnostic investigation. Similarly, 0 2 continuously administered through a large-gauge needle placed into an abdominal or caudal thoracic air sac can prevent hypoxemia and hypercapnia in apneic birds. Ventilation is assisted or controlled by positive pressure manually or mechanically. The principles and techniques of artificial ventilation are well described by Hartsfield. 9 Many of the ventilators used in smallanimal anesthesia can be modified to ventilate small patients. A commercially available combination ventilator and anesthesia machine (An esthesia WorkStation, Hallowell EMC, Pittsfield, MA) has been specifically designed for use in small patients, primarily research rodents and rabbits. The advantage of mechanical ventilation is that it frees the anesthetist to concentrate on other tasks. The disadvantages are that mechanical ventilators require a thorough theoretic and technical understanding for their safe use; they also require endotracheal intubation and p roduce positive intrathoracic (intracoelomic) pressures that interfere w ith venous return to the heart and can cause lung trauma. Manual ventilation provides the advantage, w ith a skilled anesthetist, of rapidly adjusting ventilatory pressures and volumes and a responsiveness to the surgeon who sometimes requires brief, irregular periods of ventilatory arrest to complete a surgery safely. Additional research is necessary to evaluate the efficacy and appropriateness of ventilation techniques in exotic patients. There could be situations in which some hypoventilation is preferable to the adverse effects of PPV. Oxygen Support

Administration of elevated inspired 0 2 concentrations (2:20% and -S:40%) often can overcome mild to moderate h ypoxemia, assuming that no major pulmonary shunting is present. Failure of the Pa02 to remain

PERlOPERATIVE SUPPORTIVE CARE AND MONJTORlN G

613

above 60 mmHg in the presence of an Fi02 2 40°/c, is an indication for intubation and ventilation in a mammal. Although both mammals and birds are susceptible to pulmonary 0 2 toxicity, this syndrome is unlikely to be observed in patients maintained on high inspired 0 2 concentrations for ::::: 24 hours. TEMPERATURE MONITORING AND SUPPORTIVE CARE

Body temperature measurement is a standard of care during all anesthetic procedures. Hypothermia is common in small anesthetic patients because of the large surface area-to-volume ratio. Also, many drugs used in the perianesthetic period suppress normal thermoregulatory mechanisms. The anesthetic gases used during inhalation anesthesia are of low humidity and temperature. In one report, 23 doves without thermal support lost 4°C to 5°C within 40 minutes of isoflurane anesthesia. Hypothermia is reduced by minimizing anesthesia time and the use of surgical preparation solutions, wrapping the body, increasing the room temperature, and using external heat sources. Electric heat blankets are not used because they may cause severe burns. Similarly, heated fluid bags placed in contact with the skin also can cause burns. In one report, 23 waterblankets were unable to prevent hypothermia in isoflurane-anesthetized doves, but a 680-W radiant-energy source (Radiant Ray, Infrared Heater, Model 251-C, Merco Products, Eugene, OR) maintained normothermia. Poikilotherms are immobilized at variable temperatures of less than their preferred optimal body temperature ranges, however, hypothermia alone is not a humane anesthetic method for these species. Although it produces immobility, whether it induces adequate analgesia for painful diagnostic and surgical procedures is questionable. Hypothermia impairs drug metabolism, depresses immune function, and decreases circulation time. Thermal support should be provided, with body temperatures being maintained within or slightly less than a reptile's preferred optimal temperature range. RECOVERY

Recovery is a critical period during which the patient is placed in a warm, quiet environment and monitored. Supportive care established during anesthesia is continued into the recovery period until the patient is fully alert and physiologically stable. Vascular access is maintained to allow for emergency administration of drugs and fluids . The duration and quality of recovery are determined primarily by the anesthetic agents used, the duration of the procedure, and the magnitude of physiologic dysfunction incurred. Prolonged recovery usu-

614

HEARD

ally is caused by hypothermia, hypoglycemia, anesthetic overdose, or impaired drug elimination. Caution must be exercised when rewarming an animal that is possibly hypovolemic or hypoglycemic because warming results in dilation of vasoconstricted peripheral vessels and increases the metabolic demand for glucose. These phenomena could explain some of the sudden deaths that occur a few hours into recovery. Recovering animals (especially reptiles) are unable or unwilling to move from areas of high h eat, and hyperthermia with associated burns or death can occur if temperatures are not monitored.

