John W. Ludders1 and David Sanchez-Migallon Guzman2 1 Emeritus Professor, Department of Clinical Sciences, College of Veterinary Medicine, Cornell University, Ithaca, New York, USA 2 Department of Medicine and Epidemiology, School of Veterinary Medicine, University of California‐Davis, Davis, California, USA The class Aves consists of 27 Orders, 168 Families, and approximately 10,000 species worldwide. Birds, regarded as the only clade of dinosaurs that survived the Cretaceous–Paleogene extinction event 66 million years ago [1], inhabit every continent on this planet and live in a wide range of environmental niches. For example, emperor penguins (Aptenodytes forsteri) live in the Antarctic, and one study recorded an individual ocean dive record of 540 m (1772 ft) lasting 18 min [2]. A Ruppell’s griffon (Gyps rueppelli), the highest‐flying bird, encountered a jet at 11,485 m (37,900 feet) over the west African country of Côte d’Ivoire. Bar‐headed geese (Anser indicus) routinely fly over the Himalayas, some as high as 7290 m (23,917 ft), when migrating between central and south Asia [1]. The burrowing owl (Athene cunicularia) nests underground while wandering albatrosses spend most of their life in soaring flight over the Southern Ocean. The weight and size of birds vary greatly. Bee hummingbirds (Mellisuga helenae) measure 5–6 cm (2–2.4 in) in length and weigh 1.6–2 g while the ostrich (Struthio camelus) has a height of 2.75 m (9 ft) and weighs up to 145 kg. Throughout time, humans have had a practical interest in birds as a source of food. In domesticating and selecting for desirable production characteristics, such as rapid weight gain or high egg production, a number of structural and functional changes have occurred in domesticated species, changes not seen in their wild relatives. For example, domesticated turkeys and chickens have smaller lung volumes and less gas exchange surface area compared to their wild counterparts. The tremendous diversity in form, function, and mode of life existing across avian orders and between wild and domesticated species, as well as variation that exists within any given species, pose challenges to the anesthetic management of birds. Indeed, the risk of anesthesia‐related death of birds is as high as 3.9% [3–5]. Challenges associated with anesthetizing birds can be lessened by considering and applying basic principles of avian anatomy, physiology, and pharmacology. The avian respiratory system consists of two separate and distinct functional components: one for ventilation (larynx, trachea, syrinx, bronchi, air sacs, thoracic skeleton, muscles of respiration), and one for gas exchange (parabronchial lung). These two components can be used to advantage when anesthetizing birds, especially when using inhalant anesthetics. The avian larynx is located at the base of the tongue and protrudes into the pharynx as a somewhat heart‐shaped mound [6]. Birds do not have an epiglottis, so when the tongue is pulled gently forward the larynx is easily visualized in most birds (Fig. 58.1). A notable exception is the flamingo with its ventroflexed beak and large fleshy tongue that make it difficult to visualize the larynx. The avian trachea, which consists of complete cartilaginous rings, conducts air from the nares and mouth to the bronchi while warming, moisturizing, and screening particulate matter from inspired gas [6]. From one avian class to another, there are tracheal anatomical differences that have significant implications for ventilation. For example, emu and ruddy ducks have an inflatable sac‐like diverticulum that opens from the trachea. In emu of both sexes, the sac arises from the ventral surface of the trachea approximately three‐quarters of the way down the neck where the tracheal rings are incomplete and form a slit‐like opening (Fig 58.2) [6]. This sac is responsible for the characteristic booming call of the emu. In male ruddy ducks, the sac opens in a depression on the dorsal wall of the trachea immediately caudal to the larynx, thus lying between the trachea and esophagus [6]. The sac may act as a sounding board for the bill‐drumming display of the males [6]. Some penguins and petrels possess a double trachea consisting of a median septum dividing part of the trachea into right and left channels [6]. In both groups of birds, the septum extends cranially from the bronchial bifurcation, but its length is variable; in the jackass penguin (Spheniscus demersus), it extends to within a centimeter of the larynx [7], whereas in the rockhopper penguin (Eudyptes spp.), the septum is only 5 mm in length [6]. Figure 58.1 Larynx of a chicken. Figure 58.2 Tracheal slit of an emu (Dromaius novaehollandiae). Source: Dr. Julie A. Smith, with permission. Other classes of birds have complex tracheal loops or coils in the caudal neck, within the keel, or within the thorax and keel (Fig. 58.3). Studies in cranes (Grus spp.) have demonstrated that tracheal coiling enables these birds to produce extremely loud calls using very low driving pressures [8]. Birds generally have relatively long necks, not to mention tracheal loops and coils, which affect tracheal deadspace, an important consideration during general anesthesia. The typical avian trachea is 2.7 times longer than that of comparably sized mammals, but it is 1.29 times wider, so tracheal resistance to gas flow is comparable in birds and mammals [6]. Tracheal deadspace volume in birds is about 4.5 times larger than that of comparably sized mammals, but the relatively low respiratory frequency (about one‐third that of mammals) and larger tidal volume of birds (about four times larger than in a comparably sized mammal) ensure that the effect of the larger tracheal deadspace volume is decreased [6]. The net effect is that avian tracheal minute ventilation is only about 1.5 to 1.9 times that of mammals [6]. The syrinx, the sound‐producing organ in birds, is located at the junction of the trachea and mainstem bronchi and the details of its structure vary among avian species. Its location and structure (a number of variably ossified cartilages, muscles, and vibrating soft structures) explain why gas flowing through the trachea, especially during positive‐pressure ventilation, can produce sound in an anesthetized, intubated bird. Mammals have 23 orders of bronchial branching leading to the gas exchange area of the lung (the alveoli), but birds have only three orders of branching before reaching gas exchange tissue [9]. The avian bronchial system consists of a primary bronchus (extrapulmonary and intrapulmonary), secondary bronchi, and tertiary bronchi (referred to as “parabronchi”). The parabronchi and their surrounding mantle of tissue (the peri‐parabronchial mantle) is where gas exchange occurs [6,9]. Figure 58.3 Different forms of tracheal loops: black swan (Cygnus atratus); whooper swan (Cygnus cygnus); white spoonbill (Platalea leucorodia); whooping crane (Grus americana); helmeted curassow (Crax Pauxi). Source: Adapted from McLelland [6]. The primary bronchus enters the lung ventrally and obliquely at the junction of the cranial and middle thirds of the lung, then passes dorsolaterally to the lung surface where it turns caudally in a dorsally curved direction until, at the caudal lung margin, it opens into the abdominal air sac [10]. The primary bronchi have low columnar, pseudostratified epithelium overlying a well‐developed internal circular smooth muscle layer, and longitudinally oriented smooth muscles [11] that can change the internal diameter of the primary bronchus. Acetylcholine, pilocarpine, and histamine contract bronchial smooth muscles; atropine blocks the effects of these drugs but has no effect when given alone [12]. A secondary bronchus is any bronchus arising from a primary bronchus; for a short distance, they have the same histologic structure as the primary bronchus, but subsequently develop simple squamous epithelium [13,14]. In most