Rebecca A. Johnson and David B. Brunson Department of Surgical Sciences, School of Veterinary Medicine, University of Wisconsin–Madison, Madison, WI, 53706, USA Adequate respiratory function and support are critical for safe anesthetic management, especially if inhalants are used. When patients present with coexisting respiratory disease, safe anesthetic practices can become challenging. It is therefore important to understand normal physiology, as well as the physiological changes and pathological progression of any coexisting disease, especially regarding the respiratory system [1–3]. Dogs and cats with respiratory disease may require sedation or anesthesia for diagnostic or surgical procedures that are either related or unrelated to the primary problem. Fortunately, pulmonary function support in these cases can frequently be accomplished with minimal specialized equipment. A brief overview of relevant ventilatory concepts and strategies, respiratory physiology, sedative/anesthetic agents, and specific case management is presented in the following text. Readers are referred elsewhere for a full review of basic respiratory physiology and pathophysiology [2, 3]. Ventilatory “drive” originates within respiratory centers of the central nervous system (CNS) ventral medulla. This brainstem circuitry of neurons and pathways is responsible for respiratory rhythm generation and respiratory pattern formation [4]. Although not yet fully elucidated, several models for this complex network have been proposed, many using reduced medullary slice preparations as models [5]. Respiratory rhythm and pattern are continuously altered by homeostatic control mechanisms that allow the animal to “adapt” to physiologic respiratory challenges (i.e., exercise, pregnancy) or pathologic conditions (i.e., neurologic or respiratory disease). This respiratory plasticity involves alterations via sensory (i.e., central and peripheral chemoreceptors and airway mechanoreceptors) and modulatory projections (i.e., serotonergic neurons), as well as many other conscious and unconscious processes that affect breathing (i.e., cortical inputs, cardiovascular disease, etc.). Collectively, inputs merge to form the spatiotemporal neural output that projects to the respiratory muscles. The primary inspiratory muscles are the diaphragm and inspiratory (external) intercostals which move the ribs forward and outward. However, accessory inspiratory muscles also play a role in breathing, especially during respiratory stress or disease (i.e., upper airway muscles innervated by the hypoglossal nerve) [6]. Altogether, synchronized respiratory muscle contraction generates a breath which ultimately drives alveolar ventilation and blood gas regulation (Figure 2.1) [7]. Figure 2.1 Graphic representation of the respiratory control system. The respiratory rhythm is generated within the brainstem and is modulated by multiple afferent inputs from various sensors such as mechanoreceptors and chemoreceptors. The detailed spatiotemporal output is projected to primary (diaphragm, inspiratory intercostal) and secondary (upper airway) respiratory muscles which contract to generate an adequate breath. During ventilation, respiratory mechanics and arterial oxygen and carbon dioxide levels change as conditions are altered, thus providing further sensory feedback to brainstem respiratory areas which imparts the respiratory continuum. Normal respiratory volumes (Figure 2.2) can be affected in anesthetized/sedated patients or in patients with altered physiology (i.e., pregnancy, disease, etc.). Many respiratory volumes are not measured clinically. However, tidal volume (VT) and functional residual capacity (FRC) can be measured in awake animals. Tidal volume refers to the air volume inspired or expired during a normal breath. The FRC is the amount of air remaining in the lungs after a normal expiration and can be affected by diaphragm position; although animal studies are sparse, this is especially important, as FRC likely decreases with pregnancy, bowel dilatation, abdominal masses, etc. [8]. The “inspiratory capacity” is the amount of air that can maximally inhaled beginning after a normal expiration. The “vital capacity” is the maximal amount of air that can be exhaled after filling them to their inspiratory capacity. The “residual volume” is the air left in the lungs following a maximal expiration. The “total lung capacity” is the lung volume following the greatest possible inspiratory effort (vital capacity and residual volume). Alveolar ventilation (V&c.dotab;A) is primarily driven by the arterial partial pressure of CO2 (PaCO2) sensed at central chemoreceptors such that, for a given metabolic state, V&c.dotab;A is directly and inversely related to PaCO2 (see the section titled “Carbon Dioxide”). For example, if V&c.dotab;A increases by 50%, PaCO2 is halved. It is defined as the rate that alveolar air is exchanged with atmospheric air. Although clinically it is frequently assumed to be minute ventilation (V&c.dotab;E), these concepts are not the same due to anatomic (and potential mechanical) dead space (see in the following text). Minute ventilation is composed of the tidal volume (volume of each breath; VT) and respiratory rate (f): V&c.dotab;E = VT × f. For most species, the VT is approximated at 8–20 ml kg−1; however, breathing frequency is highly variable [9]. Although smaller animals usually breathe at higher frequencies than larger animals based on metabolic oxygen requirements, birds are an exception, as they have relatively slower respiratory rates than other mammals with similar body