DEFINITIONS

A number of terms are used to describe the function of the respiratory tract, or abnormalities that arise because of a variety of diseases. Many of these terms are described in more detail in the text that follows, but a brief definition of each is provided here:

HYPOXIA

Failure of the tissues to receive an adequate supply of oxygen occurs in a number of ways and the differences are clinically relevant, in that they are associated with failure of different organ systems, different diseases, and have fundamentally different pathophysiological mechanisms

COMPENSATORY MECHANISMS

Compensation of respiratory insufficiency occurs as both short-term and long-term events. Short-term compensatory mechanisms for low arterial oxygen tension or oxygen delivery to tissues occur within seconds to minutes and include respiratory, cardiovascular and behavioral responses. Stimulation of respiratory centers in the medulla oblongata by low arterial oxygen tension (P_ao₂) and high arterial carbon dioxide tension (P_aco₂) causes an increase in respiratory minute volume mediated by an increase in tidal volume and respiratory frequency. Both low oxygen tension and high carbon dioxide tension in arterial blood, together or separately, are potent stimulators of these events. Inadequate tissue oxygenation also stimulates an increase in cardiac output, mainly as a result of increased heart rate and to a lesser extent by an increase in stroke volume. Splenic contraction, in those species such as the horse in which the spleen is an important reservoir of red blood cells, increases both blood volume and hemoglobin concentration, thereby increasing the oxygen-carrying capacity of blood. Hypoxemia also causes animals to attempt to decrease their oxygen requirement by decreasing physical activity, including moving and eating.

Longer-term compensatory mechanisms include an increase in erythropoietin secretion by the kidney with subsequent increases in bone marrow production of red blood cells and an increase in hemoglobin concentration in blood. This polycythemia increases the oxygen-carrying capacity of blood. Severe polycythemia, such as occurs with congenital cardiac anomalies causing chronic right-to-left shunting, increases the viscosity of blood and impairs tissue perfusion, increases the workload of the heart and the risk of thromboembolism. Longer-term compensatory mechanisms also include changes in ventilatory pattern, such as in horses with heaves, and behavior.

CARBON DIOXIDE RETENTION (HYPERCAPNIA)

Respiratory insufficiency results in decreased elimination of carbon dioxide and its accumulation in blood and tissues. Animals breathing room air that are hypercapnic are always hypoxemic. Increasing the oxygen tension of inspired air can alleviate the hypoxemia but, by reducing hypoxic stimulation of the respiratory center, can cause further increments in arterial Pco₂.

Acute hypercapnia causes a respiratory acidosis that reduces both blood and cerebrospinal fluid pH.² The clinical signs of acute hypercapnia are initial anxiety followed by central nervous system depression and eventual coma and death. These clinical abnormalities are attributable to declines in the pH of cerebrospinal fluid (CSF), a consequence of the ease with which carbon dioxide crosses the blood–brain barrier. Decreases in CSF pH are greater for respiratory acidosis than for a similar degree of metabolic acidosis. Severe hypercapnia also causes peripheral vasodilation, which can contribute to arterial hypotension, and cardiac arrhythmia. The acid–base effects of chronic hypercapnia are compensated by renal mechanisms that return the arterial and CSF pH to almost normal and therefore do not cause more than mild clinical disease in most instances. So long as oxygen delivery to tissue is maintained, animals can tolerate quite high arterial carbon dioxide tensions for a number of days or longer – this is referred to as ‘permissive hypercapnia’ and is sometimes an alternative to artificial or mechanical ventilation of animals with respiratory insufficiency.

RESPIRATORY FAILURE

Respiratory movements are involuntary and are stimulated and modified by the respiratory centers in the medulla. The centers appear, at least in some species, to have spontaneous activity that is modified by afferent impulses to higher centers, including: cerebral cortex and the heat-regulating center in the hypothalamus; from the stretch receptors in the lungs via the pulmonary vagus nerves; and from the chemoreceptors in the carotid bodies. The activity of the center is also regulated directly by the pH and oxygen and carbon dioxide tensions of the cranial arterial blood supply. Stimulation of almost all afferent nerves may also cause reflex change in respiration, stimulation of pain fibers being particularly effective.

Respiratory failure is the terminal stage of respiratory insufficiency in which the activity of the respiratory centers diminishes to the point where movements of respiratory muscles cease. Respiratory failure can be paralytic, dyspneic or asphyxial, or tachypneic, depending on the primary disease.

The respiratory failure that occurs in animals with pneumonia, pulmonary edema and upper respiratory tract obstruction is caused by combinations of hypoventilation, ventilation/perfusion mismatch and diffusion impairment, which leads to hypercapnia and hypoxemia. Hypercapnia and hypoxia stimulate the respiratory center and there is a potent respiratory drive evident as markedly increased respiratory rate and effort. As the disease progresses these changes become more marked until death occurs as a result of central nervous system or cardiac failure. Animals that die of the central nervous system effects of respiratory failure typically have dyspnea followed by periods of gasping and apnea just before death.

Paralytic respiratory failure is caused by depression of the respiratory centers or paralysis of the muscles of respiration. Depression of the respiratory center occurs with poisoning by respiratory center depressants, such as general anesthetics, or damage to the respiratory center, such as might occur with brainstem injury. Paralysis of respiratory muscles occurs in disease such as botulism, tetanus, strychnine poisoning, white muscle disease, severe hypocalcemia and tick paralysis. The signs of paralytic respiratory failure are a gradual or abrupt cessation of respiratory movements without preceding signs of increased respiratory effort or dyspnea. The animal is often unconscious, or unable to move, during the later stages of the disease.

The differentiation of these types of failure is of some importance in determining the type of treatment necessary. In the paralytic form of respiratory failure the optimal treatment is mechanical ventilation, along with removal of the inciting cause. Administration of respiratory stimulants is seldom effective as sole therapy. The more complex pathogenesis of respiratory failure in most diseases requires a therapeutic approach that removes each of the underlying defects. In most cases this is achieved by treating the inciting disease, for example administering antimicrobials to an animal with pneumonia or furosemide to an animal with pulmonary edema, in addition to supportive care including, potentially, nasal or pharyngeal insufflation with oxygen, or mechanical ventilation.

ABNORMALITIES IN RATE, DEPTH AND EASE OF BREATHING

Polypnea is a rate of breathing that is faster than observed in clinically normal animals of the same species, breed, age, sex and reproductive status in a similar environment.

Tachypnea also describes an increased rate of breathing, although with the implication that breathing is shallow (i.e. of a reduced tidal volume).

Hyperpnea is an abnormal increase in the rate and depth of breathing (an abnormally high minute volume) but the breathing is not labored and is not associated with signs from which one could infer represent distress on the part of the animal (i.e. the animal is not dyspneic). This assessment requires measurement of minute ventilation or arterial blood gas tensions.

Dyspnea is a term borrowed from human medicine, in which it refers to the sensation of shortness of breath or air hunger. It is used in veterinary medicine to describe labored or difficult breathing in animals that also display some signs of distress, such as anxious expression, unusual posture or stance, or unusual behavior.

Dyspnea is a physiological occurrence after strenuous exercise and is abnormal only when it occurs at rest or with little exercise. It is usually caused by hypoxia with or without hypercapnia, arising most commonly from diseases of the respiratory tract. In pulmonary dyspnea one other factor may be of contributory importance; there may be an abnormally sensitive Hering–Breuer reflex. This is most likely to occur when there is inflammation or congestion of the lungs or pleura. Rapid, shallow breathing results.

Expiratory dyspnea is prolonged and forceful expiration, usually associated with diffuse or advanced obstructive lower airway disease. The dyspnea of pulmonary emphysema is characteristically expiratory in form and is caused by anoxic anoxia and the need for forced expiration to achieve successful expulsion of the tidal air. It is commonly accompanied by an audible expiratory grunt in ruminants but less so in pigs and almost never in horses.

Inspiratory dyspnea is prolonged and forceful inspiration due to obstruction of the extrathoracic airways, such as with laryngeal obstruction or collapse of the cervical trachea. It may also be associated with abnormalities that restrict thoracic expansion, such as restrictive lung diseases and space-occupying lesions of the thorax. It is accompanied by a stridor or loud harsh sound on inspiration when the cause is obstruction of the extrathoracic airways, such as is typical of laryngeal or tracheal disease.

Open-mouth breathing is labored breathing with the mouth held open, commonly with the tongue protruded in ruminants and most commonly associated with advanced pulmonary disease or obstruction of the nasal cavities.

DISEASES CAUSING DYSPNEA AT REST OR LACK OF EXERCISE TOLERANCE

Dyspnea, along with hypoxemia and hypercapnia, are the clinical and laboratory findings most likely to attract attention to the possible presence of disease in the respiratory system. A brief summary of the causes of dyspnea is outlined in Figure 10.1. It is most important, when attempting to differentiate diseases that cause dyspnea, to include diseases of systems other than the respiratory system that can result in dyspnea. Dyspnea at rest is usually, but not always, caused by respiratory tract disease, whereas exercise intolerance can be caused by disease in the respiratory, cardiovascular, musculoskeletal and other body systems.

Fig. 10.1 The causes of dyspnea.

Respiratory tract disease

Respiratory tract diseases interfere with normal gas transfer, through the mechanisms discussed above. Characteristics of respiratory disease that lead to dyspnea or lack of exercise tolerance include:

• Flooding of alveoli with inflammatory cells and/or protein-rich fluid – pneumonia and pulmonary edema

• Atelectasis (collapsed alveoli and small airways) – pleural effusion, hemothorax, hydrothorax, pneumothorax, chylothorax, pyothorax, prolonged recumbency of large animals and diaphragmatic hernia

• Airway obstruction – nasal obstruction, pharyngeal/laryngeal obstruction, tracheal/bronchial obstruction, bronchoconstriction and bronchiolar obstruction.

Cardiovascular disease

This causes inadequate perfusion of tissues including the lungs. There is reduced oxygen delivery to tissues, even in the presence of normal arterial oxygenation:

• Cardiac disease. Cardiac dyspnea results from heart failure and is multifactorial. In animals with dyspnea attributable to cardiac disease there are other readily evident signs of heart failure

• Peripheral circulatory failure – usually due to hypovolemic shock, although shock associated with toxemia, including endotoxemia, can cause dyspnea. There are always other prominent signs of disease.

Diseases of the blood

These cause inadequate delivery of oxygen to tissues because of anemia or presence of hemoglobin that is unable to carry oxygen.

