Assisted Ventilation of Canada Geese (Branta canadensis) Anesthetized for Radio Transmitter Implantation
American Association of Zoo Veterinarians Conference 2002
R. Scott Larsen1,2, DVM, MS; Daniel M. Mulcahy1,3, PhD, DVM, DACZM; Jerry Hupp3, PhD
1Environmental Medicine Consortium, Department of Clinical Sciences, College of Veterinary Medicine, North Carolina State University, Raleigh, NC, USA; 2North Carolina Zoological Park, Asheboro, NC, USA; 3Alaska Biological Sciences Center, U.S. Geological Survey, Anchorage, AK, USA


Surgical implantation of radio transmitters allows the use of these devices in birds that do not tolerate externally applied equipment.3,5 Surgical implantation of transmitters usually involves invasion of the coelomic cavity and ablation of the right abdominal air sac. Air sac ablation may alter the respiratory dynamics of the anesthetized bird and affect anesthetic management, but the cardiopulmonary effects of this procedure have not been studied. We documented cardiorespiratory changes in Canada geese during surgical implantation of transmitters and assessed the effects of two rates of assisted ventilation.

Adult female Canada geese (Branta canadensis) were captured in Anchorage, Alaska for surgical implantation of intracoelomic transmitters. Eighteen geese were included in the anesthetic investigation. Each goose was manually restrained, and an intravenous (IV) catheter was placed in the left jugular vein. The bird was induced by face mask with 4% isoflurane (IsoFlo, Abbott Laboratories, North Chicago, IL) in oxygen, intubated, and then maintained on 1.5–4% isoflurane. Throughout surgery, half of the birds were ventilated at a low rate (LV; 2 breaths/min) and half at a high rate (HV; 6 breaths/min). Venous blood samples were taken for blood gas analysis during manual restraint (T0), at the time of skin incision (T2), after the transmitter had been secured to the body wall (T3), and at the end of surgery (T4). Body temperature, heart rate, respiratory rate, end-tidal carbon dioxide (ETCO2), and pulse oximetry values (SpO2) were recorded once each bird was intubated (T1), and at T2, T3, and T4. Respiratory rates were recorded before birds were removed from transport kennels and while they were being held for IV catheter placement.

Vaporizer settings did not differ between the two groups, with mean initial vaporizer settings at 4.2%, decreasing to 3.25% at T2, 2.5% at T3, and 0% at T4. The amount of time from induction to skin incision, time from skin incision until skin closure, and total anesthesia time did not differ significantly between the two groups. The time from the end of surgery to extubation was significantly shorter in the LV group (4±3 min) than in the HV group (10±2 min). Recovery times were also significantly shorter in the LV group (13±5 min) than in the HV group (19±4 min). Neither heart rate nor body temperature differed significantly between the groups at any time point. The resting respiratory rate did not differ between the two groups, nor did it differ when the birds were held for initial blood sampling. There was a significant decrease in respiratory rate in both groups from the time when they were held for blood sampling until induction. Respiratory rate differed significantly between the groups at T2 and at T4. Four HV birds stopped voluntary breathing while being ventilated. Oxygen saturation values did not differ between the two groups. End-tidal carbon dioxide values were significantly higher in the LV group at T2, T3, and T4, than in the HV group.

Both groups had an initial mild metabolic acidosis with normal partial pressures of carbon dioxide (PvCO2). Geese in the HV group had very little change in PvCO2 during anesthesia, while geese in the LV group experienced more severe acidosis and hypercapnia. More dramatic physiologic abnormalities would have probably occurred had birds received no assisted ventilation.4 Both isoflurane and high concentrations of inspired oxygen have been shown to be potent respiratory depressants in birds.6 These factors likely account for the decrease in respiratory rate, hypercapnia, and respiratory acidosis observed in the geese of this study.

Initial plasma lactate values were elevated, but these values decreased over time. The physiologic consequences of elevated lactate have not been well documented in wildlife, but it has been suggested that it may contribute to capture myopathy.2 Administration of 100% oxygen may have decreased the amount of anaerobic metabolism, thereby reducing lactate production. Lactate production may also have been diminished as the birds were anesthetized and became less responsive to stressful stimulation.

It is unknown why the partial pressures of oxygen (PvO2) were significantly lower in the HV group. While venous samples can provide useful information regarding PCO2 and acid-base status,1 interpretation of venous PO2 is difficult. Lower PvO2 values may have been caused by decreased oxygen delivery to the lungs, decreased uptake in the lungs, increased loss from the ruptured air sac, or increased uptake by body tissues. It would have been interesting to determine PaO2 values for the geese of this study; however, arterial catheterization can be difficult in birds and was not feasible for this investigation.

The most profound blood gas changes occurred between T0 and T2 in both groups. It was difficult to determine if this was due to coelomic air sac rupture because of effects related to the duration and depth of anesthesia. However, isoflurane requirements did not increase when the air sacs were ruptured and there were no clear patterns for changes in PvCO2 or ETCO2. It is possible that there are no physiologic consequences to disruption of the caudal abdominal air sac, but it is more likely that the time that the air sac is open to the outside during transmitter implantation is insufficient for a change to be detected.

Geese in the HV group took longer to recover than birds in the LV group. The two groups had no significant differences in the percentage of isoflurane administered at each time point, so it is likely that birds in the HV group received a higher frequency of deep breaths and, consequently, more isoflurane. Thus, the HV group was probably more deeply anesthetized. Furthermore, HV birds had lower PvCO2 values and lower pH values, which probably decreased the stimulation to breathe. This also could have contributed to longer recovery times as these birds had low respiratory rates during the initial minutes of recovery, thereby taking longer to eliminate the isoflurane.

Assisted ventilation should be performed in birds anesthetized for procedures such as surgical implantation of transmitters in order to approximate normal physiologic homeostasis. Blood gas analysis or capnography can be used for monitoring the adequacy of ventilation. The longer recovery times that we observed might be minimized by using lower concentrations of isoflurane and by continuing assisted ventilation until the birds are extubated.

Literature Cited

1.  Bailey, J.E. and L.S. Pablo. 1998. Practical approach to acid-base disorders. Vet. Clin. N. Amer. Small Anim. Pract. 28: 645–662.

2.  Dabbert, C.B. and K.C. Powell. 1993. Serum enzymes as indicators of capture myopathy in mallards (Anas platyrhynchos). J. Wildl. Dis. 29: 304–309.

3.  Korschgen, C.E., K.P. Kenow, A. Gendron-Fitzpatrick, W.L. Green, and F.J. Dein. 1996. Implanting intra-abdominal radiotransmitters with external whip antennae in ducks. J. Wildl. Manage. 60: 132–137.

4.  Ludders, J.W., G.S. Mitchell, and J. Rode1. 1990. Minimal anesthetic concentration and cardiopulmonary dose-response of isoflurane in ducks. Vet. Surg. 19:304-307.

5.  Mulcahy, D.M. and D. Esler. 1999. Surgical and immediate post-release mortality of harlequin ducks (Histrinonicus histrionicus) implanted with abdominal radio transmitters with percutaneous antennae. J. Zoo Wildl. Med 30: 397–401.

6.  Seaman, G.C. and J.W. Ludders. 1994. Effects of low and high fractions of inspired oxygen in ducks anesthetized with isoflurane. Am. J. Vet. Res. 55: 395–398.


Speaker Information
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R. Scott Larsen, DVM, MS
Environmental Medicine Consortium
Department of Clinical Sciences
College of Veterinary Medicine
North Carolina State University
Raleigh, NC, USA

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