Abstract

The use of flexible fiberoptic endoscopes to examine the respiratory tract of Bottlenose dolphins, Tursiops truncatus, and in diagnostic procedures such as bronchoalveolar lavage (BAL) has been recently reported.^1,3,4,7 As respiratory endoscopy becomes more commonplace in cetacean clinical medicine it is important to recognize normal anatomy and findings, to differentiate from abnormal, and to identify potential pitfalls of the technique.

There has been, and remains, considerable controversy over the exact source and mechanisms behind odontocete sound production. Early hypotheses based largely on dissection included the uniquely modified larynx and various components of the equally unique supracranial air sac structures as proposed sound sources.² Pressure profile metry work at our laboratory in the 1970's effectively ruled out the larynx as the sound source and strongly suggested the primary site(s) lies superior to the level of the bony nares.⁸ A recent hypothesis based on computed tomography scanning points to paired structural complexes, within the nasal passages, termed the Monkey Lips Dorsal Bursae (MLDB) complex, as a potential primary source of sound production.

To investigate the hypothesis that the MLDB complex is a key component in odontocete sound production an investigation is in progress at our laboratory using small diameter flexible fiberoptic endoscopes and high speed (400 Hz) video recording equipment to image upper respiratory structures of phonating and echo locating dolphins at the surface and underwater. An offshoot of this project has been considerable experience in imaging the various regions of the respiratory tract and a large data set of video images. This information has been useful in the clinical management of dolphins with suspected or confirmed upper respiratory disease or abnormality. This paper presents normal dolphin upper respiratory endoscopic technique and anatomy, related relevant cytology, functional observations, and examples of pathology. Observations during endoscopic examination of the upper respiratory tract of the gray whale, Eschrichtius robustus, both ante-mortem and postmortem, are presented for comparative purposes.

Technique

The entire upper respiratory tract, with the exclusion of the premaxillary and nasofrontal sacs, can be effectively imaged using a 3.5 mm outside diameter x 70 cm working length flexible fiberoptic endoscope and standard light source. The ability of the distal tip to flex 180° in one direction greatly enhances the examination by allowing retroflexed views of the inferior surfaces of the choanae and nasal plug. Endoscopes as large as 12.8 mm outside diameter have been successfully used to examine the upper respiratory tract but retroflexion is limited; use with the blowhole under water is not recommended and repeated use should be limited. Recording all examinations with a video-camera adapter and VCR is highly recommended. Post-procedure review often reveals significant observations that may be missed during the procedure when attention is focused on the well being of the patient. An archive of video images is also invaluable for inter and intra-patient comparisons.

Dolphins and whales can be trained to accept an endoscope into the nasal passages while at a hand station in the water and we have trained two animals to maintain the scope in place while positioned on a bite plate one meter below the water surface. Most clinical cases win not have had this training, however, our experience with small bore scopes is that most animals accept the examination after minimal initial objection. Light sedation with oral diazepam is useful in those animals likely to object more strongly. Dialogue with the animal's primary handler and senior training staff is essential to good case management. An untrained animal, with no experience with upper respiratory endoscopy, should not be examined in the water. These animals should be removed from the water and restrained in sternal recumbency. Adequate personnel, familiar with handling cetaceans out of the water and patient monitoring equipment should be available. Sterilization of the endoscope is recommended, and essential if the scope is to be introduced through the glottis. Thorough rinsing of the scope with sterile saline should be performed to avoid introducing potentially irritating disinfection solutions into the nasopharynx or interference with microbiology isolation attempts.

For initial examination the endoscope is grasped approximately 6-8 cm from the distal tip, between the thumb and forefinger of the dominant hand of the endoscopist and the tip positioned over the blowhole. The blowhole opening is viewed as a "U" shape, with the top of the "U" oriented rostrally. The orientation of the scope image to the blowhole opening should be noted. This will serve as an aid to orientation of the deeper structures, especially for the inexperienced endoscopist. As the blowhole is opened, during a breath, the scope is introduced into the left or right nasal passage with a deliberate movement and the operating hand braced against the animal's melon to prevent damage to the scope or trauma to the animal should it move in objection to the scope presence. Initial introduction takes practice and patience. Often the scope tip is deflected.

Once the scope is in place in the blowhole proper and the animal accepts the procedure, the scope is advanced to a depth of 12-15 cm. At this point the nasopharynx can be imaged, and with practice, the scope tip can be directed caudally and medially to view the glottis, apex of the cuneiform and epiglottic cartilages, and surrounding palatopharyngeal muscle complex.

