Abstract

General anesthesia monitoring of the bottlenose dolphin remains a significant challenge stemming from the animals' unique anatomic and physiologic adaptations to an aquatic existence. These adaptations make it difficult to apply many of the anesthesia monitoring techniques commonly used in terrestrial animals to dolphins. We believe that some of these challenges may be mediated by working with non-anesthetized dolphins--developing techniques for monitoring and gaining familiarity with values collected from these healthy and alert animals--before applying the same monitoring program to animals under heavy sedation or general anesthesia. To that end, we've had the opportunity to monitor dolphins participating in research projects underway at the Navy Marine Mammal Program that require animals to be out of the water and in sternal recumbancy for various periods of time. Our physiologic monitoring program thus far includes collecting rectal temperatures, pulse-oximetry, electrocardiography, capnography, central venous blood pressure, and blood gas and co-oximetry data.

We use a Datex S/5 Anesthesia Monitor (Datex-Ohmeda, Madison, WI) to collect most of our data. The modular unit is designed for use in the human medical field, and for our purposes is configured for data collection from multiple sources including airway gases, electrocardiogram, oscillometric blood pressure, two lines of invasive blood pressure, and two temperature sources. The machine also has pulse-oximetry and respirometry capabilities that we plan to apply to future procedures. The Datex-Ohmeda S/5 Collect 4.0 software program allows us to record all information collected by the anesthesia monitor during the procedure and to save those data for review.

Regularly scheduled procedures have allowed us to develop a systematic approach to the placement of monitoring equipment. After diazepam premedication (0.1-0.2 mg/kg), an animal is transported into the veterinary surgical suite. It is placed in sternal recumbancy on closed-cell foam padding and supported in this position by training staff for the duration of the procedure. For placement of monitoring equipment, the animal is temporarily rolled into left lateral recumbancy and gently restrained while a temperature probe is placed rectally to a standard depth of 15 cm, and a reflectance pulse-oximetry probe is placed in the genital slit. The pulseoximetry probe is attached to a HESKA Veterinary pulse-oximetry unit (HESKA, Fort Collins, CO), and in this location gives sporadic readings at best, however, the frequency of feedback from this location is better than any other site tested so far.

For central venous pressure monitoring, venous blood collection, and intravenous fluid and drug administration, a 5-French, 60 cm, polyurethane catheter (Instech Laboratories, Plymouth Meeting, PA) is placed in the brachiocephalic vein using a percutaneous, ultrasound-guided, through the needle technique. We use a customized, thin-walled, stainless steel needle (14-gauge, 16 cm) with an echogenic tip (Popper and Sons, Inc. New Hyde Park, NY) to access the vein. After catheter placement is complete and patency is well established, the animal is rolled back into sternal recumbancy. Suction-cup leads are placed around the animal's thorax for EKG measurement, and the animal's lead trainer is positioned at the head with a side-stream airway adapter to collect blowhole exhalations for expired airway gas analysis (ETCO2, FiO2, ETO2). Values recorded from 5 animals are presented in Table 1. Heart rates, rectal temperatures, and end-tidal CO2 levels we've recorded correlate well with those previously reported^2,3, while central venous pressures are similar to reference ranges published for terrestrial large animals.¹

Throughout the procedure, serial blood samples are collected from the brachiocephalic vein, and from other peripheral locations (fluke veins, peduncle, dorsal fin, and hemal arch) for comparison of blood gas data. Blood gases are measured with a Nova Stat Profile Critical Care Xpress (Nova Biomedical Corporation, Waltham, MA) blood chemistry, gas, and co-oximetry analyzer. While we are currently investigating arterial sites that can be consistently, percutaneously accessed for blood gas sampling, we have run some descriptive statistics on the samples we've collected centrally and from peripheral sites. Looking at oxygen saturation as an indicator of arterial (vs. venous or mixed arteriovenous) sample, a blood sample collected from the peduncle is significantly more likely to be arterial (p-value <.0001, Table 2).

These efforts have improved our anesthesia-monitoring plan as it is applied to dolphins. Several of these same animals have been induced to general anesthesia after base-line data was collected. Induction agents, such as propofol, have been given through the brachiocephalic catheter to induce anesthesia, and the animals were intubated and ventilated. The animals recovered uneventfully from these procedures. The data from these anesthetic events warrant further investigation and will be the focus of our continued efforts as we work towards the capability of regular clinical procedures performed on animals under general anesthesia.

Table 1. Values collected from 5 animals during physiologic monitoring. These animals received 0.1-0.2 mg/kg oral diazepam at least one-hour prior to recording.

Table 2. Oxygen saturation of blood samples collected from brachiocephalic vein, peripheral vessels (fluke, dorsal fin, and hemal arch) and peduncle.

References

1.  Muir WW, Hubbell JAE, Skarda RT, Bednarski RM. 2000. Patient Monitoring During Anesthesia. Handbook of Veterinary Anesthesia, 3rd Edition. Mosby, Inc. 250-283.

2.  Ridgway SH, McCormick JG, Wever EG. 1974. Surgical approach to the dolphin's ear. J Exp Zool. 188(3): 265-76.

3.  Van Elk CE, Epping N, Gans SJ. 2001, Pulmonary function measurements in dolphins using capnography. Vet Rec. 8; 149(10): 308-9.