birds, the secondary bronchi are arranged into four groups: medioventral, mediodorsal, lateroventral, and laterodorsal secondary bronchi [15]. The medioventral secondary bronchi arise from the primary intrapulmonary bronchus close to where it enters the lung and occupy the ventral surface of the lung [16]. The mediodorsal, lateroventral, and laterodorsal secondary bronchi arise from the caudal curved portion of the primary intrapulmonary bronchus. Many medioventral and lateroventral secondary bronchi open through ostia into the cervical, clavicular, cranial thoracic, caudal thoracic, and abdominal air sacs. With a few notable exceptions, most birds have nine pulmonary air sacs that connect to the lungs: two cervical, an unpaired clavicular, two cranial and two caudal thoracic, and two abdominal air sacs. Some exceptions include storks, where each caudal thoracic air sac is divided in two, making 11 air sacs; domestic fowl, where the cervical air sacs are fused, making eight air sacs; songbirds, where the cranial thoracic air sacs are fused to the single median clavicular sac, making seven air sacs in all; and turkeys, where the caudal thoracic air sacs are absent and they have a fused cervicoclavicular air sac, making seven definitive air sacs [15]. In contrast to the pulmonary air sacs, the cervicocephalic air sacs are not connected to the lung and are divided into cephalic and cervical portions; they connect to caudal aspects of the infraorbital sinus. Extensive cervicocephalic air sac development has been noted in budgerigars, cockatiels, conures, Amazon parrots, macaws, and cockatoos. Air sacs are thin‐walled, vessel‐poor structures composed of simple squamous epithelium; they do not significantly contribute to gas exchange [17,18]. In chickens (Gallus gallus domesticus), adrenergic and cholinergic nerve plexuses have been described in the walls of air sacs, along with vasoactive intestinal peptide and fibers containing substance P, somatostatin, and enkephalin‐immunoreactive fibers [18]. To a varying extent, depending upon the species, diverticula from the pulmonary air sacs aerate the cervical vertebrae, some of the thoracic vertebrae, vertebral ribs, sternum, humerus, pelvis, and head and body of the femur [15]. The pulmonary air sacs functionally provide a flow of air to the relatively rigid avian lung which changes in volume by only 1.4% [19]. Based on their bronchial connections, air sacs are grouped into a cranial group consisting of the cervical, clavicular, and cranial thoracic air sacs, and a caudal group consisting of the caudal thoracic and abdominal air sacs [20]. The volume is distributed approximately equally between the cranial and caudal groups [21]. During ventilation, all air sacs are effectively ventilated, with the possible exception of the cervical air sacs; the ratio of ventilation to volume is similar for each air sac [21,22]. In birds, unlike in mammals, inspiration and expiration are active processes requiring muscular activity. Birds lack a muscular diaphragm, so pressures do not differ between the thoracic and abdominal cavities [11,15,16,23]. Inspiration and expiration occur through movement of the sternum by contraction of the cervical, thoracic, and abdominal muscles [16,23]. As the inspiratory muscles contract, the internal volume of the thoracoabdominal cavity increases (Fig. 58.4). Since air sacs are the only volume‐compliant structures in the body cavity [19], pressure within the air sacs becomes negative relative to ambient atmospheric pressure, and air flows from the atmosphere into the pulmonary system, specifically into the pulmonary air sacs and across the gas exchange surfaces of the lungs. During expiration, pressure within the air sacs becomes positive relative to ambient atmospheric pressure, and air flows from the air sacs and pulmonary system to the environment. A parabronchus (tertiary bronchus) and its mantle of surrounding tissue, consisting of air capillaries and blood capillaries, is the basic unit of gas exchange. Paleopulmonic parabronchi are long, narrow tubes that anastomose and connect the medioventral to the mediodorsal secondary bronchi; the neopulmonic parabronchi profusely interconnect mainly the laterodorsal and lateroventral secondary bronchi [19] (Fig. 58.5A and B). There is a network of smooth muscle surrounding the entrances to the parabronchi; electrical stimulation of the vagus nerve causes the smooth muscle to contract, thus narrowing the parabronchial openings [24]. The inner surfaces of the tubular parabronchi are pierced by numerous openings into chambers called “atria” that are separated from each other by inter‐atrial septa (Fig. 58.6A and B) covered by a thin epithelial layer with a core of densely packed bundles of smooth muscle framing the atrial openings [15]. Since the avian lung is richly innervated with vagal and sympathetic nerves, it is possible that afferent and efferent neural pathways exist for controlling pulmonary smooth muscle, thus varying airflow through the parabronchial lung in response to a variety of stimuli [20]. Figure 58.4 Changes in the position of the thoracic skeleton during breathing in a bird. The solid lines represent the thoracic position at the end of expiration while the dotted lines show the thoracic position at the end of inspiration. Source: Fedde MR. Respiration. In: Sturkie PD, ed. Avian Physiology, 4th edn. New York, NY: Springer‐Verlag, 1986; 191–220. Figure 58.5 Two views of secondary bronchi and parabronchi in the right lung of a goose. A. Medial view. B. Dorsal view. Source: Brackenbury, JH. Ventilation of the lung‐air sac system. In: Seller TJ, ed. Bird Respiration, Vol I. Boca Raton, FL: CRC Press, 1987; with permission. Arising from the abluminal floor of each atrium are funnel‐shaped ducts (infundibulae) that lead to air capillaries measuring 3 to 20 μm in diameter, and which are globular in shape, not tubular as previously alleged, and that interconnect via short, narrow passageways; the blood capillaries with which the air capillaries entwine consist of segments that are about as long as they are wide and form an anastomosing three‐dimensional network spatially arranged in the exchange tissue [19]. It is in this highly compact tissue of air and blood capillaries that gas exchange occurs [19] (Fig. 58.7A–C). The law of Laplace (P = γ/r; where P is opening pressure, γ is surface tension, and r is radius of tubule) applied to small‐diameter tubules, such as air capillaries, indicates that high surface tensions result and generate significant negative pressure across the blood–gas barrier that could lead to influx of fluid or collapse the tubules [25]. However, air and blood capillaries possess innate structural elements that preserve their anatomic and gas exchange integrity [26]. These elements form an interdependent, tightly coupled network of tension and compression – a tensegrity state – in the avian lung that gives the lungs their shape while strengthening the air and blood capillaries, thus preserving their function [19]. There are two types of parabronchial tissue: (1) paleopulmonic parabronchial tissue (paleopulmonic lung), found in all birds, consisting of parallel stacks of profusely anastomosing parabronchi; and (2) neopulmonic parabronchial tissue (neopulmonic lung), a meshwork of anastomosing parabronchi located in the caudolateral portion of the lung, the degree of development of which is species‐dependent (Fig. 58.8A–C). Penguins and emu have only paleopulmonic parabronchi. Pigeons, ducks, and cranes have both paleopulmonic and neopulmonic parabronchi with the neopulmonic parabronchi accounting for 10–12% of the total lung volume. In fowl‐like and song birds, the neopulmonic parabronchi are more developed and may account for 20–25% of total lung volume. Paleopulmonic and neopulmonic parabronchi are histologically indistinguishable from each other [19]. Compared to mammals, specific total