General
Laboratory Testing
Basic blood work (complete blood count and biochemical panel) are often performed during the workup of a respiratory patient and may help support the presence of an underlying respiratory tract disease. In local diseases associated with inflammation or infection such as rhinitis and tracheobronchitis, hematologic changes are rarely found. With parenchymal diseases such as bacterial or aspiration pneumonia, a neutrophilic leukocytosis is often found. Neutrophilia or eosinophilia is reported in eosinophilic pneumonia/bronchopneumopathy and feline bronchial disease. Fungal pneumonia is anticipated to result in neutrophilia and monocytosis, reflecting the chronic nature of disease. Chronic hypoxemia can result in polycythemia.
Biochemical abnormalities in respiratory diseases are usually nonspecific. Hyperglobulinemia can be found in feline bronchial disease, fungal pneumonia, chronic foreign body or aspiration pneumonia, or bronchiectasis due to chronic antigenic stimulation, and concurrent hypoalbuminemia is occasionally present as a negative acute phase reactant.
Molecular diagnostics are increasingly used to document the presence of an infectious organism, such as feline herpesvirus-1, Bordetella, or Mycoplasma, in either upper or lower respiratory tract disease; however, there are important limitations to interpretation of results (see sections on specific diseases). Also, it is critical to realize that a positive molecular assay does not confirm that the organism is responsible for the clinical disease identified.
Pulse Oximetry
Pulse oximetry provides an estimate of hemoglobin saturation with oxygen and is inexpensive, noninvasive, and easy to perform. The technique relies on detection of the optical density of the pulse wave as blood passes through the arterial system. The sensor subtracts the signal between pulses from the height of the pulse wave to determine oxygenation of inflowing blood only. Because of this feature, pulse oximetry can provide a falsely low measurement in a hypotensive patient with weak pulses or in an animal with anemia. This technique cannot differentiate between methemoglobin and oxyhemoglobin.
Pulse oximetry is useful prior to anesthetizing the patient for a respiratory procedure or as a monitoring tool during therapy. Sites that can used to obtain a measurement include the lip, tongue, between the toes, on the ear pinna, and sometimes on the abdomen. The probe can be applied to various sites several times to obtain a signal, and detection of a strong pulse rate indicates that the reading is likely accurate. A pulse oximeter reading below 95% correlates with a PaO2 of less than 80 mm Hg (Figure 2.1). When such a reading is obtained, an arterial blood gas analysis should be performed, if available, to confirm the degree of hypoxemia. It is important to remember that the pulse oximeter measures only oxygenation. It provides no information on ventilatory status and thus cannot detect hypoventilation (increased PaCO2) in an animal.
Figure 2.1. Pulse oximetry measures hemoglobin saturation with oxygen, which has a sigmoidal relationship with the partial pressure of arterial oxygen. A hemoglobin saturation <95% equates to a PaO2 <80 mm Hg and indicates hypoxemia.
Pulse oximetry can be valuable in determining response to therapy in hypoxemic patients because improvements in oxygenation occur prior to radiographic changes. However, because of the sigmoidal relationship between hemoglobin saturation and arterial oxygen, oximetry remains a somewhat crude estimate of lung function.
Arterial Blood Gas Analysis
Arterial samples are obtained by direct puncture of the dorsal metatarsal artery in dogs and the femoral artery in cats and small dogs. A small needle (23–25 gauge) on a heparinized syringe is used, or a self-filling syringe can be used. The artery is palpated and stabilized with two fingers of one hand while the syringe is firmly placed through the wall of the artery. Approximately 0.5-mL blood is needed for analysis and the sample must be stoppered and stored on ice until analyzed. After withdrawal of the syringe, firm pressure is applied to the artery for 3–5 minutes. An arterial blood gas analysis measures PaO2, PaCO2, pH, total CO2, and hemoglobin saturation with oxygen, and allows calculation of bicarbonate, base excess and deficit, and oxygen content (Table 2.1).