sizes due to large anatomical dead space in conducting airways [9, 10]. Typically, the respiratory rate of awake dogs and cats varies between 10 and 30 breaths min−1. However, with sedation or general anesthesia, respiratory rates, as well as tidal volumes, are almost always reduced due to drug‐induced respiratory muscle relaxation and respiratory center depression, hypothermia, body position, and other external factors such as bandages. For example, respiratory frequencies commonly range from 6 to 20 breaths min−1 in spontaneously breathing, anesthetized dogs and cats, and V&c.dotab;E is significantly reduced from normal values of approximately 150–250 ml kg−1 min−1 [9, 11]. Because of the high variability in respiratory frequency, and the influences of VT on V&c.dotab;A, respiratory rate alone is not an adequate indicator of ventilatory function. Figure 2.2 Graphical representation of lung volumes. Total lung capacity is the volume of air in the lungs following maximal inspiration; it is the inspiratory capacity plus the functional residual capacity or vital capacity plus the residual volume after peak expiration. A normal tidal volume represents only a small portion of the total lung capacity. An important concept in the assessment of small‐animal ventilation is dead space (VD). Mechanical dead‐space ventilation occurs where two‐way flow of gases is present with no gas exchange. This mainly occurs in the endotracheal tube extending from the mouth, any monitor adapters added to the circuit, and the end of the Y‐piece before it splits into the inspiratory and expiratory tubing (Figure 2.3). Physiologic dead space is similar, except it occurs within the animal and is the arithmetic sum of both anatomic dead space and alveolar dead space. For example, no gas exchange occurs in the upper or conductive airways of the dog and cat (i.e., nares, larynx, pharynx, trachea, bronchi, nonrespiratory bronchioles). Thus, this portion of the VT is referred to as anatomic dead‐space ventilation. In addition, dead space also occurs in lung areas that are poorly perfused but adequately ventilated; little to no gas exchange occurs in these areas and this is referred to as alveolar dead‐space ventilation. The ratio of physiological dead space to the tidal volume (VD/VT) has been clinically calculated by the Enghoff modification of the Bohr equation which uses arterial CO2 (PaCO2) in the place of alveolar CO2 (PACO2) [12, 13]: PECO2 is the expired CO2 tension. VD/VT in awake dogs is approximately 35% but increases during inhalant anesthesia (to approximately 50% or more) [14]. Dead‐space ventilation is one reason that respiratory frequency should not be used to assess ventilation adequacy, since small, frequent breaths primarily ventilate only the anatomic dead space and alveolar ventilation may be inadequate. Rapid shallow ventilations may be effective for CO2 removal that reaches the upper airways, but not effective for O2 delivery to the alveoli. Slow deep ventilations are more likely to provide effective CO2 removal and O2 delivery, such as those seen in avian species. It is usually only the slow shallow ventilations that are easily recognized as ineffective by the anesthetist. Figure 2.3 An anesthetized cat demonstrating considerable mechanical dead space containing two‐way gas flow as the orotracheal tube extends at least 3–4 cm from the teeth to the capnometer adapter. In addition, the capnometer adapter adds to the mechanical dead space before attaching to the Y‐piece which contains unidirectional gas flow due to the one‐way valves of the rebreathing anesthetic system. The respiratory system is ultimately responsible for the effective transfer of O2 and CO2 between the atmosphere and the animal. Normally, the inspired air has only a trace percentage of CO2, while the animal’s arterial blood contains approximately 5% CO2. This differential gradient promotes CO2 movement out of the animal. By contrast, the normal atmospheric O2 content is approximately 21%, which is higher than the O2 in venous blood returning to the lungs, and O2 diffuses from the alveoli into the blood. Respiratory gases such as O2, CO2, and nitrogen are measured as a part of the total gases in the animal. Since the animal lives normally at a pressure of 1 atm (approximately 760 mmHg at sea level), the units used to measure gases are also in mmHg. Evaluation of the pulmonary system requires an understanding of both O2 and CO2 transfer. Each gas is somewhat independent of the other (but not exclusively; see the section titled “Interactions Between CO2 and O2 Transport”), and assessment of pulmonary function requires measurement of both gases. Thus, the “gold standard” for measuring adequacy of ventilation and patient oxygenation is the use of arterial blood gas analysis in both dogs and cats (Table 2.1) [15, 16]. Although peripheral CO2 receptors contribute to CO2‐induced ventilatory responses, the primary CO2‐/pH‐sensitive chemoreceptors are located in the CNS throughout the brainstem (i.e., retrotrapezoid nuclei, serotonergic raphe neurons, noradrenergic neurons in the locus coeruleus, nucleus of the solitary tract, pre‐Bötzinger complex, etc.) [7]. These chemoreceptors are extremely sensitive and even small deviations from a normal PaCO2 level affect ventilation linearly and dramatically. CO2 diffuses rapidly from tissues into red blood cells where it forms bicarbonate according to the reaction: CO2 + H2O ↔ H2CO3 ↔ H+ + HCO3−. The first reaction is slow in plasma, but rapid inside the red cell due to the presence of carbonic anhydrase. CO2 is then transported mostly as