SUMMARY

Supportive care minimizes the adverse physiologic effects of anesthesia, surgery, and preexisting disease. Monitoring detects physiologic perturbations, ensures appropriate anesthetic depth, and assesses the efficacy of supportive care. The principles of monitoring and supportive care are similar in small exotic species; however, an understanding of comparative anatomy, physiology, and pathophysiology is necessary to apply these principles.

References 1. Arnall L: Anesthesia and surgery in cage and aviary birds (1). Vet Rec 73:139, 1961 2. Crowe DT, Spreng DE: Doppler assessment of blood flow and pressure in surgical and critical care patients. /11 Bonagura JD (ed): Kirk's Current Veterinary Therapy XII: Small Anima l Practice. Phi ladelphia, WB Saunders, 1995, p 113 3. Curro TG: Evaluation of the isoflurane-sparing effects of butorphanol and flunixin in psittaciformes. Proc Assoc Avian Vets, Reno, NV, 1994, p 17 4. Curro TG, Brunson D, Paul-Murphy J: Determination of the ED50 of isoflurane and evaluation of the ana lgesic properties of butorphanol in cockatoos (Cacatua spp.). Vet Surg 23:429, 1994 5. Diethelm G, Mader DR, Grosenbaugh DA, et al: Evaluating pulse oximetry in the green iguana, Ig1iana iguana. In Proc Assoc Reptilian Amphibian Vet, Kansas City, KS, 1998, p 11 6. Diethelm G, Mader DR: The effect of F1 0 2 on postanesthetic recovery times in the green iguana. /11 Proc Assoc Reptilian Amphibian Vet, Columbus, 1999, p 169 7. Fedde MR: Respiration. /11 Sturkie PD (ed): Avian Physiology, ed 4. New York, Springer-Verlag, 1986, p 191 8. Gal TJ: Monitoring the respiratory system. In Lake CL (ed): Clinical Monitoring for Anesthesia and Critical Care, ed 2. Philadelphia, WB Saunders, 1994, p 213 9. Hartsfield SM: Airway management and venti lation. /11 Thurmon JC, Tranquilli WJ, Benson GJ (eds): Lumb & Jones' Veterinary Anesthesia, ed 3. Baltimore, Williams & Wilkins, 1996. p 515 10. Haskins SC: Monitoring the anesthetized patient. /11 Thurmon JC, Tranquilli WJ, Benson GJ (eds): Lumb & Jones' Veterinary Anesthesia, ed 3. Balt imore, Williams & Wilkins, 1996, p 409 11. Heard DJ: Anesthesia and analgesia. /11 A ltm an RB, Clubb SL, Dorrestein GM, et al (eds): Avian Medicine an Surgery. Ph il ad elphi a, WB Saunders, 1997, p 807 12. Jenkins JR: Diagnostic and clinical techniques. /11 Mader DR (ed): Reptile Medicin e and Surgery. Philadelphia, WB Saunders, 1996, p 264