lung volume in birds is about 27% smaller, but specific surface area of the blood–gas (tissue) barrier is ~15% greater; the ratio of the tissue surface area to the volume of the exchange tissue is 170–305% greater [27]. The harmonic mean thickness of the tissue barrier in birds is 56–67% less (less resistance to gas diffusion) and the pulmonary capillary blood volume is 22% greater. With the exception of specific total lung volume, these morphometric parameters favor the gas exchange capacity of birds [27]. The specific volume (respiratory gas volume per unit body mass) of the avian pulmonary system is between 100 and 200 mL/kg, but the volume of gas in the parabronchi and air capillaries where gas exchange occurs accounts for only 10% of the total specific volume [21]. By comparison, a dog’s specific volume is 45 mL/kg, and the pulmonary gas volume in the mammalian lung is 96% of the total specific volume. Because the ratio of residual gas volume (i.e., gas in the lungs) to tidal volume is so much smaller in birds than in mammals, it has been suggested that cyclic changes in the direction of parabronchial gas flow (i.e., tidal flow) could produce significant and intolerable cyclic changes in gas exchange analogous to breath holding [28]. The unidirectional flow of gas within the paleopulmonic lung solves this problem. Figure 58.6 Three‐dimensional drawings of a parabronchus and an atrium. A. Sagittal section of a parabronchus. On the left side are atria with infundibula departing from them and the three‐dimensional air capillary meshwork arising from the infundibula. On the right side within the interparabronchial septa are the arterioles (dense stippling) from which the capillaries originate and run radially to the lumen. The infundibula lie between the capillaries which are surrounded by a well‐developed three‐dimensional air capillary network. B. Atrium and infundibulum. At the left, two of the circular smooth muscle bundles surrounding the lumen of the parabronchus are shown in cross‐section. The atria are separated by septa running horizontally and vertically. Originating from each atrium a few infundibula pass perpendicularly into the parabronchial mantle. At the right, an infundibulum is shown in longitudinal section with air capillaries arising from it at all levels. The air capillaries cross‐link and interlace, making up a three‐dimensional meshwork around the blood capillaries. The very thin epithelium of the air capillaries and its surfactant film are shown as a single dark line. Source: Duncker [11], reproduced with permission from Elsevier. During a respiratory cycle, the direction of gas flow in the paleopulmonic parabronchi is caudocranial, continuous, and unidirectional, but in the neopulmonic parabronchi it is tidal (i.e., the direction of flow changes during the respiratory cycle [19] (Fig. 58.9A and B). The exact course of inspired air through the avian respiratory system has been debated for years, but now a general consensus exists on its route during respiratory cycles [19]. It takes two inspiratory and two expiratory cycles for inspired air to pass through the lung–air sac system [19]. During the first inspiratory cycle, the air flows through the trachea and the extrapulmonary and intrapulmonary primary bronchi (mesobronchus) and then the caudal air sacs; in the next breathing cycle (the first expiratory cycle), the air is expelled from the caudal air sacs and directed into the parabronchial system of the lung via the mediodorsal secondary bronchi; during the next breathing cycle, (the second inspiratory cycle), the air in the parabronchial system moves to the cranial air sacs; and in the subsequent breathing cycle (the second expiratory cycle), the air in the cranial air sacs is expelled to the outside through the extrapulmonary primary bronchus and the trachea. The unidirectional flow of gas through the paleopulmonic parabronchi is probably due to aerodynamic valves, not mechanical valves [19]. Although poorly understood, the mechanisms involved probably include the orientation of secondary bronchial and air sac orifices to the direction of gas flow, elastic pressure differences between the cranial and caudal group of air sacs, and gas convective inertial forces [18]. The movement of gas within the parabronchi and outwards into the atria and infundibulae is by convective flow and then by diffusion into the air capillaries [29,30]. Blood flows from the periphery via the interparabronchial artery and arterioles, ultimately flowing into the blood capillaries where it meets the outward‐moving air capillaries. The cross‐current model and its multicapillary serial arterialization system best describe gas exchange in the avian lung. The multicapillary serial arterialization system increases the duration over which the respiratory media (air and blood) are exposed to each other [18]. Thus, the quantity of oxygen and carbon dioxide in the oxygenated blood that ultimately returns to the heart via the pulmonary vein is derived from an aggregate effect of very small quantities exchanged at the infinitely many points where the air and blood capillaries contact [19]. Reversing the direction of air or blood flow in a multicapillary serial arterialization system only changes the sequence in which the blood is arterialized in the blood capillaries; the overall amount of oxygen that is exchanged does not change [19]. Within the avian lung is a counter‐current system created by the centripetal (inward) flow of deoxygenated blood and the centrifugal (radial, outward) flow of air from the parabronchial lumen [18,19]. However, since the air and blood capillaries entwine very closely, at any one point they contact over very short distances that may fall far short of the critical lengths needed for efficient gas exchange in a “conventional counter‐current system.” (Fig. 58.10). The significance of this counter‐current system for gas exchange under normal physiologic conditions is uncertain; under extreme conditions (flight at altitude), such an arrangement may contribute to gas exchange. The efficiency of the avian lung can be put into perspective by considering what happens to the partial pressures of carbon dioxide (CO2) and oxygen (O2) both in respired gas flowing through the lung and in blood perfusing the lung. As gas flows along a parabronchus, it receives CO2 and gives off O2 such that gas at the inflow end of the parabronchus has the lowest partial pressure of CO2, while gas at the outflow end has the highest partial pressure of CO2; the reverse is true of O2. The overall result, under hypoxic conditions and during exercise, is that the partial pressure of CO2 in end‐parabronchial gas (PECO2) can exceed the partial pressure of CO2 in arterial blood (PaCO2), and the partial pressure of O2 in end‐parabronchial gas (PEO2) can be lower than the partial pressure of O2 in arterial blood (PaO2) [31–33]. This potential overlap of blood and gas partial pressure ranges for both CO2 and O2 demonstrates the high gas exchange efficiency of the avian lung [31,32]. Thus, there is no equivalent of alveolar gas because parabronchial gas continuously changes in composition as it flows along the length of the parabronchus [34]. Practically speaking, this means the avian lung can extract 40 cm3 of oxygen from each liter of inspired air, whereas the mammalian lung can extract 30 cm3 of oxygen [19]. Figure 58.7 Three‐dimensional reconstruction showing the intimate intertwining of blood and air capillaries in the paleopulmonic lung of the ostrich (Struthio camelus). A. Combined three‐dimensional reconstructions of the blood capillary (BC, red) and air capillary (AC, cyan) systems. B. Three‐dimensional reconstruction of the blood capillaries. C. Three‐dimensional reconstruction of the air capillaries. Scale bars = 20 µm. Source: Maina JN, Woodward JD. Three‐dimensional serial section computer reconstruction of the arrangement of the