bicarbonate (approximately 81%) with a small amount dissolved in plasma (approximately 8%) and combined with amino groups of blood proteins (approximately 11%) [4]. Table 2.1 Normal arterial blood gas values in the unanesthetized dog and cat while breathing room air (approximately 21% O2). Values are expressed as mean (range). When breathing 100% O2 (as during anesthesia), PO2 values are expected to approach 600 mmHg with ideal gas exchange [11, 12]. As previously mentioned, in the steady‐state, PaCO2 is inversely proportional to V&c.dotab;A based on the alveolar ventilation equation: Overall ventilation is a measure of the animal’s ability to regulate the level of CO2 within the body; it is the only route for elimination. Adequate ventilation is present when CO2 is maintained at an optimal partial pressure in arterial blood (Table 2.1) [11, 15, 17]. Accordingly, hyperventilation occurs when V&c.dotab;A is increased and the resultant PaCO2 falls below normal values, while hypoventilation is defined as a reduction in V&c.dotab;A, increasing PaCO2 values from normal. Although blood gas analysis is the definitive way to assess the adequacy of ventilation, noninvasive technology such as the use of end‐tidal CO2 (ETCO2) capnometers has become increasingly popular in small‐animal (as well as exotic‐animal) practice, since they closely approximate alveolar CO2 levels under normal situations and can be easily used in most species (Figure 2.4). Since capnometry relies on adequate cardiac output to return CO2 to the lungs, as well as pulmonary function to eliminate CO2 from the body, it is an essential piece of equipment in any case where respiratory impairment or dysfunction is a concern and should become a routine monitor for any veterinary practice [18]. Figure 2.4 Capnometry (Vetcorder Airmate™ Capnograph, Sentier, Waukesha, WI) and pulse oximetry (Vetcorder™, Sentier, Waukesha, WI) used during isoflurane anesthesia on a domestic rabbit. Similar to the procedure in cats and dogs, the end‐tidal sample is drawn directly from the oral end of the endotracheal tube via a low dead‐space adapter. The capnometer produces a graphical waveform and is reading an end‐tidal CO2 level of 41 mmHg with a respiratory rate of 46 breaths min−1 in this rabbit (larger monitor). The pulse oximetry probe was placed on the foot and is reading a heart rate of 246 beats min−1 and hemoglobin saturation of 100% (smaller gray monitor). Inspired oxygen levels were approximately 100%. Once oxygen is carried from the conducting airways to the respiratory exchange tissues (respiratory bronchioles and alveoli), it quickly diffuses into the blood where it is carried both dissolved in solution and bound to hemoglobin (Hb). Approximately 1.36–1.39 ml of O2 can combine with 1 g of hemoglobin compared to only a small amount that is dissolved in blood (0.003 ml per 100 ml blood per mmHg PO2) [4, 19]. The total oxygen content of blood (CaO2) is therefore calculated as: CaO2 = (1.39 × Hb × SaO2) + (0.003 × PaO2), where SaO2 is the oxygen saturation of Hb. Thus, the amount combined with Hb impacts CaO2 to a greater extent than does the dissolved amount, and altered Hb levels (i.e., anemia) will have a profound effect on not just CaO2 but oxygen delivery (DO2) to the tissues, since DO2 = CaO2 × cardiac output. However, in an anemic animal, the dissolved portion plays an increasing important role; increasing PaO2 (through enriching inspired O2 levels, for example) will be advantageous to these patients. As determined by the oxygen–Hb dissociation curve, the amount of O2 carried by Hb increases rapidly until PO2 reaches approximately 60–70 mmHg where the curve flattens off (Figure 2.5). Many factors affect the curve placement including temperature, PCO2, pH, and 2,3‐diphosphoglycerate (2,3‐DPG) levels, an enzyme which competes with O2 for its binding site on Hb (Figure 2.5). Hemoglobin is normally nearly 100% saturated in healthy patients breathing room air (approximately 21% O2) and should always be above 90–95% [20]. In most normal anesthetized patients, the PaO2 is very high due to 100% O2 used as a carrier gas for the inhalant (frequently >500 mmHg; Table 2.1). As per the oxygen–Hb dissociation curve, a very high percentage of Hb will therefore be bound with O2. Since arterial blood gas analysis may not be available to every practitioner, pulse oximetry is a standard, yet indirect measure of oxygenation in anesthetized dogs and cats which measures pulse rate and estimates the Hb oxygen saturation (termed SpO2; Figures 2.4 and 2.6) [21]. However, one must be cautious, since significant pathology must exist in the anesthetized patient administered 100% oxygen for the PaO2 to drop to a level where the pulse oximeter will alert the anesthetist; a PaO2 of approximately 60 mmHg corresponds to an SpO2 of approximately 90%, and a PaO2 of approximately 80 mmHg corresponds to an SpO2 of approximately 95%. In addition, pulse oximetry does not indicate if adequate ventilation is present to remove CO2, since the pulse oximeter only estimates the percentage of Hb saturation with O2. Although this monitor has become one of the most widely used anesthetic monitors for a variety of species in veterinary medicine (Figures 2.4, 2.6, and 2.7), it provides little information concerning the animal’s ventilatory status during anesthesia unless severe pathology is present and should be used with care. Figure 2.5 Example oxygen–hemoglobin dissociation curves. The amount of O2 carried by hemoglobin increases rapidly until PO2 reaches approximately 60–70 mmHg when the curve flattens off. High levels