• Anemia – an abnormally low concentration of hemoglobin

• Altered hemoglobin – methemoglobinemia (e.g. in nitrite poisoning of cattle, red maple toxicosis of horses), carboxyhemoglobinemia.

Nervous system diseases

Diseases of the nervous system affect respiratory function by one of several mechanisms:

• Paralysis of respiratory muscles occurs in tick paralysis or botulism. Tetanic spasm of respiratory muscles, such as in tetanus or strychnine toxicosis, also impairs or prevents alveolar ventilation. Both flaccid and tetanic paralysis cause hypercapnia and hypoxemia and, in extreme situations, death by suffocation

• Paralysis of the respiratory center, as in poisoning by nicotine sulfate, or overall central nervous system depression, causes hypoventilation because of impaired ventilatory drive

• Stimulation of the respiratory center, so-called neurogenic dyspnea, occurs as a result of stimulation of the center by a small irritative lesion, such as in animals with encephalitis, or administration of drugs, such as lobeline, that increase sensitivity of the respiratory center to hypoxemia or hypercapnia.

Musculoskeletal diseases

• Muscle diseases. Diseases of the respiratory muscles can impair ventilation. These include white muscle disease in lambs, calves, and foals, and some congenital diseases (such as glycogen branching enzyme deficiency in foals)

• Fatigue. Animals with primary severe respiratory disease can develop fatigue of the respiratory muscles (intercostal, diaphragm, accessory muscles of respiration), which can further impair ventilation

• Trauma. Fractured ribs can impair ventilation both because of the pain of breathing and because of mechanical disruption to respiration (flail chest).

General systemic states

Tachypnea can occur in a number of systemic states in which there is no lesion of the respiratory tract or nervous system. These include:

• Pain – such as in horses with colic

• Hyperthermia – as can occur with intense or strenuous exercise

• Acidosis – as a metabolic disturbance associated with any of a number of diseases but notably gastrointestinal disease that causes excessive loss of cationic electrolytes in feces.

Environmental causes

• Low inspired oxygen tension, such as in animals at high altitude

• Exposure to toxic gases.

Miscellaneous poisons

A number of poisons cause dyspnea as a prominent sign, but in most cases the pathogenesis has not been identified.

• Farm chemicals, including metaldehyde and dinitrophenols (probable mechanism is stimulation of respiratory center)

• Organophosphates and carbamates (probable mechanism is alteration of pulmonary epithelium), urea (probably effective as ammonia poisoning)

• Nicotine depressing the respiratory center

• Poisonous plants, including fast-death factor of algae, the weeds Albizia, Helenium, Eupatorium, Ipomoea, Taxus spp. and Laburnum and ironwood (Erythrophleum spp.); all appear to act at least in part by central stimulation.

ABNORMAL POSTURE

Animals with respiratory disease, and especially those in respiratory distress, often adopt an unusual posture and are rarely recumbent except in the terminal stages of the disease. Animals in severe respiratory distress will stand with the head and neck held low and extended. Animals, except horses, will often have open-mouthed breathing. Horses, except in extreme and unusual circumstances, are unable to breath through the mouth because of the anatomical arrangement of the soft palate, which effectively provides an airtight barrier between the oropharynx and nasopharynx. Cattle with severe respiratory distress and open-mouthed breathing will often drool large quantities of saliva – probably a consequence of decreased frequency of swallowing as the animal labors to breath.

The positioning of the legs is often abnormal. Severely affected animals, and those with pleuritic pain (horses or cattle with pleuritis) or severe respiratory distress, will usually stand with elbows (humeroradial joint) abducted. The animals are reluctant to move but when forced to do so can react violently. They are resistant to diagnostic or therapeutic interventions that interfere even transiently with their ability to breath.

NORMAL AND ABNORMAL BREATH SOUNDS

Auscultation of the lungs and air passages is the most critical of the physical examinations made of the respiratory system. The examination should be performed in as quiet an environment as possible, though it is often difficult to achieve a silent listening environment in large animal practice. The animal should be adequately restrained so that the examiner can concentrate on the lung sounds, and should not be sedated or anesthetized because of the depression in lung sounds that can occur in these instances. To be effective and diagnostically reliable, auscultation must be systematic. Both the upper and lower parts of the respiratory tract must be examined in every case. It is preferable to begin the examination by auscultating the larynx, trachea and the area of the tracheal bifurcation in order to assess the rate of air flow and the volume of air sound to be heard over the lungs.

RESPIRATORY NOISES

Respiration may be accompanied by audible noises that indicate certain normal or abnormal occurrences in the respiratory tract such as sneezing, snorting, stridor, stertor or snoring, wheezing, roaring, expiratory grunting and snuffling, bubbling and rattling sounds.

Sneezing is a sudden, involuntary, noisy expiration through the nasal cavities caused reflexly by irritation of the nasal mucosae. Sneezing occurs in rhinitis and obstruction of the nasal cavities, and digital manipulation and examination of the nasal mucosae.

Snorting is a forceful expiration of air through the nostrils as in a sneeze, but a snort is a voluntary act used by horses and cattle as a device to intimidate potential predators.

Stridor is an inspiratory stenotic sound originating from a reduction in the caliber of the larynx, as occurs in laryngeal edema and abscess.

Stertor or snoring is a deep guttural sound on inspiration originating from vibrations of pharyngeal mucosa. Snoring is often intermittent, depending on the animal’s posture. For example, a fat young bull will often snore when he is dozing half asleep, with his head hung down, but the snore will disappear when he is alert and his head is held up in a more normal position. Stertor can occur during expiration in horses with dorsal displacement of the soft palate.

Wheezing is a high-pitched sound made by air flowing through a narrow lumen, such as a stenotic or inflamed nasal cavity.

Roaring may occur during exercise and is caused by air passing through a larynx with a reduced lumen, e.g. laryngeal hemiplegia in horses.

Expiratory grunting is a clearly audible grunting noise synchronous with expiration. It is most common in cattle with diffuse pulmonary disease. A painful grunt may occur in painful diseases of the thorax such as fibrinous pleuritis and is unassociated with inspiration or expiration.

Snuffling, bubbling or rattling sounds may be audible over the trachea or base of the lungs when there is an accumulation of secretion, or exudate, in the nasal cavities, larynx or trachea. These are most clearly audible on inspiration.

COUGHING

A cough is an explosive expiration of air from the lungs. It is initiated by reflex stimulation of the cough center in the medulla oblongata by irritation of sensory receptors in one of various organs, especially the respiratory tract. The stimulus may originate in the pharynx, larynx, trachea or bronchi. Coughing may also be initiated by irritation of the esophagus, as in choking. The act of coughing consists of several stages:

The purpose of coughing is to remove the excess mucus, inflammatory products or foreign material from the respiratory tract distal to the larynx. Coughing indicates the existence of primary or secondary respiratory disease.

Coughing can be assessed according to several characteristics. Coughing is infrequent in the early stages of respiratory tract disease but can become frequent as the degree of inflammation in the larynx, trachea and bronchi becomes more severe. Assessment of the severity of coughing, at least in horses, requires prolonged observation (preferably for an hour).⁷ Coughing is a fairly specific but not very sensitive indicator of pulmonary inflammation.⁸ If coughing is detected then it is quite likely that the animal has inflammation of the airways, whereas failure to detect coughing does not reliably rule out the presence of clinically significant airway inflammation.⁸ The severity of coughing in horses is closely linked to the severity of inflammation and accumulation of mucopus in the airways.^7,8 Race horses that cough are 10 times more likely to have more than 20% neutrophils in a tracheal aspirate, and more than 100 times more likely to have more than 80% neutrophils.⁸ The frequency of coughing correlates well with maximal changes in pleural pressure, extent of mucus accumulation and proportion of neutrophils in bronchoalveolar lavage fluid of horses with heaves (recurrent airway obstruction).⁷ Coughing is therefore a specific indicator of the presence of respiratory inflammation.

The frequency of coughing is an indicator of the severity of lung disease in horses⁷ and presumably in other species. Horses that cough more than four times per hour have increased likelihood of mucus accumulation and higher pleural pressure changes during breathing than do horses that cough fewer than four times per hour.⁷

A cough cannot be induced in normal adult cattle and horses by manual manipulation of the larynx or trachea. If a cough can be induced in an adult horse by manual manipulation of the larynx or trachea, then this indicates airway inflammation and is a reason for further examination of the respiratory tract.

The most common causes of coughing in farm animals are due to diseases of the larynx, trachea, bronchi and lungs, which are presented under the headings of diseases of those parts of the respiratory tract later in this chapter.

CYANOSIS

Cyanosis is a bluish discoloration of the skin, conjunctivae and visible mucosae caused by an increase in the absolute amount of reduced hemoglobin in the blood. It can occur only when the hemoglobin concentration of the blood is normal or nearly so, and when there is incomplete oxygenation of the hemoglobin. Cyanosis is apparent when the concentration of deoxygenated hemoglobin in blood is greater than 5 g/dL (50 g/L).⁹ Cyanosis does not occur in anemic animals. The bluish discoloration should disappear when pressure is exerted on the skin or mucosa. In most cases, the oral mucous membranes are examined for evidence of cyanosis, although the skin of the pinna and the urogenital mucous membranes will suffice. Examination of vaginal mucosa is preferred in horses that have severe congestion of the oral and nasal mucosa as a result of disease affecting the head, such as cellulitis or bilateral jugular thrombophlebitis. Artificial lighting and skin pigmentation affect the ability to detect cyanosis.

Methemoglobinemia is accompanied by discoloration of the skin and mucosae but the color is more brown than blue and cannot be accurately described as cyanosis.

Cyanosis is classified as central or peripheral. Central cyanosis is present when arterial oxygen saturation is below normal with concentration of deoxygenated hemoglobin exceeding 4–5 g/dL. Peripheral cyanosis occurs when there is localized desaturation of blood despite arterial oxygen saturation being normal. This usually occurs because there is diminished blood flow to tissue, with a resulting increase in oxygen extraction by the ischemic tissues and low end-capillary and venous hemoglobin saturation.