structural components of the parabronchus of the ostrich, Struthio camelus lung. Anat Rec (Hoboken) 2009; 292(11): 1685–1698, with permission. Figure 58.8 Diagram of paleopulmonic and neopulmonic lungs. A. The paleopulmonic lung found in penguins and emus. B. The paleopulmonic and neopulmonic lung found in storks, ducks, and geese. C. The paleopulmonic and the more highly developed neopulmonic lung found in chickens, sparrows, and other songbirds. Source: Fedde [20], with permission of Elsevier. Figure 58.9 Schematic representation of gas flow through the avian pulmonary system during inspiration and expiration. The neopulmonic lung has been removed for clarity. A. Inspiration. B. Expiration. Figure 58.10 Diagram depicting the three intertwined systems involved in gas exchange in the avian parabronchus: cross‐current, counter‐current, and multicapillary serial arterialization. The cross‐current system consists of air flowing in the parabronchial lumen at right angles to that of deoxygenated blood flowing from the periphery of the parabronchus (small pink arrows). The counter‐current system is formed by the centripetal flow of deoxygenated blood in the blood capillaries relative to the centrifugal diffusion of air in the air capillaries (black dashed arrows). The multicapillary serial arterialization system is formed by multiple successive contacts between the air and blood capillaries in the gas exchange tissue (dashed yellow circles). For clarity, the atria and infundibula are not shown. The shading of blood from blue to red in the intraparabronchial vein reflects the increasing oxygenation of blood as it passes through the parabronchial gas exchange tissue. Source: Maina [19], with permission. Birds have many of the same physiologic components for respiratory control as mammals, such as a central respiratory pattern generator, central chemoreceptors sensitive to PCO2, and many similar peripheral chemoreceptors [35]. Birds have a unique group of peripheral receptors located in the lung called “intrapulmonary chemoreceptors” (IPCs) that are vagal respiratory afferents inhibited by high lung PCO2 and excited by low lung PCO2, thus providing phasic feedback for the control of breathing, specifically rate and depth of breathing [35–38]. They are not mechanoreceptors and they are insensitive to hypoxia [39,40]. However, IPCs are not the sole receptors stimulated by inhaled gas containing low partial pressures of CO2; arterial and central chemoreceptors are also stimulated [41]. There may be species differences in CO2 responsiveness depending upon the ecologic niche a given species occupies. The CO2 responsiveness of IPCs in chickens, ducks, emus, and pigeons is greater than that of burrowing owls that live underground where the CO2 concentration is higher than that of above‐ground‐dwelling birds [42,43]. The avian heart is a four‐chambered muscular pump that separates venous blood from arterial blood. Birds have larger hearts, larger stroke volumes, lower heart rates, and higher cardiac output than mammals of comparable body mass [44]. Birds also have higher blood pressure than mammals [44,45]. The atria and ventricles are innervated by sympathetic and parasympathetic nerves [46], and norepinephrine and epinephrine are the principal sympathetic neurotransmitters while acetylcholine is the principal parasympathetic neurotransmitter. The conduction system of the avian heart consists of the sinoatrial node, the atrioventricular node and its branches, and Purkinje fibers [46]. Two groups of animals can be identified by the depth and degree to which Purkinje fibers ramify within the ventricular myocardium, and the pattern of ramification is classified as type 1 or 2 (or category A or B) [47]. The pattern of Purkinje fiber distribution within the ventricular myocardium is responsible for QRS morphology. In birds, Purkinje fibers completely penetrate the ventricular myocardium from endocardium to epicardium and the pattern of ventricular activation is described as type 2b, a pattern that may facilitate synchronous beating at high heart rates [48]. The avian kidney receives venous blood from the legs via the renal portal circulation, and arterial blood via the arterial circulation [49,50]. The flow of afferent venous blood, unlike in other non‐mammalian vertebrates, is not obligatory; venous blood can either perfuse the renal parenchyma or bypass it and enter the central circulation. A unique valve‐like structure, the shape of which varies from species to species, is located within the external iliac vein where the efferent renal vein joins it [49]. The valve contains smooth muscle innervated by adrenergic and cholinergic nerves. Epinephrine causes the valve to relax (open) and venous blood from the legs enters the central circulation, whereas acetylcholine causes it to contract (close) and venous blood perfuses the kidney [49,51]. The control of renal valve activity is complex and its effects on drug uptake are not fully understood. Proper physical restraint of a bird is a crucial aspect of anesthetic management. In general, a bird is restrained so that its wings and legs are controlled and not allowed to flap or kick about. For long‐necked birds such as herons and cranes, the neck must be gently controlled so that the bird does not suffer head, eye, or neck trauma; restraining the bird’s head and neck also protects the handler. Improper restraint can cause a variety of problems for a bird, including physical trauma (wing or leg fractures), or physiological stress with increased levels of catecholamines, especially epinephrine, which may predispose to cardiovascular instability [46,52]. Physical restraint can cause significant changes in heart and respiratory rates, and body temperature. Because birds cannot dissipate heat through the skin, they can become stressed and easily overheated with prolonged restraint. For example, during 15 min of restraint of Amazon parrots (Amazona spp.), the average cloacal temperature increased significantly by 4.3 °F while mean respiratory rates increased significantly from 129 to 252 breaths/min [53]. This emphasizes the importance of limiting restraint time and watching for tachypnea, even in healthy birds, so as to avoid potentially life‐threatening increases in body temperature [53]. However, in restrained birds, the behavioral indicators of stress – increased respiratory and heart rates – may vary by species. In barred owls (Strix varia), heart and respiratory rates decreased significantly during 15 min of restraint while body temperature significantly increased [54]. In a study of red‐tailed hawks (Buteo jamaicensis) manually restrained for 15 min while hooded or not hooded, heart and respiratory rates decreased significantly while cloacal temperature increased significantly regardless of hooding; when hooded, the hawks had significantly lower heart and respiratory rates than when not hooded [55]. Thus, hooding amplified the decrease in heart rate and respiratory rate compared to nonhooding but had no effect on stress‐induced hyperthermia [55]. It would seem that barred owls and red‐tailed hawks cope differently with restraint‐associated stress when compared to psittacine species [54]. In a study of geese conditioned over several weeks to 5 min of physical restraint, the humoral indices of stress – catecholamines, corticosterone, and lactate – significantly increased within 2 min after the start of physical restraint despite weeks of conditioning to restraint and the fact that the birds appeared outwardly calm [56]. Thus, the absence of stress cannot be deduced from behavioral observations only [56]. Anesthesia and sedation may ameliorate the humoral stress response. Awake and manually restrained Hispaniolan Amazon parrots had significantly higher serum corticosterone levels (33.7 ± 2.83 ng/mL) than when anesthetized with isoflurane (19.8 ± 1.97 ng/mL) [57]. In manually restrained Hispaniolan Amazon