Advanced pulmonary function testing in camelids (Proceedings)

Evaluation of respiratory distress and dysfunction in camelids – sorting out dyspnea

Respiratory distress is generally defined as outwardly evident, labored respiratory efforts or ventilation i.e. the clinically apparent inability to adequately ventilate and/or oxygenate. Primary inspiratory dyspnea may be related to alveolar hypoventilation (impaired air delivery to the lung) or inadequate gas exchange. Expiratory dyspnea may accompany inspiratory distress in any animal with severe impairment of gas exchange. However, primary expiratory dyspnea is usually associated with obstruction of the intrathoracic airways by mucous and bronchospasm (e.g. recurrent airway obstruction).

The evaluation of lung and airway disease in SAC may pose a particular challenge to veterinarians, as llamas and alpacas often merely display subtle clinical abnormalities even in the face of progressive or severe respiratory disease. Specific analysis of the breathing pattern in camelids has shown that asynchronous breathing is commonly observed during conditions of upper airway obstruction, phrenic nerve dysfunction, diaphragmatic or intercostal muscle paralysis and diaphragmatic fatigue due to chronic airway disease.

High altitude adaptations

Llamas, and other South American Camelids, are native to the Andean altiplana region and show several evolutionary adaptations to environmental hypoxia. For example, the small elliptical red blood cells of camelids have a greater affinity for oxygen, a high hemoglobin concentration and high oxygen transfer conductance (facilitated by the small RBC size).

The oxygen dissociation curves of the blood of lamoids are thus shifted to the left, even in camelids adapted to low altitudes. Furthermore, llamas were identified to carry a higher blood oxygen content than, for example, sheep.A high muscle myoglobin concentration, more efficient oxygen extraction at the tissue level5 and high lactate dehydrogenase activity further improve the camelid's resistance to hypoxemia.

Gas exchange in camelids

Arterial blood gas analysis is preferentially used to determine oxygenation and ventilation in camelids. Technically, an arterial blood sample can most easily be obtained from the saphenous artery of the medial thigh. Normal arterial blood gas result have been documented in 14 adult, healthy male and non-pregnant female alpacas (mean weight: 62 ± 21kg, mean age: 4.5 yrs) as well as 12 healthy neonatal alpaca crias (2-7days old, mean 7.8 kg body weight):

Normal arterial blood gas results in healthy neonatal (2-7 days) and adult alpacas Variable n Mean 95% confidence interval pH 14 7.47 7.46-7.48 Bicarbonate (mEq/L) 14 20.4 19.6-21.3 PaO2 (adults)  14 104.6 100.7-108.4 Neonatal  PaO2 12 92 87.7-96.4 PaCO2 (adults)  14 27.7 26.4-28.9 Neonatal PaCO2 12 30.5 27.2-33.8

The effect of storage type (plastic vs. glass), temperature (iced at 4oC vs. 22oC room temperature) and blood gas tension (normoxemia, hypoxemia, hyperoxemia) was recently evaluated in arterial blood gases (ABG) of healthy alpacas, to facilitate ABG analysis under field conditions. Statistically significant alterations in pH, PaCO2, lactate and bicarbonate were only associated with room temperature storage.

However, mean changes in arterial blood gas parameters still remained below 3.40 mmHg for PaCO2, 0.5 mmol/L for lactate, 1.22 mEq/L for bicarbonate and 0.03 points for pH after two hour storage. In contrast, the PaO2 significantly decreased in both normoxic and hyperoxic blood samples, following storage in glass syringes at room temperature. Similar decreases in PaO2 were observed in hyperoxygenated blood stored at room temperature in plastic syringes. In contrast, PaO2 significantly increased after storage of hyperoxic blood samples in plastic syringes on ice.

Overall, storage in glass syringes on ice resulted in the lowest variability of PaO2, with the highest mean PaO2 increase being 10.28 mmHg under these conditions. Based on these results, ABG from alpacas should be stored on ice in glass syringes, to reduce variability in PaO2 if analysis is delayed. In contrast to human reports, the PaO2 of cooled hyperoxygenated camelid blood increases following storage in plastic syringes.

Specific lung function testing in camelids

A) Forced oscillatory techniques (FOT)

Novel non-invasive modalities such as Forced Oscillatory Techniques (FOT) may assess the obstruction to airflow in non-sedated camelids. In short, the flow and pressure change of small air oscillations that are superimposed on the animal's spontaneous breathing via a face mask, are measured to calculate respiratory system resistance. This technique is easily tolerated in untrained camelids and may rapidly distinguish upper versus lower airway disease, without compromising the animal's breathing.