Central diseases include:

Peripheral causes of cyanosis include:

Central cyanosis is characterized by decreased arterial oxygen saturation due to right-to-left shunting of blood or impaired pulmonary function. Central cyanosis due to congenital heart disease or pulmonary disease characteristically worsens during exercise. Central cyanosis usually becomes apparent at a mean capillary concentration of 4–5 g/dL reduced hemoglobin (or 0.5 g/dL methemoglobin). Since it is the absolute quantity of reduced hemoglobin in the blood that is responsible for cyanosis, the higher the total hemoglobin content the greater the tendency toward cyanosis. Thus cyanosis is detectable in patients with marked polycythemia at higher levels of arterial oxygen saturation than in patients with normal hematocrit values, and cyanosis may be absent in patients with anemia despite marked arterial desaturation. Patients with congenital heart disease often have a history of cyanosis that is intensified during exertion because of the lower saturation of blood returning to the right side of the heart and the augmented right-to-left shunt. The inspiration of pure oxygen (100% F_io₂) will not resolve central cyanosis when a right-to-left shunt is present, but can resolve when primary lung disease or polycythemia is causing the cyanosis.

Peripheral cyanosis is caused by obstruction of blood flow to an area. This can occur as a result of arterial or venous obstruction, although it is usually more severe when arterial blood flow is obstructed. Obstruction of arterial blood flow also causes the limb to be cold and muscle and nerve function in the ischemic area to be impaired. Cyanosis can also occur as a result of cutaneous vasoconstriction due to low cardiac output or exposure to cold air or water. It usually indicates stasis of blood flow in the periphery. If peripheral cyanosis is localized to an extremity, arterial or venous obstruction should be suspected. Peripheral cyanosis due to vasoconstriction is usually relieved by warming the affected area.

Heart failure can cause cyanosis that is restricted to the extremities, probably because of reduced blood flow to extremities during this disease and the consequent markedly lower end-capillary oxygen content. Blood in the venous end of the capillaries, and in the venous bed draining these tissues, is therefore deoxygenated and cyanosis is observed. While this type of cyanosis has a peripheral distribution, its underlying cause is central.

NASAL DISCHARGE

Excessive or abnormal nasal discharge is usually an indication of respiratory tract disease. Nasal discharges are common in all the farm animal species. Cattle can remove some or all of the nasal discharge by licking with their tongue, while horses do not remove any.

EPISTAXIS AND HEMOPTYSIS

Pulmonary hemorrhage, particularly in the horse, may be manifested as epistaxis. Pulmonary hemorrhage in cattle is commonly manifested as hemoptysis and epistaxis. These are described in more detail later in this chapter.

A small amount of serosanguineous fluid in the nostrils, as occurs in equine infectious anemia and infectious equine pneumonia, does not represent epistaxis, which must also be differentiated from the passage of blood-stained froth caused by acute pulmonary edema. In this instance the bubbles in the froth are very small in size and passage of the froth is accompanied by severe dyspnea, coughing and auscultatory evidence of pulmonary edema.

THORACIC PAIN

Spontaneous pain, evidenced by grunting with each respiratory cycle, usually indicates pleural pain, such as from a fractured rib, torn intercostal muscle or traumatic injury, including hematoma of the pleura, or pleurisy. A similar grunt may be obtained by deep palpation or gentle thumping over the affected area of the thoracic wall, with a closed fist or a percussion hammer. Pain due to a chronic deep-seated lesion cannot be detected in this way. The use of a pole under the sternum, as described under traumatic reticuloperitonitis, provides a useful alternative.

AUSCULTATION AND PERCUSSION

The techniques of auscultation and percussion used in examination of the thorax are discussed in Chapter 1 and references on percussion of the thorax are available.^14,15 Percussion of the thorax is a useful means of determining lung margins and therefore of detecting the presence of overinflation, as occurs with heaves in horses,¹⁶ or areas of consolidation. Consolidation is evident as a loss of resonance, and detection of this abnormality can reveal the presence of excessive pleural fluid or pulmonary consolidation. There is excellent agreement in the assessment of lung margins determined by percussion and by ultrasonographic examination.¹⁶ Percussion is therefore a valuable diagnostic tool, especially when ultrasonographic examination is not available.

ENDOSCOPIC EXAMINATION OF THE AIRWAYS (RHINOLARYNGOSCOPY, TRACHEOBRONCHOSCOPY)

RADIOGRAPHY

Radiography of the head, neck and thorax is valuable in the diagnosis of diseases of the respiratory tract of animals. Examination is hindered by the large size of adult horses and cattle through the need for specialized, high-capacity equipment for obtaining radiographs, and the need for adequate restraint. Radiographic examination of adult animals in the field using portable radiographic units is very limited. However, large practices with fixed radiographic units capable of generating sufficient voltage and amperage can obtain diagnostic radiographs of the thorax of adult horses and cattle. Diagnostic films of smaller animals, including adult sheep and goats and foals and calves, can be obtained using portable units capable of generating 80–100 kVp and 15–20 mA.

Examination of the thorax of large animals is restricted to lateral radiographs because the large amount of tissue prevents adequate exposure for ventrodorsal views. Multiple films are required for complete examination of the thorax, and the exposure needed for optimal quality films varies among anatomical sites. Localization of focal lesions can be achieved by examining sets of radiographs that include images collected with the horse or cow standing first with one side to the plate and then with the other side toward the plate. The lesion will appear larger in views obtained with the lesion closer to the source of X-rays.

Radiographs of calves and foals can be recorded with them standing or recumbent. Images obtained with the foal or calf in lateral recumbency with the forelimbs pulled forward permit optimal examination of the cranial thorax. However, calves or foals that are recumbent for prolonged periods of time (e.g. > 30 min) can develop atelectasis of the down lung that can mimic pneumonia radiographically. Ventrodorsal views assist with localizing lesions in foals and calves. Radiographic evidence of lung disease is common in ill neonatal foals (74% having such lesions in one study),²⁵ and is not related to clinical evidence of respiratory disease or dyspnea.²⁶ The characteristics of lung lesions detected in neonatal foals are associated with likelihood of survival. Guidelines for recognition of pulmonary patterns in foals have been proposed (Table 10.2)²⁶ and these guidelines are likely to be useful aids for interpretation and description of pulmonary patterns in neonates of other species.

Table 10.2 Guidelines for radiographic pulmonary pattern recognition in foals²⁶

Radiography can assist in the recognition and differentiation of atelectasis and consolidation, interstitial and exudative pneumonias, the alveolar pattern of pulmonary disease, neoplasms, pleural effusions, pneumothorax, hydropericardium and space-occupying lesions of the thorax. Cardiomegaly, abnormalities of the cranial mediastinum, fractures of ribs and diaphragmatic hernia can also be detected.

Many pulmonary diseases do not have lesions that are readily detected on radiographic examination. Failure to detect abnormalities on radiographic examination of the thorax does not eliminate pulmonary disease. Furthermore, radiographically detectable signs of lung disease can persist after the animal has clinical and clinicopathological signs of recovery or improvement.

Bronchography utilizing contrast agents is of value in determining the patency of the trachea and bronchi, but general anesthesia is required to overcome the coughing stimulated by the passage of the tracheal catheter. Using a fluoroscope to determine the location of the catheter tip, the contrast agent can be deposited in each dependent lobe in turn. This technique is used infrequently. Computed tomographic (CT) examination of the lung is very sensitive and specific for lung disease in companion animals and is technically feasible in calves, foals and small ruminants. The technique is useful in the diagnosis of mediastinal disease in foals.²⁷

Radiographic examination of the trachea can reveal the presence of abnormalities in shape, such as occur with tracheal collapse, or the presence of foreign bodies or exudate.

Radiographic examination of the head can identify diseases of the paranasal sinuses, ethmoids and pharynx. Radiographic examination is useful in defining diseases of the guttural pouches and in detecting retropharyngeal abscesses or abnormalities, such as the presence of foreign bodies. The CT anatomy of the head of horses and foals has been described.^28-30 CT imaging of the nasal cavities and paranasal sinuses of horses is useful in the detection of diseases of these structures,^31,32 and of the teeth,³³ pharynx, larynx and guttural pouches.³⁴ The technique is technically feasible in ruminants and pigs, although there are few reports of its use in these species.³⁵

Magnetic resonance (MR) imaging is useful in diagnosis of diseases of the head, and the anatomy as visualized on MR imaging of the head of horses has been reported.³⁶ Unfortunately, the lack of units suitable for examination of large animals precludes routine use of this imaging modality.

SCINTIGRAPHY (NUCLEAR IMAGING)

The basis of pulmonary scintigraphy is detection at the body surface of radiation emitted from the lungs after injection or inhalation of radioactive substances.³⁷ The technique has been described in both horses and calves.^37,38 The technique has limited diagnostic usefulness in large animals because of the need for availability of appropriate isotopes and detection equipment. Furthermore, the large size of adult cattle and horses limits the sensitivity of the technique. The technique has been used to determine the distribution of pharmaceuticals administered by aerosolization and the presence of ventilation/perfusion mismatches. Alveolar clearance can be detected using scintigraphic examination. Currently pulmonary scintigraphy is largely a research tool.

ULTRASONOGRAPHY

Ultrasonographic examination of the thorax of farm animals and horses is a very useful diagnostic tool. Ultrasonographic examination of the thorax provides diagnostic information that is not obtained by radiographic examination. The widespread availability of portable ultrasound units and the ability to image parts of the thorax using ultrasound probes intended for examination of the reproductive tract of mares and cows makes this a potentially valuable diagnostic aid for both field and hospital-based practitioners. Furthermore, the absence of radiation exposure and the ‘real-time’ nature of images obtained by ultrasonography aid in frequent assessment and monitoring of abnormalities and performance of diagnostic or therapeutic procedures such as thoracocentesis or aspiration of masses.

There are limitations to imaging imposed by aerated lung and the bones of the ribcage. Examination of the thorax is limited by the presence of ribs and aerated lungs because the sound waves used to create ultrasound images are reflected from these surfaces. Ultrasonography cannot reveal lesions of the lungs that are not confluent with the visceral pleura. Imaging windows are restricted to the intercostal spaces but this impediment can be overcome by scanning through adjacent intercostal spaces and angling of the ultrasound beam.

Ultrasonographic examination of the thorax should be performed in a consistent manner that ensures thorough examination of the thorax. Preferences for the pattern of examination differ somewhat among examiners, but one common and successful technique is to scan each intercostal space from dorsal to ventral starting at the 17th intercostal space in horses and the 12th intercostal space in cattle. The ultrasound probe is slowly moved from dorsal to ventral while the examiner studies the images. When one scanning of one intercostal space is completed, the probe is moved to the most dorsal aspect of the next intercostal space and the examination is repeated. Each side of the chest is examined in this manner. This consistent and thorough examination ensures that no important or localized abnormalities are missed. The examination is performed in adult horses and cattle with the animal standing. The rostral thorax is scanned by pulling the ipsilateral forelimb forward. This is more readily achieved in horses than in cattle. Thorough examination of the rostral thorax might require placing the animal in lateral recumbency. Calves and foals can be examined either standing or in lateral recumbency.