parrots (Amazona ventralis) administered midazolam (2 mg/kg) intranasally, the mean rate of cloacal temperature increase was significantly slower and remained significantly lower in birds that received midazolam compared with controls [58]. Understanding the physical characteristics and defense mechanisms birds possess is crucial for effective restraint that protects both the bird and handler. Each avian species has its own unique and effective means of defense and attack. Birds of prey will use their talons and inflict severe physical trauma on a handler or assistant, and the risk of infection from such wounds is quite real. Although most birds of prey do not bite, they can and will use their talons and beak to great effect. Psittacines have very strong beaks that can cause severe soft tissue injury. Cranes and herons use their long, pointed beaks in a spearing manner, and they tend to strike at eyes, a reason for wearing eye protection when working with these birds. Cranes and ratites, such as emus and ostriches, also will jump and strike forward with their legs and clawed feet; an adult ratite can produce a very powerful and dangerous strike‐kick. Every bird should be given a thorough physical examination prior to anesthesia. A number of excellent texts describe in detail the techniques for physical examination and what to look for in specific avian species [59–61]. In general, quiet observation of a bird in its cage will provide a great deal of information. A bird’s awareness of and attention to its surrounding environment, body form, and posture, feather condition, and respiratory rate provide clues to its physical condition. Birds should be removed from their cage and examined, with particular attention given to the respiratory and cardiovascular systems. For small species, a stethoscope with a pediatric head should be used for cardiopulmonary examinations. For those species with a crop, it should be palpated in the thoracic inlet to make sure it is empty. The pectoral muscles over the keel should be assessed as they are a good indicator of body condition. A reasonable approach for healthy birds is to withhold food long enough for the upper gastrointestinal tract to empty, usually overnight in large birds and 4 to 6 h in smaller birds [62]. For raptorial species, a 24 h fast is recommended. In an emergency, a bird with a full crop should be held upright during induction with a finger positioned just below the mandible so as to block the esophagus [62]. Once the bird is anesthetized, the crop can be emptied by placing a finger covered with gauze over the choanal slits to prevent food from entering the nasal cavity and then milking the food contents out of the crop and esophagus [62]. At the end of anesthesia, the oral cavity should be checked for and cleaned of food material to prevent aspiration. Sedation offers numerous benefits in avian medicine, including attenuation of stress responses caused by manual restraint and ease of diagnostic sample collection and imaging, and it may decrease the risk of morbidity and mortality in critically ill patients [63,64]. There are a number of routes by which sedative and anesthetic drugs can be administered to birds. Subcutaneous injection sites include the area over the back between the wings, the wing web, and the skin fold in the inguinal region. Intramuscular injections can be made in the pectoral and thigh muscles. In birds, drugs injected intramuscularly in the hind limb or caudal body might be carried in the blood through the renal portal system and perfuse the kidneys such that some portion of the injected drug may be excreted by the kidney without reaching the general systemic circulation, or it may bypass the kidneys and undergo first‐pass hepatic metabolism [65]. In great horned owls administered butorphanol either via the jugular vein or the medial metatarsal vein, the AUC0→∞ after medial metatarsal vein administration was significantly less than the AUC0→∞ after jugular vein administration of the drug, results indicating a possible renal portal first‐pass effect, but no other kinetic parameters were significantly different [66]. In a pharmacokinetic study of florfenicol, an antibiotic, there was no evidence of an influence of the renal portal system on its kinetic parameters when injected into the leg muscles [65]. For intravenous injections as well as for catheterization, the ulnar vein, dorsal metatarsal vein, and jugular vein can be used; the right jugular vein is larger and easier to visualize than the left jugular vein. The ulnar vein can be catheterized in birds as small as cockatiels. An alternative non‐invasive route that is safe and rapidly induces sedation in birds (first effect within 3–5 min) is the intranasal route [64]. When using this route, tissue trauma associated with IM injections and subsequent elevations of muscle enzymes in clinical chemistry tests are avoided, and it is not painful. Its limitations are that there may be incomplete drug delivery to a bird due to sneezing, physiologically narrowed nostrils, such as in cockatoos, and upper airway disease, such as blocked or stenotic nostrils [64]. Intranasal delivery of drugs is made with either an insulin syringe (0.5 mL) or tuberculin syringe (1.0 mL) [64]. The intranasal route (IN) has been used to administer a variety of sedative and analgesic drugs to facilitate minor therapeutic procedures in a variety of birds. Xylazine, diazepam, and midazolam have been administered IN to budgerigars (Melopsittacus undulatus) and finches [67,68], and juvenile ostriches (Struthio camelus) weighing 50–61 kg and aged 4–5 months [69]. Midazolam, detomidine, or combinations of midazolam, xylazine, and ketamine have been administered IN to ring‐necked parakeets and successfully reversed with flumazenil, yohimbine, or atipamezole [70]. Intranasal midazolam and diazepam produced rapid and effective sedation in canaries, and xylazine and detomidine produced sedation but not sustained recumbency; specific antagonists (flumazenil and yohimbine) effectively reversed sedation [71]. In pigeons, a combination of midazolam and dexmedetomidine administered IN produced an effective degree of immobilization lasting 20 to 30 min, which was partially antagonized with atipamezole 10 min after its application [72]. In wild blue‐and‐yellow macaws (Ara ararauna), midazolam (2 mg/kg) divided into two equal volumes, was administered IN over 3–5 s via a 24 gauge catheter inserted approximately 2–5 mm into each nostril; sedation occurred in 77% of the macaws and lasted on average 20 min without adverse side effects [73]. In a project involving implantation of intracoelomic satellite transmitters in female surf scoters (Melanitta perspicillata), 26 were administered midazolam (4.6–5.9 mg/kg) intranasally, and the same volume of saline (1 mL) was given to another 26 adult female surf scoters [74]. The odds of presumed death in the saline group were 5.3 times higher than in the midazolam‐treated group; the presumed mortality at 30 days for the midazolam group (23%) was lower than for the saline group (61%) [74]. There is a large number of avian species, each one of which has species‐specific pharmacokinetic and pharmacodynamic profiles for each injectable drug [52]. Recognizing this reality, the following discussion of anesthetic and analgesic drugs will highlight a few key pharmacological realties pertaining to birds and focus primarily on drugs for which there are pharmacokinetic and pharmacodynamic data. The rational use of drugs is based primarily on our understanding of pharmacokinetic behavior coupled with information on pharmacodynamic activity, an approach that advances avian anesthesia and analgesia [75]. The reader is referred elsewhere for a more comprehensive review of anesthetic and analgesic drugs used in birds [76]. There are many reasons for using injectable drugs for anesthesia and analgesia in birds, including low cost, ease