B) The “open pleth”

The breathing pattern and lung function of llamas and alpacas may also be assessed using a plethysmographic technique called Open Pleth, as previously described. Simplistically, this system evaluates the synchrony of chest and abdominal breathing, while determining the patient's effort needed to overcome respiratory resistance. During this test, two elastic bands that contain a sinusoidal conducting wire are temporarily placed around the animal's chest and abdomen. Stretch and contraction of these bands due to normal breathing movements are measured as voltage alterations and quantify the changes in circumference and volume of the chest and abdomen during breathing.

Simultaneously, we measure the animal's airflow at the level of the nares, thus rapidly assessing asynchrony of the patient's breathing pattern due to airway obstruction and breathing muscle (diaphragmatic) dysfunction. This technique has been used to describe the breathing patterns of healthy alpacas, llamas, and a llama with diaphragmatic paralysis. Evidence of lower airway obstruction may be identified by measured differences between plethysmographic and nasal flow over the first 25% of expiration (delta flow). The average delta flow recorded in healthy alpacas is near zero (0.18+/- 0.07 L/s) as found in llamas (0.04 +/- 0.14 L/s).

C) Functional residual capacity (FRC)

FRC is defined as the volume of gas that remains in the respiratory system at passive end-expiration, and is determined by the balance of chest wall compliance and lung elastance. FRC may thus serve as an index of mechanical lung failure and assist in the diagnosis of lung fibrosis, airway inflammation, and inspiratory airway obstruction (e.g. laryngeal paralysis). The normal FRC in adult llamas was non-invasively measured via helium dilution, with a mean FRC of  39.18 ml/kg (± 11.45).

In this study, FRC was positively correlated (p = .05) to body length (point of shoulder to ischial tuberosity) but not to weight, height, or circumference in adult llamas. Similarly, the mean FRC in alpacas was reported as 3.19 + 0.53 L, which translated to 46.29 + 7.50 ml/kg. A positive correlation was observed between FRC and body weight in the study participants (P = 0.003; r = 0.645). A mean FRC of 45.13 ml/kg (95% CI: 40.9 – 49.4) was similarly obtained in eleven clinically healthy alpaca crias (age: 1–18 days).

Non-invasive pulmonary function testing (PFT) has been validated in 10 non-sedated llamas to established normal parameters of respiratory mechanical function. Measurements of pulmonary function included: functional residual capacity (FRC) via helium dilution, respiratory inductive plethysmography (RIP) to assess breathing pattern and flow limitations, esophageal-balloon pneumotachography and monofrequency forced oscillatory techniques (FOT: 1-7Hz) before and after sedation with 0.2 mg/kg xylazine IM.

The following measurements of respiratory function were obtained in non-sedated llamas: FRC (5.60 +/-1.24 L), tidal volume (VT = 1.03 +/- 0.3 L), dynamic compliance (Cdyn= 0.83 +/-0.4 L/cmH2O), pulmonary resistance (RL=1.42 +/-0.54 cmH2O/L/s) and respiratory system resistance (RRS in cmH2O/L/s) at 1 Hz (2.4 +/-0.9), 2 Hz  (2.3 +/-0.7) , 3 Hz (2.2 +/-0.6), 5 Hz (2.7 +/-0.7) and 7 Hz (2.5 +/-0.5) using FOT. Measurements of flow limitations via RIP were comparable to other species.

However, xylazine sedation induced significant increases in RL and maximal pleural pressure changes. A mean 127% increase in RL and mean 116% increase in RRS across 1-7 Hz, were observed following sedation. The magnitude of change in RRS increased with decreasing impulse frequency, suggesting bronchoconstriction. Similar lung function tests were recently performed in non-sedated alpacas, identifying the following results: tidal volume (VT =0.8 +/-0.13 L), respiratory system resistance (RRS in cmH2O/L/s) at 1Hz (2.70 +/-0.88), 2Hz  (2.98 +/-0.70) , 3Hz (3.14 +/-0.77), 5Hz (3.45 +/-0.91) and 7 Hz (3.84 +/-0.93).

The phase angle, as a measurement of thoracoabdominal asynchrony, was 19.59 +/-10.06° and the delta flow between nasal and plethysmographic flow measurements was 0.18 +/-0.07 L/s. The VT, peak inspiratory and expiratory flows were significantly higher in cushed compared to standing animals. The authors of both studies concluded that PFT was well tolerated in untrained, awake camelids and may have extensive application in clinical practice. Due to alteration in respiratory function, testing should not be performed after xylazine sedation.

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