of PO2 and hemoglobin saturation occur in the lung, whereas lower values occur in the tissues (near the bottom of the curve). The curve is shifted right by increases in temperature, PCO2, and 2,3‐DPG levels, and decreases in pH. Left shifts occur with opposite changes in temperature, PCO2, pH, and 2,3‐DPG levels. Figure 2.6 Pulse oximeter (Nonin Medical Incorporated, Plymouth, MN) used on an isoflurane‐anesthetized dog. As measured with a lingual probe, the heart rate was 42 beats min−1 and SpO2 read 100%. Isoflurane in 100% O2 was delivered via the anesthesia machine and a large animal mechanical ventilator. Hypoxemia (low PaO2) is commonly encountered in patients with respiratory impairment and is primarily due to one of the five reasons: low inspired O2 fraction, hypoventilation, diffusion impairments, ventilation–perfusion mismatching, and right‐to‐left shunts (cardiac or pulmonary). Discerning between these causes is sometimes difficult. However, some information can be obtained from the alveolar gas equation which predicts that the arterial (a) or alveolar (A) O2 tension will decrease with an increase in PCO2: PAO2 = PIO2 − (PACO2/R), where PIO2 is the inspired O2 level and R is the respiratory exchange ratio (approximately 0.8 in normal animals). The difference between alveolar and arterial oxygen levels is defined as the (A‐a) gradient. Substituting PaCO2 for PACO2, the (A‐a) oxygen gradient equation becomes: (A‐a) gradient = PAO2 − PaO2 = (PIO2 − 1.25PaCO2). Generally, values between 5 and 15 mmHg in animals breathing room air are considered normal [4]. Figure 2.7 Pulse oximeter (Vetcorder™, Sentier, Waukesha, WI) used on an isoflurane‐anesthetized domestic guinea pig (Cavia porcellus). This monitor can be a valuable tool to measure oxygenation in a variety of species (including cats and dogs). The probe is placed on the foot and displays a waveform (bottom trace) with an SpO2 of 98%. In addition, this monitor is reading the electrocardiogram (top trace). Hypoxemia due to low inspired O2 levels (i.e., low atmospheric pressure, use of nitrous oxide without appropriate O2 flows) does not usually result in increased (A‐a) O2 gradients, since frequently there is a subsequent increase in V&c.dotab;A (thus, decreasing PaCO2) secondary to hypoxemia. In addition, if hypoxemia is solely due to hypoventilation, the PAO2 and PaO2 decrease, while the PaCO2 and PACO2 must increase and the (A‐a) difference should not change. However, in the case of ventilation–perfusion abnormalities or right‐to‐left shunting (both very common during anesthesia), the (A‐a) gradients are significantly elevated. These two are differentiated from each other, as administration of 100% O2 to patients with ventilation–perfusion abnormalities significantly increases the PaO2, whereas PaO2 levels fail to return to normal in patients with significant shunting, as blood traverses across the lungs or through the heart and does not pick up the oxygen [4]. Although management of CO2 and O2 within the body appears independent, they are not mutually exclusive of each other [9]. For example, acidity (lower pH) associated with carbonic acid produced from CO2 in tissues promotes O2 release from Hb with no change in dissolved oxygen levels, whereas CO2 release from the lungs promotes oxygen uptake; this is known as the Bohr effect. In addition, the Haldane effect states that deoxygenation of Hb causes Hb to become a weaker acid, this increases its ability to carry hydrogen ions [9]. Normally, there are three ventilatory cycle phases: inspiration, expiration, and an expiratory pause (which are important to understand in evaluating breathing or during adjustments of ventilatory support). Inspiratory pauses are not normally present in conscious dogs and cats. Expiration, in most cases, is a passive process related to the elastic recoil of the chest. One exception is the horse in which expiration normally has an active phase even at rest [22]. The respiratory cycle is primarily changed by adjustments of the “pause” phase of the cycle. Ventilatory rates are usually increased by decreasing the pause time between each inhalation and exhalation. For mechanical ventilation, adjustment of the inspiratory to expiratory ratio (I:E) is possible and is important when changing the respiratory rate (Figure 2.8). As a guideline, the inspiratory time should be less than or equal to the expiratory time. Thus, the I:E ratio will be at least 1:1, but may vary to a ratio as high as 1:5. For example, if inspiratory time is kept at 1–2 s in duration and the respiratory rate is 10 breaths min−1, then the I:E ratio will be between 1:5 and 1:2. Typically, the I:E ratio is usually not less than 1:1, which allows adequate time for lung filling and emptying. Mechanical ventilation differs significantly from spontaneous ventilation. Normally, air enters the respiratory system because of contraction of inspiratory intercostal muscles and the diaphragm. During normal inspiration, the change in alveolar pressure is only about 1 cm of water pressure [23]. However, IPPV is not a “normal” situation for the animal, as positive pressure is “forced” into the lungs; too much pressure can result in detrimental effects such as reducing cardiac output, volutrauma, barotrauma, etc. (Figure 2.9) [24]. Figure 2.8 A mechanical ventilator used for small animals. Respiratory rate, inspiratory flow, maximum pressure, and I:E ratio can all be adjusted to reach the targeted end‐tidal CO2 level. Ultimately, the concern with IPPV is that intermittent high thoracic pressures can lead to tissue perfusion failure. For