of use, rapidity of anesthetic inductions, no need for expensive equipment to deliver or maintain anesthesia, minimal if any anesthetic pollution of the work environment, and possible reduction of inhalant anesthetic requirements when used in combination. However, injectables also have inherent disadvantages including: (1) difficulty in delivering a safe volume to small birds, (2) ease in overdosing by any route, (3) difficulty in maintaining surgical anesthesia without severe cardiopulmonary depression, (4) the potential for prolonged, violent recoveries, and (5) significant variation in dose and response among and within species [52,77]. Allometric scaling has been used to determine doses of drugs used in birds [78–81]. Indeed, in the absence of pharmacokinetic data for a drug, general principles of allometric scaling may serve as a rational basis for understanding how body mass or metabolic rate affects drug doses, but there are limitations in its application. Allometric equations, with some exceptions, are based on the assumption that all birds are pharmacologically similar, and only distinguish between two large groups of birds – passerines and non‐passerines – a distinction that ignores the many structural and functional differences in the kidneys of birds within these two groups [51]. Drugs used in birds are formulated primarily for mammals. Differences in anatomy and physiology of the two classes of animals make it difficult to achieve pharmacokinetic equivalency between birds and mammals using allometric equations that do not take these differences into consideration [51,82]. Using allometric scaling, it may be possible to derive some pharmacokinetic parameters for a drug, but not all necessary parameters that would allow for safe administration of a drug [83,84]. Allometric scaling is not useful for extrapolating doses of non‐steroidal anti‐inflammatory drugs (NSAIDs) between avian species [85]. The pharmacologic reality is that for safe use of drugs in birds, we need pharmacokinetic and pharmacodynamic data. Early pharmacodynamic studies identified significant differences in response among avian species administered the same drug [52,86,87]. For example, the commercially available form of ketamine, which consists of a racemic mixture of S(+)‐ and R(−)‐enantiomers, produces poor‐quality chemical restraint and anesthesia in great horned and snowy owls [87]. When great horned owls receive only the S(+)‐enantiomer of ketamine, anesthesia induction is smoother and there are fewer cardiac arrhythmias, whereas the R(−)‐enantiomer is associated with inadequate muscle relaxation, cardiac arrhythmias, and excitement during recovery [87]. It is unknown whether these differences are due to differing metabolic pathways among birds, production of pharmacologically active metabolites, or differences in types of receptors or receptor sensitivity. Recognizing and assessing pain in birds can be difficult because it relies on an understanding of species and individual normal behaviors that could be masked in the presence of an observer. Birds tend to respond to noxious stimuli with a fight‐or‐flight response (i.e., escape reactions, vocalization, and excessive movement), and/or conservation–withdrawal responses (i.e., no escape attempts or minimal vocalization and immobility). Pain can change a bird’s social interactions, such as perching away from the flock, reduction in social grooming, or interactions with the owner. Grooming activity may decrease when a bird is painful, but conversely over‐grooming and feather‐destructive behaviors have been associated with chronic pain, which may include neuropathic pain [88]. These behavioral signs coupled with studies involving psittacines, raptors, domestic fowl, and pigeons, and using a variety of analgesiometric techniques and pharmacokinetic studies, have been used to determine the plasma concentrations of drugs that produce analgesia in birds (for examples, see [89–94]). Few studies have been conducted regarding the distribution, quantity, structure, and function of each opioid receptor type in birds. In an earlier study in pigeons, the regional distribution of μ (mu), κ (kappa), and δ (delta) opioid receptors in the forebrain and midbrain were similar to mammals, but the κ‐ and δ‐opioid receptors were more prominent in the pigeon forebrain and midbrain than μ‐opioid receptors and 76% of opioid receptors in the forebrain were determined to be κ‐type [95]. These findings, together with earlier studies in opioids in birds, led to a paradigm in which κ‐opioid receptor agonists such as butorphanol were considered the drugs of choice for pain management in birds. More recent studies of opioid receptors in chicks [96], pigeons, and cockatiels [97] challenge this paradigm and suggest that μ‐opioid receptor agonist drugs might actually be better choices for some avian species when treating pain. The results of recent studies are described in the following paragraphs. Interestingly, when looking at the structure of opioid receptors (μ, κ, and δ) in the peregrine falcon (Falco peregrinus), snowy owl (Bubo scandiacus), and blue‐fronted Amazon parrot (Amazona aestiva), nucleotide homologies with humans ranging from 77–82% were found, which might account for some of the differences seen in the efficacy of these drugs [98]. Most importantly, the large variability in pharmacokinetics and pharmacodynamics between species, and in some cases significant age‐ and sex‐dependent variability, has clearly shown that making pharmacologic generalizations across the class Aves, and extrapolating drug doses from one avian species to another must be exercised cautiously [75]. Butorphanol has been studied in gray parrots and Timneh parrots (Psittacus erithacus and Psittacus timneh) [89], domestic pigeons [99], red‐tailed hawks (Buteo jamaicensis) and great horned owls (Bubo virginianus) [66], Hispaniolan Amazon parrots (Amazona ventralis) [90,100–104], broiler chickens (Gallus gallus domesticus) [105,106], Indian peafowl (Pavo cristatus) [107], sulphur‐crested cockatoos and yellow‐crested cockatoos (Cacatua galerita, Cacatua sulphurea cintrinocristata and Cacatua sulphurea sulphurea) [108], green‐cheeked conures (Pyrrhura molinae) [101], American kestrels (Falco sparverius) [109], and orange‐winged Amazon parrots (Amazona amazonica) [104]. Most of these studies have evaluated the pharmacokinetics of the standard butorphanol tartrate formulation and some sustained‐release formulations, but only a few have evaluated the potential analgesic effects. The pharmacokinetic studies have shown that butorphanol has excellent parenteral and poor oral bioavailability (less than 10%), is quickly absorbed following SC and IM injection, and has a very short half‐life (e.g., Hispaniolan Amazon parrots 0.51 h, red‐tailed hawks 0.93 h, great horned owls 1.78 h, American kestrels 1.48 h, and chickens 1.19 h), which makes frequent intermittent administration (every 2–3 h) or administration by constant rate infusion (CRI) necessary. Pharmacodynamic studies have shown that relatively high doses are required in psittacines (e.g., 1 mg/kg in gray parrots when assessed using electrical antinociception and 5 mg/kg in Amazon parrots when using thermal antinociception) and are associated with a very short duration of action (between 20 min and 1.5 h) and a relatively small effect; however, studies are lacking to evaluate the effect of different doses. A study involving gray parrots, white cockatoos (Cacatua alba), and blue‐fronted Amazon parrots (Amazona aestiva aestiva) and using the minimum anesthetic concentration (MAC)‐sparing technique to determine the analgesic properties of butorphanol (1 mg/kg IM) resulted in a reduction of isoflurane MAC in cockatoos and gray parrots, but not in the blue‐fronted Amazon parrots indicating species variability to either the dose or the drug itself [110]. In white cockatoos, butorphanol significantly