Indiscriminate advancing of the scope is to be avoided. The scope may easily pass along side the larynx and be misdirected entering the oropharynx (it is possible to view, if required, the ventral aspect of the larynx and the oropharynx by introducing the scope orally). A moderate amount of foamy white secretion is often present in the nasopharynx of clinically normal dolphins. Retroflexion of the scope brings the inferior surface of the paired choanae, separated by the boney nasal septum, into view and each distal nasal passage can be inspected separately. Along the lateral walls of the nasopharynx and into the depths of the nasopharyngeal fornix, crypt openings of multiple mucous glands are visible At the level of the choanae, on the lateral wall of the nasopharynx, the right and left Eustachian tube openings can be inspected. These are best viewed with the scope tip straight and retracted to a level just superior to the choanae.

The spiracular cavity is best inspected by slow retraction of the scope from depth as the tip is directed alternately laterally and medially. The inexperienced patient will often chuff during manipulation of the scope in the spiracular cavity, especially as the MLDB complex is approached. The spiracular cavity can also be viewed by retracting the scope while it is in the full retroflexed position, however, caution must be exercised to avoid hanging up on the nasal septum or diagonal membrane and little advantage is offered. Examination of the spiracular cavity is hindered by tissue collapsed over the distal optics of the scope between respirations, i.e., when the "blowhole is closed." This can be overcome by placing a clear flexible sheath or "standoff' of a diameter slightly larger than the scope over the distal end. The standoff displaces tissues from the distal optics and allows a small degree range of motion of the scope. It is hard to image a clinical situation that would require this but it does have research applications and has been successfully employed in our dynamic imaging of the MLDB complex.

Examination of the supracranial air sacs is realistically limited to the vestibular sacs. They are easily entered by directing the scope laterally just below the blowhole opening along the right or left side when initially introducing the scope. Most patients do not object to the scope in the vestibular sac, in fact many attempt to manipulate the scope tip into this area during examinations of the spiracular cavity. The entire interior mucosal surface of the sacs can be examined. Infusion with small amounts of sterile saline can be used to distend the sacs and facilitate inspection. Our experience with post mortem dissections suggests it is not currently possible to intubate the nasofrontal sacs in the live dolphin. We also conclude it may be possible to examine the premaxillary sacs but it is unlikely to be clinically beneficial and to date has not been attempted.

Observations

Early hesitations by marine mammal clinical veterinarians at performing respiratory endoscopy were in large part due to the uncertainty of potential compromise to the effective seal of the blowhole/nasal plug mechanism. Certainly the unique cetacean respiratory cycle, with prolonged apneustic plateau and breath holding, is a beneficial adaptation to aquatic life. Concerns over compromise to the respiratory pattern, with resultant ventilation and potential acid base disturbances, were appropriate. Concern over sea water aspiration of an animal in the water is valid. Experience with monitored animals has shown us that upper respiratory endoscopy, when performed as above, is well tolerated by most patients and has not affected core body temperature, ECG patterns, or pulse oximetry readings. Our in-water investigations have additionally brought to light several aspects of the functional anatomy that allow this to be a safe, clinically valuable procedure.

The exquisitely complex groups of muscles surrounding the upper airways of the dolphin provide multiple well controlled physical barriers to the passage of air or fluid through the airways between breaths.⁶ The tissues superior to the level of the MLDB complex are very muscular and are held in tight apposition to each other as are the MLDB complexes themselves. The nasal plug, when "closed," is held tightly against the posterior aspect of the boney nares and diagonal membrane. The palatopharyngeal muscle complex can constrict down to completely close over the apex of the larynx. And, the lumen of the larynx in the region of the elongated cuneiform and epiglottic cartilages collapses the entire length, bringing the mucosal surfaces in contact with each other for the full length, as opposed to merely at the glottis as in terrestrial domestic species.

Two other key adaptive mechanisms function to avoid the potential complications discussed above. Pressure studies mentioned previously, and experimental observations with animals undergoing under-water endoscopy, show that the pressure in the nasopharynx remains above ambient pressure at all times between breaths. Thus a dolphin, under the water surface, that experiences a breach of the upper respiratory tract will merely "leak air" rather than aspirate water. This is evident from pressure profilemetry as wen as the clinical observation of occasional bubble streams from the blowhole of submerged dolphins. Endoscopic observations suggest the animal may "recharge" the nasopharynx with additional air by opening the glottis and exhaling if necessary, although concurrent profilemetry during this observation has not been performed.