example, although during the positive pressure breath, pressurization of chest can augment cardiac ejection, increased intrathoracic pressure often exceeds central venous pressure, resulting in a decrease in venous return to the heart. The resulting decrease in ventricular filling may reduce cardiac output on the subsequent heartbeat and often decrease systolic arterial pressure [24]. To minimize the negative effects of IPPV, the inspiratory time should be kept short and should not exceed 2–3 s. Additionally, the lowest peak inspiratory pressure (PIP) capable of inflating the lungs should be used. In many animals, a PIP of less than 15 cmH2O will be sufficient to fully inflate the lungs. In mammals, the guideline is to keep the PIP below 20 cmH2O; for reptiles and birds the guideline is a maximum of 15 cmH2O [25]. In animals with normal thoracic compliance, if PIP is set to approximately 10 cmH2O, respiratory rate is approximately 10 breaths min−1, and VT is approximately 10–15 ml kg−1, the PaCO2 frequently approaches normal levels; minor adjustments can then be made to achieve the precise, desired PaCO2 for the patient. Figure 2.9 Thoracic radiograph depicting subcutaneous emphysema secondary to barotrauma following excessive intrathoracic pressure delivered to cat. The skin has been separated from the body wall by a large volume of radiolucent air (white arrow). Because of the aforementioned concerns, mechanical ventilation may not be warranted in every patient. If the patient is healthy, having a short procedure (<1h), does not display airway obstruction, is not apneic, and is oxygenated, mild‐to‐moderate hypercapnia is usually well tolerated. However, ventilatory support should be available to all animals during general anesthesia and can include assisted “hand ventilation” by an anesthetist or the use of mechanical ventilation. This is because multiple factors contribute to an inability to adequately ventilate including drug‐induced respiratory depression, age, body weight, and preexisting disease. Ventilation not only ensures that the patient is oxygenated and CO2 is removed, but also ensures consistent delivery of anesthetic gases. Delivery, maintenance, and recovery from inhaled anesthetics uniquely depend on the movement of the agents through the lungs. Consistent and effective ventilation with controlled or assisted ventilation makes inhalant anesthesia both more precise and easier to perform. However, if the goal is to keep PaCO2 or ETCO2 at normal levels, consistent ventilation must occur, as CO2 levels quickly rise when ventilation is stopped, even for a few minutes. Any patient can benefit from effective ventilatory support; however, in the patient with preexisting respiratory disease, it is essential. In many patients with respiratory insufficiency, O2 supplementation can be beneficial. Criteria for preoxygenation include animals that are marginal or unable to maintain Hb O2 saturation when breathing room air. Cyanosis is a sign of an oxygenation crisis that warrants immediate O2 supplementation. The ability to see cyanosis indicates that approximately 5 g of Hb is in the reduced form or is not carrying O2. However, it can be difficult to visually determine cyanosis in certain situations, such as in poor lighting conditions or if the animal is severely anemic. Oxygen can be administered via a face mask, an O2 chamber, or via nasal O2 cannulas (Figure 2.10). However, inspired O2 levels may not reach 100% with these techniques. For example, low‐flow nasal insufflation with 100% O2 only resulted in approximately 32–61% inspired O2 and was dependent on insufflation rate, respiratory rate, and tidal volume in cadaveric dog preparations [26]. However, high‐flow nasal insufflation shows more promise [27]. In addition, mask oxygenation should not be forced on the patient if the process produces anxiety or stress due to increased patient O2 demands associated with struggling. Unless the animal is cyanotic, the benefit of preoxygenation is that it provides a longer time before a patient becomes hypoxic during apneic periods; for example, following anesthetic induction [28, 29]. Specifically, 3 min of preoxygenation in dogs via tight‐fitting face mask increased the time to Hb desaturation from approximately 70 s (room air) to 298 s (O2 supplementation) following propofol induction [28]. Preoxygenation with 100% delivered O2 theoretically fills the alveolus and FRC with a higher‐than‐normal O2 concentration. If the animal becomes apneic or if tracheal intubation is delayed, additional O2 is then available to the pulmonary blood. Effectively washing out gases normally found in the alveoli and replacing them with a higher O2 concentration take multiple minutes of preoxygenation [28], and any break in the breathing of the supplemental O2 will require restarting of the oxygenation procedures. Thus, unless the animal is in respiratory distress and is hypoxic prior to anesthesia, preoxygenation may not benefit the animal and may actually precipitate additional failure through excitement or anxiety. Figure 2.10 Bilateral nasal oxygen cannulas and supplementation in a dog. The tubing is secured to the lateral nasal area and upper lip to reduce the chances for dislodgment of the cannula. The tubing is further connected to an oxygen flowmeter to deliver gases enriched with oxygen. Adequate patient ventilation results from careful integration of all components of the respiratory system. For example, the (i) neural control system, (ii) upper and lower airways, (iii) pulmonary parenchyma, and (iv) thoracic bellows mechanism all need to