PERIOPERATIVE SU!'PORTI VE CA RE AN D MON ITORINC

615

13. Kaplan PM: Monitoring. /11 Murtaugh RJ, Kc1plan PM (ed s): Ve terinary Emergency and Cr itical Care Medicine. St Louis, Mosby -Yea r Book, 1992, p 21 1-l. Kerr J, Frankel HM: Inadequacy of blood drawn by ca rdi,ic puncture as a so urce for res piratory gas measurem ents in turtl es !Pscudc111_11s scrip/a). Comp Bi ochem Ph ysiol 41A:913, 1972 15. King AS, Pa yne DC: Norma l brea thing and the e ffects of posture in Gallus domcslirns. J Ph vs iol 174:340, 1964 16. Kru~1 H: lntraosseous fluid administration in sea turtles. /11 Proc Assoc Reptilian Amphibian Vet, Houston, TX, 1997, p 125 17. Lill ywhite HB: Circu latory adaptations of snakes to gravitv. Am Zoo! 27:81, 1987 18. Ludders JW, Mitche ll GS, Schaefer SL: Minimum anestheti c dose and cardiopulmonary dose response for halothane in chickens. Am J Vet Res 49:929, 1988 19. Ludders JW, Rode J, Mitchell GS: Isofluran e anesthesia in sa ndhill cranes (Grus ca11ade11sis): Minimal anesthetic concentration and cardiopulmon ary dose-response during spontaneous and controlled breathing. Anesth Anal g 68:511, 1989 20. Martin HD, Kolli as GV: Evaluation of water depri vation and fluid thera py in pigeons. J Zoo Wild! Med 20:173, 1989 21. Muir WW, deMorai s HSA: Acid-base balance: Traditional and modified approaches. /11 Thurmon JC, Tranq uilli WJ, Benson GJ (eds): Lumb & Jones' Veterinary Anesthesia, ed 3. Baltimore, Williams & Wilkins, 1996, p 558 22. Murray MJ: Cardiology and circulation. ill Made r DR (ed): Reptile Medicine and Surgery. Philadelphi a, WB Saunders, 1996, p 95 23. Phalen DN, Mitchell ME, Cava zos-Martinez ML: Eva lu ation of three hea t sources for their abi lity to maintain core bod y tempera ture in the anesthetized avian pati ent. J Avian Med Surg 10:174, 1996 24. Prezant RM, Jarchow JL: Lactated fluid use in reptiles: Is there a better solution? In Proc Assoc Reptilian Amphibian Ve t, Houston, 1997, p 83 25. Quasha AL, Eger EI, Tinker JH: Determination and app li cation of MAC. Anes thesiology 53:315, 1980 26. Rosenthal K, Mille r M: Cardiac disease. /11 Altman RB, Clubb SL, Dorrestein GM, et a l (eds): Avian Medicine and Surgery. Philadelphi a, WB Saunders, 1997, p 491 27. Schmidt-Nielsen K: Scaling: Wh y is anima l size so important? New York, Cambridge University Press, 1984 28. Schmitt PM, Gobe l T, Trautvetter E: Eva luati on of pu lse oximetry as a monitoring method in avian anesthesia. J Avian Med Surg 12:91, 1998 29. Seaman GC, Ludders JW, Hollis NE, et al: Effects of low and hi gh fractions of inspired oxygen on ventilation in du cks anesthetized with isofluran e. Am J Vet Res 55:395, 1994 30. Sedgwick C: All ometrically scaling the data base for vital sign assessment used in genera l anesthesia of zoological species. Proc Ann Meet Am Assoc Zoo Vet, 1991, p 360 31. Tremper KK, Barker SJ : Monitoring of oxyge n. in Lake CL (ed): Clinical Monitoring for Anesthesia and Critical Care, ed 2. Philadelphia, WB Saunders, 1994, p 196 32. Wang T, Smits AW, Burggren WW: Pulmonary function in reptiles. /11 Gans C, Gaunt AS (eds): Biology of the Reptilia, Vol 19. Morphology G, Visceral Organs. St. Louis, Society for the Study of Amphibians Reptiles, 1998, p 297 33. White FN: Circulation. In Gans C, Dawson WR (eds): Biology of th e Reptili a, Vol 5. Physiology A. New York, Aca demic Press, 1976, p 275 34. Wood SC, Lenfant CJM: Respira tion: mechani cs, contro l and gas exchange. In Gans C, Da wson WR: Biology of the Reptilia, Vol 5. Physiology A. New York, Academic Press, 1976, p 225

Address reprint requests lo Darryl Heard, BSc, BVMS, PhD, Dip!. ACZM College of Veterinary Medicine University of Florida Gainesv ille, FL 32610- 0126 e- mail: heardd @mail. ve tm ed .ufl.edu