Alveolar–arterial oxygen gradient and PF ratio
The alveolar–arterial (A–a) oxygen gradient estimates the difference between the calculated alveolar oxygen level expected for the animal and the measured arterial oxygen level. Thus, the A–a gradient corrects for the level of ventilation performed by the animal and allows comparison of blood gas data through the course of disease that is not impacted by the effect of an increase or a decrease in PaCO2 on PaO2. The A–a oxygen gradient is calculated as
where FiO2 is the fraction of inspired oxygen (0.21 on room air), PB is the barometric pressure (inmmHg), PH2O is the water vapor pressure (47 mm Hg at 37°C), and R is the respiratory quotient (ratio of CO2 production to O2 consumption, usually assigned a value between 0.8 and 1.0). PaO2 and PaCO2 are obtained from blood gas analysis. Normal value is <15.
The PaO2/FiO2 ratio (PF or oxygenation ratio) provides a measure of the ability of the lung to oxygenate as the fraction of inspired oxygen changes from room air to 100% oxygen. This is calculated by dividing arterial oxygen by FiO2 (ranging from 0.21 to 1.0). Normal animals have a PF ratio of >500. Values between 300 and 500 indicate mild impairment of oxygenation, while values <200 indicate serious decrements in oxygenation. A PF ratio <200 is one of the requirements for a diagnosis of acute respiratory distress syndrome.
Table 2.1. Normal blood gas values for dogs and cats
| Dog | Cat | |
| PaO2 (mm Hg) | 90 (80–105) | 100 (95–105) |
| PaCO2 (mm Hg) | 37 (32–43) | 31 (26–36) |
| pH | 7.35–7.45 | 7.35–7.45 |
| HCO3 (mmol/L) | 22 (18–26) | 18 (14–22) |
Table 2.2. Respiratory causes of hypoxemia
| Mechanism | Clinical Attributes | Causes |
| Hypoventilation | High PaCO2 Normal A–a gradient Improved by oxygen supplementation Improved by increasing alveolar ventilation | Anesthesia Upper airway obstruction Neuromuscular weakness CNS disease |
| V/Q mismatch | Increased A–a gradient Mildly increased PaCO2 Markedly improved by oxygen supplementation | Virtually any lung disease |
| Shunt | Increased A–a gradient Not improved by oxygen supplementation Not improved by increasing alveolar ventilation | Congenital right to left cardiac shunts Acute respiratory distress syndrome |
| Diffusion impairment | Increased A–a gradient Seldom a major cause of hypoxemia at rest Causes hypoxemia during exercise or with low inspired oxygen Improved by oxygen supplementation | Interstitial lung disease Pulmonary edema |
| Reduced inspired oxygen | Improved by oxygen Causes hypoxemia during exercise or when diffusion is impaired | High altitude |
Causes of hypoxemia
Obtaining an arterial blood gas, calculating the A–a gradient, and assessing response of hemoglobin saturation or PaO2 to exogenous oxygen supplementation allow assumptions to be made about the most likely mechanism causing hypoxemia (Table 2.2). This can help determine the most likely underlying cause of hypoxemia, although ventilation/perfusion mismatch contributes to the pathophysiology of hypoxemia in almost all lung diseases.
Imaging
Radiography is often the key to creating an appropriate list of differential diagnoses for the respiratory case and for determining the type of sampling method that is most likely to achieve a final diagnosis, such as endoscopy, fine-needle aspiration (FNA), or a tracheal wash (Table 2.3). It will also help determine the need for advanced imaging including fluoroscopy, ultrasound, nuclear scintigraphy, or computed tomography. Specific features of these tests are presented in the relevant disease sections.