function normally to maintain patient oxygenation and ventilatory capacity. If alterations occur in any or all these constituents, hypoxia or hypercapnia may occur; these untoward effects may be enhanced in anesthetized patients due to the inability to compensate for such changes. Thus, each part of the respiratory control system must be considered when creating an anesthetic plan for not just normal patients but also for patients with disorders of ventilation. Since respiratory function is frequently diminished simply by using respiratory‐depressant agents such as inhalants [30, 31] and opioids (although variably) [32] in many species, respiratory complications from coexisting diseases can impair ventilation even further. Anesthetic management of these cases should focus on keeping the patient oxygenated, ventilated, and perfused. Frequently, the anesthetic agents themselves are not as important as the strict patient management associated with these high‐risk cases. Many pure opioid analgesics depress minute ventilation via effects at mu‐opioid receptors near and pathways projecting to brainstem respiratory centers. These dose‐dependent effects can be seen as a reduction in respiratory frequency, tidal volume, or both [24], and are more pronounced with the pure mu‐opioid receptor agonists when compared to the mixed agonist–antagonists or partial agonists [33, 34]. Mixed agonist–antagonists such as butorphanol may be used to reverse, at least in part, the respiratory depression seen with pure mu‐opioid agonists [34, 35]. The ventilatory response to hypoxia is reduced and the CO2 response is shifted to the right and the slope is decreased, resulting in a rightward shift of the apneic threshold (level of CO2 where breathing no longer occurs and the animal becomes apneic; Figure 2.11) [24, 36]. Interestingly, ventilatory responsiveness may show maturational changes, as the newborn dog appears to have increased sensitivity to the respiratory‐depressant effects of some opioids (e.g., morphine) [36]. In addition, these effects may be reversed with opioid receptor antagonists such as naloxone or naltrexone [24]. In any small‐animal patient with significant respiratory disease, caution should be used when opioids are administered systemically; careful monitoring of ventilation (PaCO2) and oxygenation (PaO2) is highly recommended. Similar to opioids, other tranquilizers and sedatives such as acepromazine [37] and alpha‐2‐adrenergic receptor agonists (e.g., medetomidine, dexmedetomidine) [38] may also have at least some respiratory‐depressant effects in dogs and cats, although they appear to be minimal when the sedative agents are used alone and at clinical doses. For example, in dogs administered medetomidine, (the racemic mixture of levomedetomidine and dexmedetomidine (20 and 60 μg kg−1)), respiratory rate significantly decreased and PaCO2 significantly increased, albeit with little effect on PaO2 [33,39–42]. By contrast, cats administered dexmedetomidine alone did not show decreased respiratory rates and medetomidine alone did not significantly alter arterial blood gas values significantly [43, 44]. It is important to recognize that the respiratory‐depressant effects of the alpha‐2‐adrenergic receptor agonists are likely to be exaggerated when used in combination with other anesthetic/analgesic agents (i.e., propofol, opioids) [42]. Figure 2.11 Example ventilatory response curves to inspired CO2 levels. As CO2 levels rise, ventilation increases rapidly as the central chemoreceptor system is tightly regulated. Pharmacologic agents such as inhalants and opioid analgesics may shift this curve to the right (decreasing sensitivity) and increase the apneic threshold (when ventilation ceases). Acepromazine has minimal effects on the respiratory control system in unanesthetized patients, at least at clinical doses. Studies in small animals show a reduction in respiratory rate following acepromazine administration with minimal changes in minute ventilation, PaO2, PaCO2, and arterial pH [34, 45, 46]. Similarly, benzodiazepines (diazepam, midazolam) are associated with minimal pulmonary effects in small animals [47]. However, as always, in animals with respiratory disease, care should be used when administration of the agents is required; proper respiratory monitoring should be performed. Similar to the pure mu‐opioid agonists, most induction and inhalational agents depress minute ventilation significantly. PaCO2 increases because of decreased respiratory rates and volumes; these changes are frequently dose‐dependent. For example, propofol administered to small animals significantly reduces diaphragm contractility [48], arterial pH, and PaO2 while significantly increasing PaCO2; large doses can result in apnea [49]. Propofol also suppresses carotid body chemoreceptor activity involved in hypoxia‐induced respiratory responses [50]. Alfaxalone dose‐dependently decreases respiratory rate and minute volume similar to propofol in cats [51] and in dogs [52], although apnea appears to be more of an issue in dogs. By contrast, intravenous ketamine administration appears to have less respiratory‐depressant effects than other agents, especially at low doses. Indeed, ketamine infusions improve respiratory (and cardiovascular) depressant effects associated with inhalant anesthetics in small animals when used as part of a balanced anesthesia technique [53]. All inhalant agents shift the CO2–ventilatory response curve to the right and reduce the slope in a dose‐dependent manner, and the CO2 response curve is almost horizontal at high inhalant levels (2.0 × the