Ultrasound examination of the thorax is particularly useful for detecting diseases of the pleura, pleural space or lung surface. This is in addition to the well-documented utility of ultrasonographic examination of the heart and great vessels (see Ch. 8). The normal ultrasonographic anatomy of the thorax of cattle, horses and calves has been determined.^39-41 The following is a partial list of disorders or abnormalities detectable by percutaneous ultrasonographic examination of the thorax of farm animals or horses (excluding cardiac abnormalities):

Ultrasonographic examination is more sensitive and specific than radiographic examination in detecting the presence of pleural fluid⁴² and is particularly useful in the diagnosis and management of pleuritis in horses and cattle^43,44 and pneumonia in calves.⁴⁵ The extent of pulmonary lesions detected at necropsy correlates closely with the results of ultrasonographic examination of calves with pasteurellosis.⁴⁶ Ultrasonographic examination is useful in diagnosis of thoracic diseases of cattle.⁴⁷ Ultrasonography can identify pulmonary lesions in horses with infectious viral pneumonia⁴⁸ and is a viable alternative, though not as sensitive, to radiology in the evaluation of foals with Rhodococcus equi pneumonia.⁴⁹ Ultrasonography is very useful in identifying the presence of pleural fluid and guiding thoracocentesis to sample and drain this fluid.

LABORATORY EVALUATION OF RESPIRATORY SECRETIONS

TRACHEOBRONCHIAL SECRETIONS

The collection and evaluation of tracheobronchial secretions is a useful method for assessing lower airway disease and is widely used in the determination of the etiology of infectious pneumonia (viral, mycoplasmal, fungal, and parasitic) or the severity of disease (bronchoalveolar lavage fluid cytology in horses with heaves, exercise-induced pulmonary hemorrhage in athletic horses). It is also used as a tool in evaluating respiratory health of intensively housed animals, such as in piggeries.⁵³ Cytological examination of recovered fluid can provide valuable information about the severity, extent and etiology of disease of the lower airway. There are two methods of sampling tracheobronchial secretions – aspiration of tracheal fluid or lavage of bronchioles and distal airways. Each sampling method yields fluid of differing characteristics and source and interpretation of the results of examination of these fluids depends on their source and the method of collection.

Comparison of tracheal aspirates and bronchoalveolar lavage fluid

Examination of tracheal aspirates and bronchoalveolar lavage fluid yields different, but often complementary, information about the lower respiratory tract. The differences between tracheal aspirates and bronchoalveolar lavage fluid arise because cell populations, and types of cell, differ markedly among segments of airways. There is no correlation between cytological features of tracheal aspirates and bronchoalveolar lavage fluid of horses, and this is probably the case in other species.⁵⁴ Tracheal aspirates are representative of cell and bacterial populations of the large conducting airways (trachea and mainstem bronchi), which can originate in both the large and small conducting airways and the alveoli.⁵⁴ Secretions of more distal airways can be modified during rostrad movement, such that fluid in a tracheal aspirate is not representative of processes deeper within the lung. Furthermore, disease localized to one region of the lung can alter tracheal fluid. Examination of tracheal aspirates is useful for detecting inflammation of the large airways and for isolation of microorganisms causing disease in these structures. There is no good evidence that findings on examination of tracheal aspirates correlate with abnormalities in pulmonary function, and tracheal aspirates do not accurately reflect lesions in the lungs of horses.⁵⁵

Bronchoalveolar lavage is useful for sampling secretions in the more distal airways. It provides a sample of secretions that have not been contaminated by upper respiratory tract organisms or secretions before collection and the sample is therefore assumed to be more representative of small airway and, to a lesser extent, pulmonary parenchymal and alveolar secretions or exudates. Bronchoalveolar lavage is useful in the detection of widespread lung disease but not necessarily in the detection of localized disease. Tracheal aspirates, because they in theory represent a composite sample of secretions from all regions of the lung, are likely to be more sensitive in detecting focal disease, such as a pulmonary abscess. Bronchoalveolar lavage fluid composition correlates well with pulmonary function in horses.^56,57

There is little agreement in cytological examination of tracheal aspirates and bronchoalveolar lavage fluid of sick and healthy horses, and this difference probably exists in other species. Typically, the proportion of cells that are neutrophils is much higher in tracheal aspirates than in bronchoalveolar lavage fluid of both horses with heaves and normal horses.^54,58 Mast cells are detected more frequently, and eosinophils less frequently, in bronchoalveolar lavage fluid than in tracheal fluid of normal horses.⁵⁹

Tracheal aspirates

Indications for collection of tracheal aspirates include the need for microbiological and cytological assessment of tracheal fluids. The primary indication is collection of samples for microbiological diagnosis of infectious respiratory disease. Other indications include detection and characterization of inflammation of the conducting airways. Contraindications include severe respiratory distress, although this is not an absolute contraindication, inability to adequately restrain the animal, and severe, spontaneous coughing. Percutaneous tracheal aspirate collection performed in animals with severe coughing can result in development of severe subcutaneous emphysema as a result of the high intratracheal pressures associated with the early phase of coughing. Most animals in which percutaneous tracheal aspirates are collected subsequently have radiographic evidence of pneumomediastinum.

Tracheal aspirates can be collected either by percutaneous puncture of the trachea or through an endoscope passed through the upper airways. The advantage of percutaneous collection of tracheal aspirates is that there is minimal risk of contamination of the sample by upper respiratory tract or oropharyngeal secretions. Microbiological examination of the samples is therefore likely to accurately reflect microbes present in tracheal fluid. Collection of tracheal aspirates through an endoscope markedly increases the risk of contamination of the sample with oropharyngeal fluids, and compromises the diagnostic utility of culture of the sample. This disadvantage is partially alleviated by the use of guarded catheters inserted through the endoscope.^60,61 The disadvantage of percutaneous collection of tracheal fluid is that it is invasive and there is a risk of localized cellulitis and emphysema at the site of puncture. Endoscopic collection is relatively noninvasive and readily accomplished.⁵ We prefer use of the percutaneous method when accurate microbiological assessment of samples is desired.

Percutaneous transtracheal aspiration

This is a practical method that has been used extensively in the horse⁶⁰ and is adaptable to cattle,⁶² sheep and goats. For the horse, a 60 cm no. 240–280 polyethylene tube is passed through a 9–14-gauge needle inserted into the trachea between two rings. Commercially prepared kits for performing tracheal aspirates in horses are available that include all catheters and needles required. An alternative to polyethylene tubing is to use an 8–10 French male dog urinary catheter inserted through an appropriately sized cannula. The site for insertion of the needle or cannula is at the junction of the proximal and middle one-thirds of the ventral neck. The horse is usually sedated prior to insertion of the needle or cannula. The skin site is prepared aseptically and a short stab incision is made after the area has been anesthetized. The cannula is removed to avoid cutting the tube and the tube is pushed in as far as the thoracic inlet. Fluid typically pools in the trachea at the thoracic inlet in horses with lung disease (the tracheal lake or pool) and it is this fluid that is aspirated. Thirty to 50 mL of sterile saline (not bacteriostatic saline) is rapidly infused. The catheter or tubing should be rotated until tension is felt on aspiration by a syringe. Fluid is aspirated and submitted for cytological, microbiological or other examination.

Complications such as subcutaneous emphysema, pneumomediastinum and cellulitis can occur, which necessitates care and asepsis during the procedure. Sudden movement of the cannula during insertion of the tubing may cause part of the tube to be cut off and to fall into the bronchi, but without exception this is immediately coughed up through the nose or mouth.

Endoscopic sampling of tracheal secretions

The flexible fiberoptic endoscope can be used to obtain tracheal lavage samples and at the same time visualize the state of the airways.^15,60 The process is as for rhinolaryngoscopic examination with the addition of passage of a catheter through the biopsy port of the endoscope. Tracheal fluid is then visualized and aspirated through the catheter. The clinical advantages of the endoscopic collection include noninvasiveness, visual inspection of the airways, guidance of the catheter, and speed.⁶⁰ The use of an endoscope with a guarded tracheal swab minimizes contamination by oropharyngeal secretions but does not eliminate it.^5,61

Assessment of results

Microbiological examination can yield any one or more of a variety of bacteria, depending on the species examined, the animal’s age and its clinical condition. Tracheal aspirates of normal animals rarely yield any bacterial growth on culture. Growth of unusual organisms or known oropharyngeal commensal bacteria from samples obtained by endoscopic examination should not be given undue clinical significance as they probably result from contamination of the tracheal aspirate during collection. Pseudomonas spp. and anaerobes isolated from tracheal aspirates collected by endoscopy are almost always contaminants and of no clinical significance.⁵ The extent of contamination of tracheal aspirate samples by oropharyngeal bacteria can be estimated from the number of squamous epithelial cells in the sample.⁵⁹ There is an apparent approximate linear relationship between the number of squamous cells per milliliter of fluid and the number of colony-forming bacterial units in tracheal aspirates. Samples containing over approximately 10 squamous epithelial cells per milliliter of tracheal aspirate had markedly greater bacterial contamination.⁵⁹ Examination of Gram-stained smears of tracheal fluid is specific but not very sensitive for detection of bacteria, compared with culture.⁶³ In other words, if examination of a Gram-stained smear of tracheal fluid reveals bacteria, then the sample is likely to yield bacteria on culture, whereas failure to detect bacteria predicts poorly the likelihood of growth of bacteria on culture of the sample. This indicates that examination of Gram-stained samples of tracheal fluid does not reliably predict bacterial isolation, and if an infectious etiology is suspected the fluid should be cultured. Results of the microbiological examination of the tracheal fluid should be consistent with the animal’s clinical condition and expected isolates.