Table 2.3. Airway-sampling techniques for various lung patterns
| Radiographic Pattern | Differential Diagnoses | Sampling Technique |
| Interstitial | Viral pneumonia Rickettsial pneumonia Protozoal pneumonia Hemorrhage Vasculitis Pulmonary fibrosis Neoplasia Early pulmonary edema Aspiration pneumonia | Fine-needle aspirate Lung biopsy Bronchoscopy Tracheal wash |
| Bronchial | Chronic bronchitis Feline bronchitis/asthma Bronchiectasis Parasitic bronchitis Early bronchopneumonia | Tracheal wash Bronchoscopy |
| Alveolar | Bronchopneumonia Aspiration pneumonia Fungal pneumonia Hemorrhage Pulmonary edema Neoplasia Noncardiogenic pulmonary edema | Tracheal wash Bronchoscopy |
| Consolidation | Neoplasia Lung lobe torsion Consolidating pneumonia Granuloma Bronchial obstruction Feline bronchitis Foreign body inhalation | Bronchoscopy Fine-needle aspirate Tracheal wash |
| Vascular | Congenital heart disease Congestive heart failure Heartworm disease Pulmonary hypertension Pulmonary thromboembolism | Echocardiography |
| Effusion | Hydrothorax Pyothorax Hemothorax Chylothorax Neoplasia Diaphragmatic hernia | Thoracocentesis |
Airway Sampling
Transoral Tracheal Wash
Transoral tracheal wash is appropriate for use in large and small dogs or in cats. A sterile endotracheal tube and a sterile polypropylene or blunt-ended red rubber catheter (3.5–8 French) are needed. Prior to doing a tracheal wash, the catheter should be measured against the animal and marked at a position that estimates the location of the carina, which is approximately at the fourth intercostal space. Passing the catheter too far distally can result in airway damage. The animal is anesthetized with a short-acting anesthetic agent. Options include propofol, ketamine–valium, or a balanced protocol using a narcotic agent. Prior to intubation, the function of the larynx is assessed. In the cat, local lidocaine can be used to facilitate intubation and avoid contamination of the tube through contact with oropharyngeal or laryngeal mucosa. An assistant holds the tube in place during the lavage; however, if retrieval of fluid is less than desired, the cuff of the endotracheal tube can be inflated to improve suction.
With the endotracheal tube held in place, the polypropylene or red rubber catheter is passed sterilely to the level of the carina, and the three-way stopcock with syringe is attached to the outer port. An aliquot of saline (4–6 mL) is instilled into the airway, and suction is used to retrieve the fluid and cells from the lower airway. Either hand suction or house suction can be applied as needed. Retrieval of fluid can be enhanced by having the assistant compress the chest or by stimulating a cough during suction. Instillation and aspiration of fluid can be repeated several times until an adequate sample has been retrieved (0.5–1.0 mL is usually sufficient for culture and cytology).
A modification of the transoral tracheal wash can be performed, which yields a sample more closely approximating that of a bronchoalveolar lavage. In the dog, this is achieved using a 16-French ArgyleTM stomach tube (Sherwood Medical Co/Tyco Healthcare Kendall, Deland, FL) (Hawkins and Berry 1999). The dog must be large enough to accommodate an endotracheal tube that will not be fully occluded by the stomach tube. The stomach tube is shortened to remove any side holes on the distal end of the tube and to create a length that approximates the distance from the endotracheal tube to the last rib. The distal end of the tube can be tapered with a pencil sharpener to improve the ability to wedge within the airway, and the tube should be sterilized prior to use. Endotracheal intubation is carried out using a short-acting anesthetic agent (e.g., propofol) and a sterile endotracheal tube. The dog is placed in dorsal recumbency, and the modified stomach tube is passed through the lumen of the endotracheal tube until it meets resistance. Gentle pressure should be used when passing and wedging this tube to avoid perforating the lung. As soon as slight resistance is encountered, the tube is withdrawn 1–2 cm and lavage is initiated with 20 mL of sterile saline followed by 5 mL of air. Gentle hand suction is applied to retrieve the fluid, and a second aliquot can be instilled as needed.