minimum alveolar concentration) (Figure 2.11) [54, 55]. In addition, the ventilatory response to hypoxia is also diminished via significantly depressed carotid body chemoreflexes [56]. When inhalant anesthetics are used in animals with respiratory disease (as well as in normal patients), the anesthetist should be ready to assist or control ventilation, since the combination of anesthetic agents used (i.e., opioids along with sedatives and inhalants) is expected to result in significant respiratory depression in almost every anesthetized case. As previously stated, the choice of anesthetic/analgesic agents themselves is likely not as important as specific case management during respiratory disease (i.e., monitoring, ventilatory support, etc.). In normal anesthetized dogs and cats, as well as in patients with concurrent respiratory disease, general guidelines have been created by McDonell and Kerr (2015) when approaching ventilatory support: [9] The following serve as examples of commonly occurring respiratory diseases in companion animal practice with some suggested approaches to case management [9]. They are separated into: (i) upper airway disease, (ii) lower airway or parenchymal disease, and (iii) extrapulmonary disease or disorders of the thoracic wall. Diseases may also be classified as obstructive (e.g., asthma, laryngeal paralysis [may also be restrictive], etc.) and restrictive respiratory disease (e.g., pneumonia, obesity, etc.). The main difference is that with obstructive disease, patients have difficultly exhaling a complete breath, whereas with restrictive disease, patients have difficulty in inhaling a complete lung volume. The clinical manifestation of restrictive disease is a decrease in the vital capacity of the lung (Figure 2.2); this contrasts with obstructive disease where inspiratory flow rates and volumes may be normal [1]. Disturbances in the upper airway include, but are not limited to, laryngeal paralysis, brachycephalic airway syndrome, collapsing trachea, intraluminal or extraluminal masses or abscesses, etc. Patients may present for anesthetic procedures unrelated to the airway disturbance or for surgical intervention to correct the primary airway problem. If an airway crisis is reached, patients frequently present anxious, dyspneic, and hypoxemic. Unless an emergent situation arises, a thorough history and physical exam should be performed, emphasizing respiratory system evaluation. Thoracic radiographs are almost always warranted in these patients to rule out concomitant pulmonary disease. Further diagnostics, including blood work, should be directed toward individual pathologic conditions. Arterial blood gas analysis and pulse oximetry measurements in the conscious patient should be performed as needed. In addition, these patients are more likely to overheat under conditions that would not make a normal dog hyperthermic and body temperature should be initially taken and monitored often during the perianesthetic period. A thorough history and signalment can be highly suggestive for specific disorders. For example, brachycephalic airway syndrome is usually identified in younger to middle‐aged animals and includes abnormalities such as stenotic nares, aberrant nasal conchae, elongated soft palate, everted laryngeal saccules, laryngeal collapse, hypoplastic trachea, and bronchial collapse, all of which may predispose to upper airway obstruction [57]. Brachycephalic dogs are at a higher risk for perianesthetic and postanesthetic complications compared with non‐brachycephalic dogs, including aspiration pneumonia and regurgitation, among others [57]. However, corrective upper airway surgery in brachycephalic dogs reduced the odds of postanesthetic complications during subsequent anesthetic events in these same patients [58]. Laryngeal paralysis is recognized mostly in older dogs; however, it can also be found in cats [59]. It can slowly develop until it reaches a stage that causes significant dyspnea, as the larynx is unable to adduct properly to allow sufficient air flow into or out of the trachea (Figure 2.12). A sedated laryngeal exam may also be necessary to rule out laryngeal dysfunction in brachycephalic patients or patients with suspected laryngeal paralysis (see the sections titled “Sedation and Analgesia” and “Anesthetic Induction”). Tracheal collapse usually occurs in middle‐aged, small‐breed dogs that present with signs ranging from mild airway irritation to severe paradoxical cough and dyspnea [60]. Definitive diagnosis is made through dynamic radiographs, fluoroscopy, or bronchoscopy, and treatment consists of various medical and surgical corrective measures [60]. If the animal’s disposition is anxious or easily stressed, and the situation is not emergent, Fear Free techniques, including the use of anti‐anxiety drugs, should be used prior to entering the clinic [61]. For example, “Chill” protocols including agents such as gabapentin (10–30 mg kg−1 PO), trazodone (2–10 mg kg−1 PO), melatonin (0.5–3 mg kg−1 PO), acepromazine (0.025–0.05 mg kg−1 PO), dexmedetomidine solution (20–30 μg kg−1 oromucosal; or Sileo (Zoetis, Kalamazoo, MI) 125 μg m−2 oromucosal), and their combinations can be useful to reduce patient stress; however, debate about whether these drugs cause anxiolysis or simply sedation still exists [62]. Prior to anesthesia, systemic abnormalities should be treated, dehydration corrected, and the patient stabilized if possible. Preoxygenation as described in the preceding text is advised if tolerated by the patient. Food should be withheld as appropriate to reduce chances for