Cytological examination of tracheal fluid is an important diagnostic tool. Various stains are available to aid identification of cell types and numbers in tracheal aspirates. Neutrophils, macrophages, lymphocytes and epithelial cells are readily identified on the basis of their classical morphology and staining using fast Romanowsky stain (Diff-Quik®), but this stain is not suitable for identifying mast cells in equine tracheal fluid and probably that of other species.⁵⁹ Leishman’s stain is useful to identify mast cells.⁵⁹ Clinically normal horses typically have fewer than 20–30% of cells as neutrophils with the majority of remaining cells being macrophages, lymphocytes and epithelial cells.^54,61 Animals with inflammation of the airways typically have increased cell counts and proportion of neutrophils, and large amounts of mucus. Horses with inflammatory airway disease such as heaves typically have more than 20% of the cells as neutrophils (see Heaves, below), and those with infectious pneumonia often have 50–90% of cells as neutrophils. The presence of eosinophils is considered abnormal and is consistent with parasite migration (Parascaris equorum in foals, Dictyocaulus viviparus in calves). The presence of hemosiderin-laden macrophages is evidence of prior pulmonary hemorrhage.⁶⁴

Bronchoalveolar lavage

Bronchoalveolar lavage provides a sample of secretions and cells of the distal airways and alveoli, referred to as bronchoalveolar lavage fluid. It is a widely used procedure in horses and, to a lesser extent, cattle and calves,^65-67 sheep⁶⁸ and pigs.^53,69 Analyses performed on bronchoalveolar lavage fluid include measurement of cell number and type, culture (usually in pigs and cattle) and analysis of immune proteins and surfactant. It is a relatively noninvasive procedure that allows cytological and biochemical evaluations of the lower airways and alveoli, which are useful diagnostic aids when evaluating animals with lung disease. While fiberoptic bronchoscopy and tracheal aspirates permit assessment of the major bronchi and upper airways, bronchoalveolar lavage offers an extension of the diagnostic potential by sampling the terminal airways and alveolar spaces.

The primary indication for collection of bronchoalveolar lavage fluid is acute or chronic lung disease. This includes both infectious and noninfectious diseases, although interpretation of samples collected by passage of the collection tube through the nostrils or mouth is complicated by the inevitable contamination of the sample by oropharyngeal commensal bacteria. Despite this shortcoming, the technique has been used to detect pneumonia associated with Mycoplasma sp. in cattle.⁷⁰ Contraindications are few, with respiratory distress being an obvious one. Complications of bronchoalveolar lavage are also few, and include a mild neutrophilia in lavaged sections of lungs and changes in phagocytic function of pulmonary macrophages, and microbial content, for several days after the procedure.^31,71

A shortcoming of bronchoalveolar lavage is that it lavages only a small region of the lung, with the risk that focal lung disease is not detected. This is best exemplified in pneumonia in horses, in which bronchoalveolar lavage fluid from pneumonic horses can contain large numbers and a high proportion of neutrophils or can be normal, depending on the area of lung lavaged. Therefore, the bronchoalveolar lavage procedure is a very specific but not very sensitive test for pneumonia in horses.⁷² Abnormal lavage fluid is helpful diagnostically, whereas normal results do not exclude the presence of foci of pulmonary disease. The lavage samples may be normal in horses affected with pneumonia or pleuropneumonia and, because of these false-negative results, this is not the best diagnostic technique to evaluate a horse with pneumonia.¹³ In contrast, the tracheobronchial aspirates are more sensitive and most horses with pneumonia have cytological abnormalities.¹³

Endoscopic bronchoalveolar lavage

Endoscopic bronchoalveolar lavage has the advantage of permitting visual examination of the airways during the procedure and selection of the region of the lung to be lavaged. This technique does require access to sophisticated endoscopic equipment. The technique described below for horses can be modified for use in other species.⁷³

Horses for bronchoalveolar lavage should be appropriately restrained. Sedation is usually essential and is achieved by administration of alpha-2-agonists. Coadministration of narcotics is recommended by some authorities to reduce the frequency and severity of coughing. Butorphanol tartrate 10 mg for a 400 kg horse is recommended, although this drug is not as effective as intratracheal lidocaine at reducing the frequency or severity of coughing when combined with detomidine for collection of bronchoalveolar lavage fluid.⁷⁴ Effective suppression of coughing during collection of bronchoalveolar lavage fluid can be achieved by instillation of lidocaine (60 mL of a 0.7% solution – made by diluting 20 mL of 2% lidocaine solution by addition of 40 mL of isotonic saline). The lidocaine solution is administered as the endoscope enters the rostral trachea. A twitch can be applied to the nares. The endoscope must be at least 2 m in length and the external diameter should be 10–15 mm. Endoscopes of 10 mm diameter will pass to about the fifth-generation bronchi, whereas endoscopes of larger diameter will not pass quite as far into the lung. The endoscope is passed until it wedges and then 300 mL of warmed (to reduced bronchospasm) isotonic saline is introduced in 5 × 60 mL aliquots. Air is infused after the last aliquot to ensure that all fluid is instilled. After the horse has taken between one and three breaths the fluid is withdrawn and the aliquots are mixed. There is no difference in the cytological composition of the first and subsequent aliquots.⁷⁵

Blind bronchoalveolar lavage

Commercial bronchoalveolar lavage tubes are available for use in horses, and are suitable for use in adult cattle and calves.⁶⁷ The tubes are made of silicone and are therefore considerably more pliable than stomach tubes (which are not suitable for this procedure). The tubes are 2 m in length and have an external diameter of about 8 mm. The horse is restrained and sedated as for endoscopic bronchoalveolar lavage and the tube is passed through one nostril into the trachea. The tube is then advanced until it wedges, evident as no further insertion of the tube with mild pressure. Continued vigorous attempts to pass the tube can result in the tube flexing in the pharynx and a loop of the tube entering the mouth. After the tube wedges, the cuff on the tube is inflated to prevent leakage of fluid around it, 300 mL of warm isotonic saline is instilled, the tube is flushed with air and fluid is aspirated. The fluid should be foamy and, if cell counts are high, slightly cloudy.

Bronchoalveolar lavage can be performed in conscious sheep by insertion of 1.7 mm external diameter polyethylene tubing through a cannula inserted percutaneously in the trachea.⁶⁸ The tubing is inserted until resistance is detected (about 40–45 cm in an adult sheep) and the lung is lavaged with 30 mL of sterile isotonic saline.

Laboratory assessment of tracheobronchial secretions

A problem with comparison of cell counts of bronchoalveolar lavage fluid reported by different authors is the use of inconsistent quantities of fluid to perform the lavage. The use of different volumes alters the extent of dilution of the fluid. There is a need for uniformity in technique.⁷⁶ An approach to this problem has been to measure substances in the bronchoalveolar lavage fluid that can provide an indication of the extent of dilution of the sample. Both endogenous (urea, albumin) and exogenous (inulin, methylene blue) markers have been used. Dilution factors using urea concentration in plasma and in bronchoalveolar lavage fluid appear to be useful.⁷⁷ The assumption is that urea concentrations in bronchial and alveolar secretions will be identical to that in plasma. The formula for correcting for dilution that occurs during collection of bronchoalveolar lavage fluid is:

Dilution factor = Urea concentration in bronchoalveolar lavage fluid/Urea concentration in plasma,

where urea concentration in bronchoalveolar lavage fluid and in plasma is expressed in the same units. The volume of the pulmonary epithelial lining fluid can then be calculated:

Pulmonary epithelial lining fluid volume = dilution factor × volume of bronchoalveolar lavage fluid.

Samples for cytology are submitted for preparation involving centrifugation of the sample to concentrate cells for preparation of slides for staining and microscopic examination. At least for samples from horses, examination of smears made directly from the sample, without centrifugation, is diagnostically useful.⁷⁸ As for tracheal fluid, the proportion of mast cells in equine bronchoalveolar lavage fluid is underestimated if cells are stained with fast Romanowsky stain (Diff-Quik®).⁷⁹

Diagnostic value

The aspirates from normal animals contain ciliated columnar epithelial cells, mononuclear cells and a few neutrophils with some mucus. The concentration of the cells depends on the volume of fluid infused and the disease status of the animal. Representative values for various species are listed in Table 10.3. The general pattern is that animals with inflammatory airway disease, either infectious or noninfectious, have a higher proportion of neutrophils than do disease-free animals. However, ranges of normal values vary considerably depending on the species, the age of the animal and its management (primarily housing conditions). Care should be taken not to overinterpret findings on examination of tracheal aspirates or bronchoalveolar lavage fluid. While there is good correlation between microbiological results and cell counts in bronchoalveolar lavage fluid of calves with pneumonia⁶⁵ and Thoroughbred race horses with inflammatory airway disease,⁸⁰ this association might not hold for all diseases or species.

Table 10.3 Representative results of cytology of bronchoalveolar lavage fluid of cattle, sheep, pigs, and horses

Thoracocentesis (pleurocentesis)

Paracentesis of the pleural cavity is of value when the presence of pleural fluid is suspected and, in the absence of ultrasonographic examination, needs to be confirmed, and when sampling of pleural fluid for cytological and bacteriological examination is indicated. The primary indication for sampling pleural fluid is the presence of excess pleural fluid. Sampling of pleural fluid is usually accompanied by therapeutic drainage, in which case the cannula used for sampling is larger than if only collection of pleural fluid is desired. Contraindications are minimal, especially if the procedure can be performed under ultrasonographic guidance. The principal contraindication is the inability to restrain an unruly animal, as this increases the risk of laceration of the lung or a coronary vessel, or cardiac puncture. Complications include hemorrhage from lacerated intercostal or pleural vessels, pneumothorax secondary to laceration of the lung or introduction of air through the cannula, cardiac puncture and sudden death, irritation of the myocardium and ventricular arrhythmia (premature ventricular contractions), or coronary artery laceration and subsequent cardiac tamponade and death. There is a risk of cellulitis at the site of centesis, especially if indwelling cannulas are maintained for more than a day.

The procedure is performed with the animal standing. Sedation or systemic analgesia is usually not needed, unless it is medically indicated or the animal is not easily restrained. The equipment for sampling of pleural fluid from adult horses or cattle is a blunt 10–15 cm cannula of approximately 3 mm diameter (such as a bovine teat cannula) or a 7.5 cm spinal needle. The blunt-tipped cannula is preferred because use of it reduces the risk of laceration of vital structures. A three-way stopcock or similar device should be attached to the hub of the needle or cannula and closed to prevent aspiration of air when the pleural cavity is entered. The site for centesis is best identified by ultrasonographic examination of the thorax or, if that is not available, by percussion and auscultation of the chest to identify the fluid level. A commonly used site is the seventh intercostal space on the left side and the sixth intercostal space on the right side. The skin should be clipped of hair and aseptically prepared. The region can be anesthetized with approximately 10 mL of 2% lidocaine, mepiricaine or similar product. The cannula or needle should be introduced over the rib and then directed cranial to the rib (the intercostal vessels and nerves course along the caudal edge of the rib). If a cannula is used then a slight ‘popping’ sensation is felt as the cannula perforates the parietal pleura. A syringe is attached to the cannula or needle and fluid is aspirated from the pleural space.