reduced isoflurane MAC from 1.44 ± 0.07% to 1.08 ± 0.05% [108,110]. Based upon these studies, doses of 1–5 mg/kg have been recommended in psittacines [111]. In raptors, results of the only pharmacodynamic study of butorphanol have been very different from those in psittacines. In American kestrels, doses of 1, 3, and 6 mg/kg IM had no thermal antinociceptive effects and males showed hyperalgesia at the higher dose [109]. These results discourage the use of butorphanol for analgesia in American kestrels and other birds of the Falco genus; it is unknown if these findings can be extrapolated to other raptors. Lame broiler chickens receiving butorphanol 2 mg/kg IV finished an obstacle course faster than the control group, and together with the latency‐to‐lie results reported in the same study, this suggests that butorphanol may be analgesic for up to 2 h [106]. The butorphanol sustained‐release formulations used with osmotic pumps, while promising, unfortunately, are not readily available to clinicians and are not frequently used. Nalbuphine has been studied in Hispaniolan Amazon parrots with standard and sustained‐release formulations. The pharmacokinetic studies have shown that the standard formulation of nalbuphine hydrochloride has a similar profile as butorphanol, with a slightly shorter half‐life. The pharmacodynamic studies using thermal antinociception resulted in significant results at 12.5 mg/kg for up to 3 h, while higher doses of 25 and 50 mg/kg were not different from control [92,112]. A sustained‐release formulation of nalbuphine had a longer half‐life and duration of effect (up to 12 h) [113], but it is not readily available at this time. Hydromorphone has been studied in American kestrels, cockatiels, and orange‐winged Amazon parrots. The pharmacokinetic studies demonstrated that hydromorphone has good parenteral (IM) bioavailability, is quickly absorbed, and has a longer half‐life (e.g., 1.26 h in American kestrels [114], 0.99 h in cockatiels [115], and 1.74 h in orange‐winged Amazon parrots [116]) than butorphanol in these species, indicating that frequent administration (approximately every 3–6 h) would be required. The analgesiometry studies have shown that relatively high doses are required in psittacines when compared to raptors. In raptors, the studies in American kestrels have shown that hydromorphone at 0.1, 0.3, and 0.6 mg/kg IM had a dose‐dependent effect using the thermal antinociception model [117]. In psittacines, in contrast, studies in cockatiels using the same antinociceptive dose as in kestrels showed no significant effect [115]. In orange‐winged Amazon parrots, higher doses of hydromorphone at 1 and 2 mg/kg showed thermal antinociception for 3–6 h, respectively [118]. Agitation, nausea‐like behavior (including vomiting in one bird), ataxia, and pupillary constriction were observed following administration of the 1 and 2 mg/kg hydromorphone doses which suggest that lower doses be used in this species. Morphine has been evaluated in multiple studies involving chickens (Gallus gallus domesticus). The only pharmacokinetic study has followed IV administration of 2 mg/kg IV and resulted in a half‐life of 68 min, with a large volume of distribution and rapid clearance [119]. Pharmacodynamic results from earlier studies have been conflicting and difficult to interpret likely due to the high doses evaluated, but more recent studies suggest that morphine might have analgesic and MAC‐sparing effects in avian species. In an early study, 200 mg/kg of morphine was injected to suppress nociception in chicks using the toe pinch test [120], but in a later study using noxious electrical stimulation, morphine produced analgesia in chicks at 30 mg/kg [121]. Further investigations using noxious thermal stimulation reported strain‐dependent analgesic effects of morphine, requiring doses of 15, 30, and 100 mg/kg for two different White Leghorn lines and a cross of Rhode Island Red × Light Sussex, respectively [122]; analgesic and hyperalgesic effects were reported with a 30 mg/kg dose in the Rhode Island Red cross, and White Leghorn and Cal White strains, respectively [123]. In a MAC reduction study in chickens, three doses of morphine injected IV caused a dose‐dependent decrease in isoflurane MAC in all birds. The baseline isoflurane MAC of 1.24 ± 0.05% was reduced 15.1 ± 2.7%, 39.7 ± 3.1%, and 52.4 ± 4.0% at doses of 0.1, 1.0, and 3.0 mg/kg, respectively [124]. In a recent study evaluating the analgesic efficacy of morphine, lame broiler chickens underwent an obstacle course and latency‐to‐lie test before and at 30 min and 2 h after injection of 2 mg/kg IV of morphine to assess their walking and standing abilities [106]. Morphine treatment caused sedation even in the sound chickens and resulted in an increased time to finish the obstacle course as the chicks tended to sit down rather than walk thereby confounding assessment of its analgesic effects [106]. Fentanyl has been evaluated in white cockatoos [125], red‐tailed hawks [75,126], helmeted guineafowl (Numida meleagris) [127], chickens [128,129], and Hispaniolan Amazon parrots [75,130]. The pharmacokinetic studies have shown significant differences in the half‐life, ranging from 30–90 min, a fact that discourages extrapolating drug doses between avian species. In chickens, 25 μg/h transdermal fentanyl patches have a large variability in plasma concentrations, but all chickens reached the human target plasma concentrations of 0.2–1.2 ng/mL within 2–4 h and maintained concentrations above that target for 72 h with a rapid decrease in plasma concentrations following removal of the patch [129]. The transdermal veterinary formulation evaluated in helmeted guineafowl was discontinued [127]. Pharmacodynamic studies have shown that significantly higher fentanyl doses or rates of infusion are needed to achieve desired analgesic or MAC‐sparing effects in psittacines as compared to other species. However, adverse effects occur at these higher doses or rates, such as agitation in awake white cockatoos receiving 0.2 mg/kg IM [125], or dose‐dependent decreases in heart rate and blood pressure in anesthetized Hispaniolan Amazon parrots [130]. In raptors, the findings have been very different, and the rates required to achieve the same MAC‐sparing effects have been 17 times lower than in psittacines, and without any significant negative effects on heart rate or blood pressure. Rates of 10–30 μg/kg/h IV decreased isoflurane MAC in a dose‐dependent manner by 31–55% [126]. Methadone has been studied in orange‐winged Amazon parrots [118] and chickens [131,132]. The pharmacokinetic studies have shown a longer half‐life than other opioid drugs but with differences between species (e.g., orange‐winged Amazon parrots 2.3 h, chickens under anesthesia 3 h). The pharmacodynamic study assessing thermal antinociception suggests that relatively high doses are required in orange‐winged Amazon parrots with an IM dose of 6 mg/kg, but not 1 or 3 mg/kg, resulting in thermal antinociception. Agitation, ataxia, and nausea‐like behavior were observed following administration, and consideration of lower doses for clinical use in this species is warranted. In chickens, isoflurane MAC‐sparing effects were documented after IM administration of 6 mg/kg, but not 3 mg/kg, with reductions of 29%, 27%, and 10% noted at 15, 30, and 45 min after methadone administration, respectively [132]. Buprenorphine has been studied in gray parrots [89,133], domestic pigeons [99], American kestrels [134–137], red‐tailed hawks [138], cockatiels [94], and orange‐winged Amazon parrots [Guzman, unpublished data]. The pharmacokinetic studies have been performed with standard, sustained‐released, and concentrated formulations. Those studies have shown major differences in the half‐life between species (e.g., 1.5 h in American kestrels [135] and 6.23 h in red‐tailed hawks [138]). In red‐tailed hawks, 0.3 mg/kg and 1.8 mg/kg doses