Nonbronchoscopic bronchoalveolar lavage (BAL) has also been reported in the cat, and the cell distribution obtained on cytology matches that found with bronchoscopy (Hawkins et al. 1994). For this procedure, the cat is anesthetized with a short-acting anesthetic agent (ketamine–valium in a 1:1 mixture or propofol) and intubated with a sterile endotracheal tube. The cuff is inflated and the cat is placed in lateral recumbency with the most affected side down. Aliquots (1–3 as needed) of warmed sterile 0.9% saline (5 ml/kg) are instilled directly into the endotracheal tube using a 35-mL syringe with syringe adapter. Fluid is retrieved by hand aspiration. Elevating the hindquarters can facilitate collection, and approximately 65–70% fluid retrieval should be expected. Alternately, a dog urinary catheter (6–8 French) can be passed gently through a sterile endotracheal tube until resistance is met, in a manner similar to that employed when a modified stomach tube is used to perform blind BAL in a dog (Foster et al. 2004). Instillation of 5–10 mL of sterile saline provides an adequate lavage sample for cultures and cytology. With either procedure, respiratory rate and pulse oximetry should be monitored to detect untoward reactions and oxygen supplied as needed.
Figure 2.2. An over-the-needle catheter can be used as a cannula to perform a tracheal wash using a sterile urinary catheter.
Transtracheal Wash
Transtracheal wash is appropriate for larger dogs (>8 kg) or those that cannot be anesthetized for a transoral tracheal wash. Generally only local anesthesia is needed although a mild sedative may facilitate completion of the procedure. The easiest way to perform a transtracheal wash is with a through-the-needle jugular catheter. An alternate approach is to use a 16-gauge over-the-needle catheter as a cannula to penetrate the trachea and a long, sterile 3.5-French urinary catheter to pass down into the airways for sample collection (Figure 2.2). The trachea can be entered at the cricothyroid notch but is preferable to enter the trachea between the tracheal rings lower on the neck to avoid potential damage to laryngeal structures. This will also facilitate collection of a sample from more distal airways since the jugular catheter is relatively short in length. For either technique, the ventral portion of the neck is clipped and lightly scrubbed with antiseptic solution followed by alcohol wipes. A more complete surgical preparation is performed after local anesthesia is instilled.
Local anesthesia with lidocaine (0.25–0.5 mL) is used at two sites that will be penetrated by the needle: at the skin and between the tracheal rings. The needle of the jugular catheter will penetrate the skin low on the neck, and the skin will then be drawn upward prior to entering the airway lumen. This creates a subcutaneous seal that limits air leakage between the skin entry and the airway.
To begin catheter placement, the needle is placed into the trachea with the bevel of the needle facing downward so that the catheter will pass over the short edge of needle rather than the long edge as it is advanced through the lumen. This makes it less likely that the sharp edge of the needle will cut off the catheter during passage. Tent the skin and pass the needle or catheter through the site that has been infiltrated with lidocaine. The needle or catheter is directed perpendicular to the trachea initially (Figure 2.3). When preparing to enter between the tracheal rings, stabilize the trachea firmly against the neck to prevent it from moving away from needle. After the needle has passed through the skin and subcutaneous tissue, the skin is retracted upward to enter the airway one to two tracheal rings above the site where the skin was penetrated. A distinct pop is usually felt when the needle enters the tracheal lumen, and the dog often coughs. With the needle in the airway, the angle made between the catheter and trachea is decreased to 60° to facilitate passage of the catheter down the lumen of the airway (Figure 2.3). The needle is withdrawn at this stage to pass the sampling catheter through the short catheter to the level of the carina (Figure 2.4).
Figure 2.3. To initiate a tracheal wash, the needle is directed into the trachea at a perpendicular angle (a). After the needle has penetrated between the tracheal rings, the hub of the needle (or the catheter being used as the cannula) is moved toward the neck to create a more parallel angle for passage of the long catheter down the trachea (b).
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