Collected fluid should be examined visually. Normal pleural fluid, which is present in small quantities in normal animals, is clear and slightly yellow. Abnormal fluid can be bloody, thick and yellow, suggestive of purulent material, or flocculent. The material should be smelled – a foul odor is usually present when the pleural fluid is infected by anaerobic bacteria and is a sign of a poor prognosis. Cytological examination should be performed, including white cell count and measurement of total protein concentration. Ancillary measurements on pleural fluid include pH, Pco₂, Po₂, bicarbonate, glucose and lactate. Sterile pleural fluid has a pH, Po₂ and Pco₂ and lactate, glucose and bicarbonate concentrations similar to those of venous blood.⁸¹ Infected pleural fluid is acidic, hypercarbic and has an increased concentration of lactate and decreased concentrations of bicarbonate and glucose compared to venous blood.⁸¹ Pleural fluid should be cultured for aerobic and anaerobic bacteria and mycoplasmas. Antimicrobial susceptibility should be determined for isolated organisms. Fungal cultures are rarely indicated.

Ultrasound-guided needle puncture of a suspected lung abscess to determine the species of bacteria present is sometimes practiced but there is the risk that infection will be spread to the pleura by this technique. This technique is not recommended as a routine procedure as microbiological examination of tracheal aspirates will probably yield the offending bacteria.

PULMONARY FUNCTION TESTS

Pulmonary function tests provide quantitative assessment of pulmonary ventilatory function through measurement of expired and inspired gas volumes, intrathoracic pressures and derivations of these variables – sometimes referred to as pulmonary mechanics. The techniques are widely used in research into pulmonary diseases, especially heaves, in horses, and have been adapted for use in ruminants.⁸² A relatively simple assessment of pulmonary function is measurement of pleural pressure changes during respiration. This can be achieved by either insertion of a blunt cannula through the intercostal space or passage of a balloon catheter into the thoracic esophagus. The pressure changes during respiration are then recorded and the maximal pressure change between inspiration and expiration is calculated. The pressure change is closely correlated with airway resistance to airflow and is an excellent indicator of the severity of bronchoconstrictive diseases.

More complex measurements are made by application of an airtight face mask containing a flow meter to the animal. Combined with measures of airway pressure, air flow during tidal breathing yields measures of tidal volume, minute volume, respiratory rate, pulmonary resistance and pulmonary dynamic compliance. Measurements made with the animal at rest are relatively insensitive to small changes in pulmonary function and the sensitivity of these tests to detect heaves is low.⁵⁷ The sensitivity of changes in maximal pleural pressure and resistance of the lower airways are 44% and 22%, respectively.⁵⁷ The sensitivity of the test can be increased by measuring these variables during exercise. Measurement of pulmonary mechanics in horses with heaves is reproducible over both short (hours) and long (months) periods of time, indicating the usefulness of these techniques for monitoring of disease progression and response to therapy.⁸³

Measurement of flow–volume loops has been performed for both stationary and exercising horses.^84,85 A number of variables are derived from these measures and used as indicators of pulmonary function. However, the large variability in these measures in stationary horses (16–32%) severely limits the utility of this test to detect mild or subclinical respiratory disease. Similarly, flow–volume loops in exercising horses with obstructive lung disease of moderate severity do not differ markedly from those of the same horses when they do not have lung disease. Flow–volume loops have limited use in evaluation of lung function in animals.

Other tests of pulmonary function include the nitrogen dilution test and the single-breath diagram for CO₂. For the nitrogen dilution test concentrations of nitrogen in exhaled air are measured while the animal breathes 100% oxygen. A number of variables are calculated from the decay curve of nitrogen concentration in exhaled air, including the functional residual capacity.⁸⁶ There are clinically significant differences between animals with normal respiratory function and those with pulmonary disease. However, this test is not readily adapted for routine clinical use. Volumetric capnography is the graphic examination of expired breath CO₂ concentrations versus expired volume to create a single-breath diagram for CO₂.⁸⁷ The results are divided into phase I, which represents the relatively carbon-dioxide-free air from the proximal or orad conducting airways, phase II, which is the transitional phase, and phase III, which is the carbon-dioxide-rich air from the alveoli. Measures of pulmonary function obtained include estimates of dead space ratio, physiological dead space volume and alveolar efficiency.⁸⁷ The clinical utility of this test and its ability to detect mild or subclinical disease in animals have not been demonstrated.

Impulse oscillometry offers the potential of being a potentially clinically useful test of respiratory function in both horses and cattle.^46,88,89 The test measures impedance of the respiratory system and provides estimates of respiratory resistance and reactance.⁴⁶ The technique has the advantage of being more sensitive to changes in pulmonary function than measurement of pleural pressure changes,⁹⁰ is minimally influenced by respiratory rate and tidal volume⁹¹ and is relatively easier to perform than more complex measures of respiratory mechanics. The test involves fitting an airtight facemask containing a pneumotachograph for measurement of respiratory volumes and tubing to a horse. The tubing is attached to a loudspeaker, which is used to generated square-wave signals containing harmonics between 0 and 10 Hz. Information from the system is analyzed using a computer program and indices of pulmonary resistance and reactance are determined. The forced oscillation technique in feedlot cattle with naturally occurring shipping fever indicates the presence of a large increase in pulmonary resistance and a decrease in dynamic compliance with obstructive lung disease located mainly at the level of large airways but also in small airways.⁹² The clinical utility of the technique remains to be determined.

The sensitivity of these tests can be increased by provocative tests in which animals are administered agents, such as histamine or methacholine, that cause bronchoconstriction in animals with reactive airways.⁹⁰

Measurement of forced expiratory flow–volume curves and forced vital capacity in horses is a sensitive indicator of bronchoconstriction.^57,93 The test involves the heavily sedated horse having a nasotracheal tube inserted. The nasotracheal tube is then attached to a large vacuum reservoir and a valve is opened abruptly. The maximum rate of forced expiratory airflow is measured and various variables indicative of pulmonary function are calculated, including forced expiratory volume in one second (FEV₁).⁹³ The clinical utility of this test of pulmonary function is limited by the extensive instrumentation of the animal and the need for sophisticated electronics.

A portable system for monitoring cardiovascular and respiratory function in large animals is available. Pulmonary function testing of cattle is also being examined and may provide some understanding of the pathophysiology of respiratory tract disease. Calves between 1 and 8 months of age with chronic respiratory disease have:⁸²

Arterial oxygen and carbon dioxide tensions are the only variables which correlate with clinical scores.

ARTERIAL BLOOD GAS ANALYSIS

Measurement of P_ao₂, P_aco₂ and arterial oxygen content (C_ao₂) provides valuable information about pulmonary function and oxygen delivery to tissues. The arterial oxygen tension and arterial oxygen content are not equivalent. The arterial oxygen tension (P_ao₂) is a measure of the partial pressure of oxygen in arterial blood determined by the amount of oxygen dissolved in the blood (not the amount bound to hemoglobin) and the temperature of the blood – it is not a direct measure of arterial oxygen content. Arterial oxygen content is the amount of oxygen per unit of blood and includes both dissolved oxygen and that bound to hemoglobin. The oxygen tension can be viewed as the driving force for diffusion of oxygen from capillaries into mitochondria (in which the oxygen tension is about 2 mmHg), whereas the oxygen content is the amount of oxygen delivered to tissue. Both are important measures of pulmonary function and oxygen delivery to tissue.

Measurement of oxygen tension in blood is achieved by analysis of an appropriately collected sample of arterial blood using a blood gas analyzer (oxygen electrode). Instruments designed for medical or veterinary clinical use measure pH, Po₂ and Pco₂ at a temperature of 37°C. Depending on the software included with the instrument, various derived values are also reported, including bicarbonate concentration, base excess and oxygen saturation. It is important to understand that oxygen saturation reported by blood gas instruments is a calculated value and might not be correct. Oxygen saturation is measured by a co-oximeter, which is different from a blood gas machine, and the amount of oxygen carried by hemoglobin is then calculated from this value and the assumption that each gram of hemoglobin, when fully saturated, carries approximately 1.34–1.39 mL of oxygen. The total oxygen content of blood is calculated by adding the amount carried by hemoglobin to the amount of oxygen dissolved in the aqueous phase of the blood. The formula is:

where O₂content is in mL/100 mL, S_ao₂ is the arterial oxygen saturation (%), 1.34 is the amount of oxygen carried by fully saturated hemoglobin (mL/g), [Hb] is the concentration of hemoglobin in blood (g/100 mL), 0.003 is the amount of oxygen dissolved in the aqueous phase of 100 mL of blood for each 1 mmHg increase in Po₂, and P_ao₂ is the oxygen tension in arterial blood. The appropriate substitutions can be made to calculate the oxygen content of venous blood.

The oxygen content of arterial blood is the critical factor (with cardiac output) in determining oxygen delivery to tissues. However, measurement of arterial oxygen content is not as readily accomplished as measurement of arterial oxygen tension. Therefore, in animals with normal hemoglobin concentration and function the arterial oxygen tension is used as a surrogate measure of arterial oxygen content. In doing so, it must be recognized that the extent of hemoglobin saturation with oxygen is dependent on both the affinity of hemoglobin for oxygen and the oxygen tension of the blood. The oxygen tension/percentage saturation relationship is sigmoidal, with 50% saturation occurring at about 30 mmHg in most species (there are minor variations) and 80% saturation at a Po₂ of 45–55 mmHg.⁹⁴ The sigmoidal shape of the oxygen– hemoglobin saturation curve has important clinical consequences. Small decrements in P_ao₂ from normal values (usually 95–105 mmHg in animals breathing ambient air at sea level) have a minimal effect on oxygen content of blood. Many modern blood gas analyzers have software that calculates oxygen content of blood, but it must be recognized that these calculations often use an assumed, not measured, hemoglobin concentration (usually 15 g/dL) and values for the human So₂–Po₂ relationship. These assumed values may not be correct for animals and one should always check the assumptions used to calculate oxygen content of blood before accepting and acting on those values. Direct measurement of blood oxygen content is restricted to research laboratories – indirect estimates gained from oxygen saturation and hemoglobin concentration are usually sufficiently accurate for clinical use.