aspiration, as this can be a complication associated with upper airway dysfunction (see “Sedation and Analgesia”). An IV catheter should always be placed in these patients as soon as possible either prior to or immediately following sedation. Sedatives should be chosen on an individual patient basis and doses adjusted toward individual pathologies. However, sedation with low doses of tranquilizers (i.e., acepromazine 0.01–0.05 mg kg−1 IM, IV) or anxiolytics (i.e., dexmedetomidine at 0.001–0.010 mg kg−1 IM, IV) and nonstressful handling are important during the preanesthetic period. Although some upper airway disorders (i.e., laryngeal paralysis, tracheal collapse, brachycephalic airway syndrome, etc.) are not believed to be painful conditions in and of themselves, opioid analgesics are frequently administered (i.e., butorphanol at 0.02–0.05 mg kg−1 IM, IV; hydromorphone at 0.1–0.2 mg kg−1 IM, IV, etc.). If respiratory depression is a concern, and any anticipated pain is considered mild‐to‐moderate, buprenorphine (0.02–0.05 mg kg−1 IV, IM, TM in cats) can also be used. The synergism of sedatives with the opioid often reduces anxiety and stress and improves the animal’s comfort and breathing without significant respiratory depression; if inhalant anesthetics are subsequently used, levels may be reduced. Figure 2.12 Laryngoscopic view of (a) the normal canine larynx and (b) a canine larynx with laryngeal paralysis showing the cuneiform (cun) and corniculate (cor) processes of the arytenoid cartilages. In the normal larynx, the arytenoid cartilages open, allowing sufficient air to move through the rima glottidis (rg) during a respiratory excursion (vf = vocal fold). With laryngeal paralysis, the arytenoid cartilages fail to adduct properly, resulting in a narrowed rima glottidis and increased resistance to breathing. Source: Photo courtesy of Dr. Robert Hardie, University of Wisconsin. One caveat associated with sedation in patients with upper airway disease is the potential for vomiting and subsequent aspiration. Thus, if possible, antiemetics such as maropitant (1 mg kg−1 IV, SC; 2–4 mg kg−1 PO) should be administered at least 1–2 h prior to sedation (depending on route) and/or general anesthesia [63]. In addition, mu‐opioid agonists and alpha‐2‐adrenergic receptor agonists should be administered cautiously, especially when used in combination. The IV route should be considered over IM opioid administration, since IV use may somewhat reduce the chances for vomiting [64]. In any case, if the procedure is anticipated to be painful, opioid agonists should not be withheld; however, the analgesic plan should be tailored to the specific case. For example, a multimodal approach including the use of ketamine and nonsteroidal anti‐inflammatory drugs (NSAIDs) may be warranted. However, if steroid administration is likely to occur during the procedure to reduce airway inflammation, NSAIDs should be avoided. If appropriate, local anesthetic techniques including nerve blocks, fascial plane blocks, and epidurals should be used if appropriate. Depending on required duration of action, lidocaine, bupivacaine, mepivacaine, or ropivacaine can be chosen. In addition, the intraincisional use of liposomal bupivacaine (Nocita®, Elanco Animal Health, Greenfield, IN) may be an excellent adjunct in the analgesic plan, as analgesia occurs locally with no systemic sedative effects. However, no studies exist concerning the use of liposomal bupivacaine in airway surgeries. Along with preoxygenation, securing an airway quickly is quite important in these patients. All equipment including orotracheal tubes, laryngoscopes, ties, and cuff inflators should be organized and accessible. Orotracheal intubation is the most common and efficient means to deliver 100% oxygen and inhalants. However, in an emergent situation, supraglottic airway devices (v‐gel®, Jorgensen Laboratories, Loveland, CO) may be used in cats (and rabbits) (Figure 2.13). An important factor associated with respiratory fatigue is airway resistance. Resistance to laminar flow is governed by the Hagen–Poiseuille law, and is estimated by:
2
Respiratory Disease
Introduction
Ventilatory Control
Respiratory Volumes and Ventilation
Respiratory Gases
Carbon Dioxide (CO2)
Dog
Cat
pH
7.41 (7.35–7.46)
7.39 (7.31–7.46)
PCO2 (mmHg)
36.8 (30.8–42.8)
31.0 (25.2–36.8)
HCO3− (mEq/l)
22.2 (18.8–25.6)
18.0 (14.4–21.6)
PO2 (mmHg)
92.1 (80.9–103.3)
106.8 (95.4–118.2)
is the metabolic production of CO2 and 0.863 is a constant that corrects for dissimilar units. CO2 is approximately 20–24 times more diffusible than oxygen; as such, there is complete equilibration across the alveolar membrane under normal situations. Thus, the PaCO2 is essentially the same as the PACO2 (within a few mmHg, primarily due to a small amount of dead‐space ventilation) in healthy animals. However, these values may significantly diverge in the presence of pulmonary disease states such as profound ventilation–perfusion inequalities or pulmonary shunting.
Oxygen
Interactions Between CO2 and O2 Transport
Ventilatory Cycle Phases and Ventilatory Support
Ventilatory Cycle
Ventilatory Support with Intermittent Positive Pressure Ventilation (IPPV)
Oxygen Supplementation
Ventilation in Conscious Patients
Pharmacologic Effects on Ventilation
Opioids
Other Sedatives and Tranquilizers
Injectable and Inhalant Anesthetics
Anesthetic Management of Specific Disorders
Upper Airway Disorders or Dysfunction
History, Signalment, and Diagnostics
Preanesthetic Considerations
Sedation and Analgesia
Ventilatory Support
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