using concentrated buprenorphine resulted in plasma concentrations being maintained above 1 ng/mL for at least 24 and 48 h, respectively [138]. Results of pharmacodynamic studies in psittacines have been discouraging because of the lack of significant effect with electrical and thermal antinociception in the species and doses evaluated (e.g., 0.1 mg/kg IM in gray parrots [89] and 0.6, 1.2, and 1.8 mg/kg IM in cockatiels [94]). Recent unpublished studies have shown a small but significant effect in orange‐winged Amazon parrots following 2 mg/kg IM, but not at 1 and 0.1 mg/kg doses [Guzman, unpublished data]. In contrast, pharmacodynamic studies in raptors have shown a longer duration of action than other opioids (e.g., up to 6 to 9 h in American kestrels following 0.6 mg/kg IM [134]) with mild sedation. High doses have been shown to be well tolerated and provide longer analgesic effects with moderate sedation. Sustained‐release formulations are commercially available (e.g., Buprenorphine SR‐LABTM) and have also been shown to be well tolerated and associated with prolonged effects (e.g., 24 h following 1.8 mg/kg SC in American kestrels), thus minimizing handling and the need for repeated injections [137]. Tramadol has been evaluated in bald eagles (Haliaeetus leucocephalus) [139], peafowl (Pavo cristatus) [140], red‐tailed hawks [141], Hispaniolan Amazon parrots [142–145], American kestrels [146], African penguins (Spheniscus demersus) [147], and Muscovy ducks (Cairina moschata domestica) [148]. Tramadol half‐life varies significantly between avian species (e.g., red‐tailed hawks 1.3 h versus African penguins 7.3 h). In those species for which pharmacokinetics studies have been published, the primary active metabolite O‐desmethyltramadol (M1) has also been measured and at concentrations that would be analgesic in other species; in most cases, it also has a longer half‐life than the parent drug. As there are large differences in oral bioavailability between species, such as 24% in Hispaniolan Amazon parrots [144] and almost 98% in bald eagles [139], pharmacokinetic realities must be taken into consideration when extrapolating doses between studies and routes of administration. Following oral administration, pharmacodynamic studies of psittacine and raptor species have shown large differences in the doses required for thermal antinociception, which could be explained by differences in pharmacokinetics alone. For example, Hispaniolan Amazon parrots showed thermal antinociception at 30 mg/kg PO and 5 mg/kg IV [144], but not at 10 or 20 mg/kg PO [145]. On the other hand, American kestrels treated with 5 mg/kg PO had significant antinociception while higher doses of 15 and 30 mg/kg PO resulted in a shorter duration of action and gastrointestinal adverse effects in a few birds [146]. No sedation or agitation was seen in these psittacine and raptor studies. In Muscovy ducks, 30 mg/kg PO effectively improved a number of pain‐associated variables in a temporary intertarsal joint arthritis model as assessed by ground‐reactive forces measured by a pressure‐sensitive walkway system [148]. Over the past two decades the advances in avian pharmacology, especially pharmacokinetics and pharmacodynamics, have been impressive. Studies of analgesics have provided insights as to which drugs in which species are efficacious, and which drugs are ineffective or potentially toxic. This reality is especially true of non‐steroidal anti‐inflammatory drugs (NSAIDs). To present a laundry list of NSAIDs and their doses would detract from the central message of this section which is that NSAIDs can be effective analgesics in birds, but they have potential for toxicity. This spectrum of activity varies with the drug, the dose, and the avian species in question. The following provides a historical perspective of NSAIDs and discusses a few select drugs that highlight the challenges inherent to their use in birds. A 2004 report in Nature documented that > 95% of old‐world vultures (Gyps spp.) died in India, Nepal, and Pakistan as a result of consuming carcasses of animals dosed with the veterinary drug diclofenac, an NSAID [149–151]. Cows are sacred in India and Hindu religion, and they often die naturally rather than being slaughtered for meat; open disposal of livestock carcasses is the norm. Scavengers, including vultures which are considered nature’s most successful scavengers and provide an array of ecological, economic, and cultural services [150], have year‐round widespread access to carrion. Studies of the vulture deaths subsequently documented that the carcasses contained high levels of diclofenac and other NSAID residues [151] that cause renal failure in birds. NSAID toxicity has been reported for raptors, storks, cranes, and owls, suggesting that the potential conservation impact of NSAIDs may extend beyond Gyps vultures and could be significant for New World vultures [152]. However, there are no reported mortalities for the NSAID meloxicam, which has been administered to over 700 birds from 60 species, a finding that supports other studies indicating the suitability of this NSAID to replace diclofenac in Asia [152]. Meloxicam, the most frequently used NSAID in birds, has a highly variable elimination half‐life among avian species when administered orally or intravenously (Tables 58.1 and 58.2) [82,85,153,154]. For example, the elimination half‐life (t1/2el) after oral administration of meloxicam (1 mg/kg) ranges from a low of 3.83 h in American flamingos to a high of 36.3 h in brown pelicans [155–157,160,163]. The clearance (in L/kg/h) of meloxicam can also vary significantly within the same species. For example, IV administration to three different age groups of emus yielded significantly faster clearance in emu chicks (1.32 L/kg/h) compared to adults (0.387 L/kg/h) [164]. Thus, from species to species and within species, there are clinically significant variations in meloxicam metabolism and elimination. As previously stated, this large variability in pharmacokinetics and pharmacodynamics from one avian species to another, and even significant age‐ and sex‐dependent variability, show that making pharmacologic generalizations across the class Aves must be exercised cautiously [75]. Table 58.1 Pharmacokinetic parameters of a single dose of meloxicam administered orally to a variety of avian species.
58
Comparative Anesthesia and Analgesia – Birds
Introduction
Respiratory system
Ventilation components
Larynx, trachea, syrinx
Primary and secondary bronchi
Air sacs
Muscles of respiration and the thoracic skeleton
Gas exchange component
Tertiary bronchi (parabronchi) and air capillaries
Paleopulmonic and neopulmonic parabronchi: lung volumes and direction of gas flow
Direction of gas flow and gas exchange
Cross‐current model of gas exchange
Control of ventilation
Cardiovascular system
Renal portal system
Preanesthetic patient evaluation and preparation
Physical restraint
Physical examination
Fasting
General pharmacologic considerations
Routes of drug administration
Allometric scaling and pharmacokinetics
Drugs used for analgesia and anesthesia
Opioids
Butorphanol
Nalbuphine
Hydromorphone
Morphine
Fentanyl
Methadone
Buprenorphine
Tramadol
Non‐steroidal anti‐inflammatory drugs
Meloxicam
Group and species
Cmax
(μg/mL)
Tmax
(h)
t1/2el
(h)
AUC0–last
(μg·h/mL)
AUC0–∞
(μg·h/mL)
Oral
bioavailability (%)
Reference
Psittacines
Hispaniolan Amazon parrots
(n = 8; 1 mg/kg)
3.7 ± 1.1
5.0 ± 3.2
15.8 ± 8.6
102
113 ± 71
49 to 75
(range)
[155]
Gray parrots
(n = 6; 1 mg/kg)
4.69 ± 0.75
13.2 ± 3.5
33.3 ± 3.1
81.0
(SE 5.73)
97.3
38.1 ± 3.6
[156]
Cockatiels
(n = 24; 1 mg/kg)
0.102
(0.019)a
1.27
(17.32)a
0.90
(32.34)a
–
0.295
(0.019)a
11
[82]
Raptors
Great horned owls
(n = 5; 0.5 mg/kg)
0.36 ± 0.08
7.8 ± 4.2
5.07 ± 4.5
–
–
62 ± 0.15
[157]
Red‐tailed hawks
(n = 5; 0.5 mg/kg)
0.18 ± 0.16
0.73 ± 0.23
3.97 ± 3.32
–
–
74 ± 0.48
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