The oxygen tension in blood is proportional to the amount of oxygen dissolved in the aqueous phase of the blood and the temperature of the blood. For a given amount of oxygen dissolved in blood, the tension varies according to the temperature of the animal. Almost all blood gas analyzers measure the Po₂ at 37°C. If the animal’s body temperature is markedly different from that then the reported Po₂ can be erroneous. For instance, the P_ao₂ of a horse with a body temperature of 40°C measured using an analyzer with a temperature of 37°C would be 80 mmHg (the Pco₂ would be 35 mmHg). If the P_ao₂ was adjusted for the difference between the horse’s body temperature and that of the analyzer, then the reported P_ao₂ would be 100 mmHg (and the P_aco₂ would be 44 mmHg). Failure to make the appropriate temperature corrections can result in errors of 6–7% per °C.⁹⁵ When interpreting blood gas values, attention should be paid to the temperature of the animal and consideration given to adjusting gas tension values according to the animal’s body temperature. This is probably only clinically important when there are extreme deviations from normal temperature and oxygen tension. Most blood gas analyzers include software that makes the appropriate corrections.

The arterial oxygen tension is determined in the alveolus by the alveolar oxygen tension and the alveolar–arterial difference. The alveolar oxygen tension (P_Ao₂) can be calculated from:

where F_io₂ is the inspired oxygen fraction (21% for ambient air), P_B is the barometric pressure (760 mmHg at sea level), PH₂o is the partial pressure of water vapor in the alveolar air (4 mmHg at 37°C), and RQ is the respiratory quotient (usually assumed to be 0.8 for resting animals). The alveolar–arterial Po₂ difference (A–a Po₂) is calculated:

The A–a Po₂ difference has clinical significance in that it is an indicator of pulmonary function that is somewhat independent of inspired oxygen fraction and is therefore useful in animals being supplemented with oxygen (there is a small increase in A–a difference with marked increases in F_io₂). Increases in A–a Po₂ difference are indicative of ventilation/perfusion mismatches, with the A–a Po₂ difference increasing with worsening ventilation/perfusion abnormalities.

Normal values

Values obtained from clinically normal animals breathing room air at sea level vary slightly between species, with most animals having an arterial P_ao₂ of 95–105 mmHg and a P_aco₂ of 35–45 mmHg. Oxygen saturation in clinically normal animals breathing air at sea level is above 98% and oxygen content of arterial blood is 16–24 mL/dL of blood (this depends on the hemoglobin concentration in blood). The difference in oxygen content of arterial and mixed venous blood is usually 4–8 mL/dL of blood. Values can be influenced substantially by changes in physiological state (exercise, hyperpnea), positioning, pulmonary disease and altitude (Table 10.4). Positioning of the animal can be important, especially in neonatal foals, in which the compliant chest wall can impair ventilation in laterally recumbent foals – foals have lower arterial oxygen tension when in lateral recumbency than when in sternal recumbency.⁹⁶

Table 10.4 Changes in blood gas tensions in various disease states compared to values in normal animals breathing air at sea level

Collection of arterial blood gas samples

Arterial samples can be collected from any of the appropriate peripheral arteries, which vary depending on species. An arterial sample is representative of aortic blood in almost all instances. Samples can be collected from the carotid, transverse facial, metacarpal and metatarsal arteries in horses and foals, and from the carotid, radial and coccygeal arteries in cattle and calves. Minimally invasive arterial access is difficult in pigs.

Samples should be collected in glass or plastic syringes in which the dead space has been filled with heparin solution. Typically, a 3 mL plastic syringe containing approximately 0.1 mL of sodium heparin and attached to a 22–25-gauge needle is used. All air should be expelled from the syringe before collection of the sample, and care should be taken to not introduce air into the syringe until blood gas tensions are measured. Air in the syringe will increase the measured oxygen tension of blood from normal animals. The sample should be measured as soon after collection as possible (within minutes). If immediate analysis is not available, the sample should be stored in iced water until analysis to prevent consumption of oxygen, production of carbon dioxide and a decrease in pH. Storage of arterial samples in plastic syringes in iced water can increase the oxygen tension from 100 mmHg to 109 mmHg in as little as 30 minutes.⁹⁷ This does not occur when samples are stored in glass syringes in iced water. The pH_a and P_aco₂ are not affected by the type of syringe.

VENOUS BLOOD GAS ANALYSIS

Measurement of gas tensions in venous blood is of limited value in assessing pulmonary function because of the extensive and variable effects of passage through the capillary beds on gas tensions. However, measurement of venous oxygen tension, saturation or content can be useful in assessment of the adequacy of oxygen delivery to tissue. The oxygen tension, saturation and content of venous blood depends on the extent of oxygenation of arterial blood, the blood flow to the tissues, the metabolic rate of the tissues drained by the veins from which blood is sampled, and the transit time of blood through capillaries. The multiplicity of these factors means that determining the precise reasons for abnormalities in venous blood gas tensions is not possible. However, some generalizations can be made about venous oxygen tension, saturation and content.

In normal, resting animals, oxygen delivery to tissues exceeds oxygen needs (demand) of the tissue, with the result that venous blood draining these tissues is only partially desaturated. Hence, venous blood from the pulmonary artery (mixed venous blood) has oxygen tension, saturation and content of approximately 35–45 mmHg, 80–90% and 12–18 mL/100 mL (the latter depending on hemoglobin concentration in addition to hemoglobin saturation). However, in situations in which oxygen delivery to tissue is decreased to levels that only just meet or do not meet the oxygen needs of tissue, there is extraction of a greater proportion of the oxygen in blood and venous oxygen tension, saturation and content decline and the arterial–venous difference in oxygen content increases. Reasons for oxygen delivery to tissue not meeting the oxygen needs of that tissue are decreased perfusion of tissue, such as can occur with shock or circulatory failure, anemia or decreased P_ao₂. Additionally, tissues with a high metabolic rate, such as exercising muscle, have high oxygen demands that can outstrip delivery.

Ideally, whole body assessment of oxygen delivery by measurement of venous blood gas tensions is best achieved by examination of mixed venous blood. Mixed venous blood represents an admixture of blood draining all tissues and is collected from the pulmonary artery (although samples collected from the right ventricle or atrium are also appropriate in most instances). While this blood is optimal for assessment of oxygen delivery to tissue, collection of mixed venous samples is not routine because of the need for catheterization of the pulmonary artery. Samples from peripheral veins are therefore used, but care should be taken when interpreting these values as venous blood gas tensions can vary considerably among veins.⁹⁸ For animals with normal circulatory status, blood gas tensions in jugular vein blood are likely to be reasonable estimates of mixed venous gas tensions. However, if circulatory function is not normal, then samples from peripheral veins may not be indicative of values in mixed venous blood.

Samples for venous blood gas analysis should be collected into syringes in which the dead space is filled with sodium or lithium heparin solution. The volume of heparin should not be more than 2% of the amount of blood. Samples should be processed promptly. If samples cannot be processed within an hour they should be stored in iced water. Samples stored in iced water for 24 hours have values that are minimally different from those before storage, while samples stored at 25°C change markedly in 2–3 hours.^99,100

PULSE OXIMETRY

Pulse oximeters are devices for measurement of blood oxygen saturation that attach to skin or mucous membranes and sense the absorption spectrum of light by hemoglobin (the same principle is used in bench top co-oximeters) in the underlying tissues. The devices are widely used for noninvasive monitoring of oxygenation in humans and have been adopted for use in animals. However, important challenges to their use exist in animals, not least of which is the presence of hair and densely pigmented skin in most farm animals. The devices have important deficiencies when used in foals and adult horses but those applied to the ear, lip or tongue of foals have good sensitivity and specificity for detecting arterial So₂ of less than 90 mmHg (12 kPa).^101,102 The devices consistently underestimate arterial So₂ at low saturations.^101,102 Care should be taken when using these devices to monitor arterial hemoglobin saturation in animals.

BLOOD LACTATE CONCENTRATION

Measurement of blood lactate concentration is useful in assessing the adequacy of oxygen delivery to tissues. Hypoxia causes a shift to anaerobic metabolism and the production of lactate. Lactate production is related to the severity and duration of hypoxia, with more severe hypoxia resulting in greater accumulation of lactate in tissues and its subsequent diffusion or transport into blood. Hypoxia also reduces the rate of removal of lactate from blood. The combination of increased production and decreased removal causes lactate to accumulate in blood. Measurement of blood lactate concentrations (which are usually lower than plasma lactate concentrations) is gaining increasing clinical usefulness as ‘point-of-care’ analyzers become more readily available and testing more affordable.

Samples for measurement of blood lactate can be collected into syringes containing heparin solution (as used for measurement of blood gas tensions) if the sample is to be analyzed within 30 minutes.¹⁰³ Samples should be stored in iced water until analysis. Prolonged storage at room temperature results in increases in blood lactate concentration. If sample collection is anticipated to be delayed, then samples should be collected into evacuated tubes containing sodium fluoride and potassium ethylenediamine tetraacetic acid (EDTA) – the sodium fluoride inhibits glycolysis. However, plasma lactate concentrations collected in these tubes are approximately 10% lower than in samples collected into tubes containing heparin – probably because of the osmotic effect of sodium fluoride/potassium EDTA on red cells. Samples for clinical analysis should be collected into syringes containing a heparin solution and analyzed within 30 minutes of collection. Measurement of blood or plasma lactate concentrations can be made using ‘point-of-care’ analyzers, although these can yield results that differ markedly from traditional analyzers, especially in animals with extreme values for hematocrit (severe anemia or polycythemia). Ideally, blood and plasma lactate concentrations should be measured only on analyzers that have been validated for the species and clinical situation being studied.¹⁰⁴

Blood lactate and plasma lactate concentrations are not equal, with blood lactate concentration being lower because of the dilutional effect of red blood cells, which have a lower lactate concentration than plasma. However, most clinical assessments are based on blood lactate concentrations. Mixed venous or arterial blood lactate concentrations in most farm animal species are less than 2 mmol/L in normal, healthy animals. Tissue hypoxia, in addition to other conditions such as toxemia and septic shock, can increase blood lactate concentration. Blood lactate concentrations between 2 and 4 mmol/L should be interpreted with caution whereas values above 4 mmol/L are indicative of clinically important disruption of oxygen transport and cellular metabolism. Repeated measurements over time can be useful for assessing progression of disease or efficacy of treatment. For instance, plasma lactate concentrations above 4 mmol/L in cattle with pneumonia are predictive of death within 24 hours.¹⁰⁵

COLLECTION AND ANALYSIS OF EXHALED BREATH CONDENSATE

Collection and analysis of exhaled breath condensate has use primarily in research studies at the current time. Breath condensate is collected and analyzed for markers of pulmonary or systemic disease. Induction of pneumonia in calves by infection with Pasteurella multocida causes increases in concentration of leukotriene B₄ in breath condensate.¹⁰⁶ Horses with heaves have higher concentrations of hydrogen peroxide than normal horses – probably a result of the airway neutrophilia in affected animals.¹⁰⁷

LUNG BIOPSY

Percutaneous biopsy of the lung is useful in confirming diagnosis of lung disease by providing tissue for histological and microbiological examination. The procedure in cattle, sheep and horses is described.^108-110 Indications for the procedure include the presence of diseases of the lungs in which a diagnosis cannot be arrived at through other forms of examination, including tracheal aspiration or bronchoalveolar lavage. It can also be used for assessing the severity of histological changes and response to therapy. The procedure is best suited for widespread diseases of the lung, but can be used for diseases that produce focal lesions if the biopsy is performed with ultrasonographic guidance. Contraindications include abnormalities in clotting function, pneumothorax and severe respiratory distress. The danger in performing lung biopsy in animals in severe respiratory distress is that complications of biopsy, such as pneumothorax, hemothorax or hemorrhage into airways, could further impair lung function and cause the death of the animal.

Complications include pneumothorax, hemothorax, hemorrhage into airways with subsequent hemoptysis or epistaxis, pulmonary hematoma and dissemination of infection from infected lung to the pleural space. Pneumothorax, which is usually not clinically apparent, occurs in most horses in which the procedure is performed.¹⁰⁸ Coughing and epistaxis occur in about 20% and 10% of horses, respectively.¹⁰⁸ Life-threatening hemorrhage occurs uncommonly (= 2% of cases). Bleeding into the airways, detected by tracheobronchoscopic examination, occurred in 16 of 50 horses after use of the manually discharged biopsy needle and in five of 50 horses after use of the automatically discharged needle.¹⁰⁸ Two of 60 cows collapsed immediately after the procedure, but subsequently stood and recovered.¹⁰⁹ The remaining cows had no clinical abnormalities detected after biopsy, although necropsy examination 24 hours later revealed small lesions in the pulmonary parenchyma at the site of biopsy.¹⁰⁹ One of 10 healthy sheep had coughing and bloody nasal discharge after lung biopsy.¹¹⁰

The procedure is performed in adult horses and cattle using a 14-gauge biopsy needle, either manually operated or one that discharges automatically. Such instruments yield tissue in over 95% of attempts in cattle.¹⁰⁹ The area for examination is best determined by radiographic or ultrasonographic examination of the thorax. A common site for biopsy is at the junction of the dorsal and middle thirds of the thorax at the ninth intercostal space in cattle and sheep^109,110 and the 13th intercostal space in horses. The procedure is best performed with the animal standing. The skin over the area should be clipped of hair and aseptically prepared and local anesthesia induced by injection of 2% lidocaine or a similar compound into the intercostal space. A 0.5 cm incision is made through the skin and the biopsy instrument is advanced through the caudal intercostal space (intercostal vessels and nerves course along the caudal aspect of the ribs) and into the lung perpendicular to the skin surface. The instrument is advanced approximately 2 cm into the lung and tissue is collected at the end of inspiration. The procedure is repeated as necessary for collection of samples for histological and microbiological examination. The skin incision is closed with a single suture if necessary. The animal is then monitored closely for 12–24 hours for signs of coughing, epistaxis, hemoptysis, fever or respiratory distress. Hemorrhage into the airways is usually evident, often within minutes of completing the procedure, by the animal coughing. Hemorrhage into the airways is often evident as hemoptysis, even in horses. Respiratory distress can be caused by pneumothorax, hemothorax or hemorrhage into airways. Treatment includes percutaneous aspiration of pleural air, administration of oxygen by insufflation or, in extreme instances, mechanical ventilation.

RESPIRATORY SOUND SPECTRUM ANALYSIS

Analysis of respiratory sounds has utility in the diagnosis of disorders of the upper respiratory tract of horses. Respiratory sounds can be detected by a small microphone near the horse’s nostril with the recording made by a tape recorder or similar device worn on the saddle or girth strap.^111,112 Studies can be performed with horses running on either a treadmill or outside over ground. Dorsal displacement of the soft palate produces broad-frequency expiratory noises with rapid periodicity (rattling), whereas dynamic unilateral collapse of the arytenoid causes an increase in inspiratory broad band high-frequency noise.^111-113 The technique correctly identifies more than 90% of horses with dynamic collapse of the left arytenoid cartilage (‘roarers’).¹¹³

EXERCISE TESTING

Exercise testing for assessment of respiratory tract function is essentially limited to horses. Such tests are usually conducted on a treadmill, although some are amenable to use in the field. Tests available for use on horses running on a treadmill include endoscopic examination of the upper airway, respiratory noise analysis, blood gas analysis and measurement of respiratory mechanics. The most important of these in a clinical setting is videoendoscopy, during exercise on a treadmill, to detect dysfunction of the upper airway of horses.¹¹⁴ Some disorders of the upper respiratory tract, such as progressive weakness of the laryngeal abductor muscles, axial deviation of the aryepiglottic fold and epiglottic retroversion, can only be diagnosed by endoscopic examination performed during strenuous exercise.¹¹⁵ The interested reader is referred to texts devoted to this topic.¹¹⁶

TREATMENT OF RESPIRATORY DISEASE

Treatment of diseases of the lower respiratory tract depends on the cause of the disease. However, the common principles are:

CONTROL OF RESPIRATORY DISEASE

Infectious diseases of the respiratory tract of farm animals are caused by a combination of infectious agents and predisposing causes such as inclement weather, the stress of weaning or transportation and poorly ventilated housing, each of which can weaken the defense mechanisms of the animal. Prevention and control of these diseases include:

PULMONARY CONGESTION AND EDEMA

Pulmonary congestion is caused by an increase in the amount of blood in the lungs due to engorgement of the pulmonary vascular bed. It is sometimes followed by pulmonary edema when intravascular fluid escapes into the parenchyma and alveoli. The various stages of the vascular disturbance are characterized by respiratory compromise, the degree depending upon the amount of alveolar air space which is lost.

PULMONARY HYPERTENSION

Pulmonary hypertension is an increase in pulmonary arterial pressure above normal values due to structural or functional changes in the pulmonary vasculature. Primary pulmonary hypertension occurs in cattle with high-altitude disease. Chronic pulmonary hypertension results in right-side congestive heart failure due to right ventricular hypertrophy or cor pulmonale.

ATELECTASIS

Atelectasis is collapse of the alveoli due to failure of the alveoli to inflate or because of compression of the alveoli. Atelectasis is therefore classified as obstruction (resorption), compression or contraction. Obstruction atelectasis occurs secondary to obstruction of the airways, with subsequent resorption of alveolar gases and collapse of the alveoli. This disease is usually caused by obstruction of small bronchioles by fluid and exudate. It is common in animals with pneumonia or aspiration of a foreign body. Compression atelectasis occurs when intrathoracic (intrapleural) pressure exceeds alveolar pressure, thereby deflating alveoli. This occurs when there is excessive pleural fluid or the animal has a pneumothorax. In large animals it also occurs in the dependent lung or portions of lung in recumbent animals. Compression atelectasis is the explanation for the large shunt fraction and hypoxemia that occurs in anesthetized horses.¹ Compression atelectasis and secondary bronchopneumonia can occur in horses kept in flotation tanks for up to several weeks for treatment of skeletal injuries.² Contraction atelectasis occurs when there is compression of parts of the lung by fibrotic changes in the pleura. Patchy atelectasis occurs in the absence of surfactant, such as can occur in newborns. Failure of the lung to inflate, or development of atelectasis of the lungs of the newborn, usually those born prematurely, occurs because of lack of pulmonary surfactant. The disorder can progress to hyaline membrane disease. Affected newborn animals are severely dyspneic, hypoxemic, cyanotic, weak and commonly die in a few hours.

The clinical signs of atelectasis are not apparent until there is extensive involvement of the lungs. Animals develop respiratory distress, tachypnea, tachycardia and cyanosis. Blood gas analysis reveals hypoxemia, with or without hypercapnia. Thoracic radiographs reveal pulmonary consolidation. Ultrasonographic examination of the thorax demonstrates consolidated lung.

Atelectasis is reversible if the primary obstruction or compression is relieved quickly before secondary consolidation and fibrosis occur.

ACUTE RESPIRATORY DISTRESS SYNDROME

This is a well-recognized clinical syndrome of humans characterized by acute onset of hypoxemia and pulmonary infiltrates without increases in left atrial pressure (i.e. without evidence of cardiogenic pulmonary edema). Precipitating causes include both direct and indirect lung injury, including sepsis, multiple transfusions, trauma, near-drowning, smoke inhalation, pancreatitis and more. The underlying lesion is diffuse alveolar capillary damage with secondary severe pulmonary edema. The disease occurs spontaneously in domestic animals¹ and, although the spontaneous disease is not extensively documented, the disease produced experimentally as a model of the human disease is better described.²

Acute respiratory distress syndrome (ARDS) in animals occurs in newborns and in adult animals. The disease in some newborn farm animals is related to lack of surfactant but except for animals born prematurely this is more the exception than the rule. Most young animals and all adult animals with ARDS have some inciting acute lung injury that then progresses to ARDS.^1,3,4 The causes can be infectious (e.g. influenza virus infection), physical (smoke inhalation) or toxic (endotoxin).

The pathophysiology of the disease involves a common final pathway that results in damage to alveolar capillaries. The initial injury can be to either the endothelium of pulmonary capillaries or to alveolar epithelium. Damage to these structures leads to extravasation of protein-rich fluid and fibrin with subsequent deposition of hyaline membranes. The capillary injury is attributed to activated leukocytes (macrophages and neutrophils) and cytokines. Accumulation of hyaline membranes and ventilation/perfusion mismatches impair respiratory gas exchange and cause hypoxemia.

The clinical signs are characteristic of acute, progressive pneumonia. Animals are anxious, tachycardic, tachypneic and have crackles and wheezes on thoracic auscultation. Severely affected animals can be cyanotic. Thoracic radiographs reveal diffuse pulmonary infiltrates. Hematologic changes are characteristic of the inciting disease but usually include leukopenia. There is arterial hypoxemia.

Treatment includes administration of anti-inflammatory drugs (NSAIDs with or without glucocorticoids), colloids, antimicrobials and oxygen. The arterial blood gas response to oxygen therapy is often minimal in severely affected animals. If it is available, mechanical ventilation can be useful, although the prognosis is grave. Inhalation of nitric oxide is beneficial in some humans with the disease, and there are anecdotal reports that it has been used to treat foals with ARDS.