Lastly, we have observed that dolphins, under the water surface, when infused with a 60 cc volume of fluid via an endoscope placed into the nasopharynx simply swallow the fluid by activity of the palatopharyngeal muscle complex with a closed glottis. All above mechanisms are expected by the animal's requirement to exist as an exclusively aquatic mammal and should not surprise us.

Functional movements of the structures of the nasopharynx and upper passages are consistent between animals and should be evaluated. The apex of the larynx is readily observed opening during breaths. Extrinsic muscles of the larynx also pull it in a cranio dorsal direction during inspiration, however, we have not observed it to insert into the choanae as has been suggested. If the laryngeal apex is viewed while the animal is fed and swallows, remarkably little motion is observed. In fact it may not be possible to tell when the animal swallows a fish. Detailed systematic observations during deglution have not been carried out but there is no reason to believe that our observations are different than the usual course of events. There has been some controversy as to whether or not dolphins displace the larynx into the oropharynx during swallowing. Our observations confirm that the Atlantic Bottlenose dolphin is able to swallow food items without displacing the larynx. There are anecdotal reports of an Orca that was trained to chuff a squid out of it's blowhole on command after swallowing it. This would indicate that at least some cetaceans may be able to voluntarily displace the larynx.

During sound production the walls of the nasopharynx constrict by action the palatopharyngeal muscle complex and the opening to the Eustachian tube will often simultaneously widen. We have successfully introduced the scope into the Eustachian tube by first introducing a cytology brush through the biopsy channel and following it with the scope. Clinical indications for this procedure have not been identified and we can not recommend it be included in routine upper respiratory endoscopic surveys. With the scope tip retroflexed the inferior surfaces of the nasal plug can be inspected. Although not observed by the authors, it is anticipated that neurologic or other functional deficits of the plug opening would easily be observed by comparing the symmetry between left and right. During phonation the, lateral aspect of the nasal plug is often seen to move. The MLDB complex is a very dynamic area during phonation. Functional pathology of this area has not been observed and is unlikely to present in most clinical settings.

Samples for cytology or microbiologic analysis are easily obtained by standard endoscopic brush or wash techniques. The inexperienced endoscopist should consult references if unfamiliar with these.⁵ Grasping forceps may also be used to obtain larger volumes of exudate for cytology or tissue biopsies if indicated. Video-microscopic evaluation of exudates and secretions from clinical animals at our laboratory have revealed numerous organisms. Most of these are interpreted as commensals, however, the recent finding of an unidentified ameboid organism in a pleural effusion and ciliated protozoans in multiple somatic lymph nodes from dolphins at this lab have raised the issue that the nasal flora may have been the source. Further work in this area is badly needed and ongoing. When examining wet mounts of nasopharyngeal washes ambient temperature sea water provides better preservation of organism morphology and motility than saline or fresh water.

The bacterial and fungal flora of the upper respiratory passages is diverse. The clinical use of swabs or "chuff plates" has been common practice in marine mammal facilities in the past. The significance of isolated microbes from the nasal passages of dolphins is usually unknown and often indiscriminately used in clinical management. Endoscopic visualization and specific sample site selection is expected to lead to more significant findings. Additional work on the normal flora of the nasopharynx, nasal passages, and supracranial air sacs is needed.

Repeat localized endoscopy of the MLDB complex of two dolphins has lead to localized mucosal erosion of the tissue just superior to the MLDB complex corresponding to the location of the tip of the endoscope or stand off during imaging. This can be prevented by reducing the frequency of observations especially when larger diameter endoscopes are used. Light lubrication of the scope or stand off tip with an insoluble lubricant (mineral oil or petrolatum) may also be useful. We have not observed any erosion or ulceration subsequent to routine clinical examinations.

The mucosa overlying the laryngeal cartilages is vulnerable to trauma induced by scope manipulations. It is not uncommon to observe hyperemia of the mucosa that has contacted the scope while retracting the scope at the conclusion of an evaluation. This same phenomenon has been observed after BAL at the site the scope is wedged in the bronchiole. We have not observed post-procedure laryngeal edema or dyspnea however this possibility must be considered. Inexperienced endoscopists attempting to enter the glottis may also easily pass the scope into the oropharynx around the larynx with the possibility of traumatizing the intrinsic muscles of the larynx or their innervation. Laryngeal hemiparesis has not been described in dolphins but is theoretically a possible iatrogenic complication.

Conclusion

The performance of respiratory tract endoscopy on cetacean patients using a variety of scope configurations is likely to become more commonplace as marine mammal veterinarians advance clinical capabilities. The systematic complete evaluation of the nasopharynx and adnexa, laryngeal apex, nasal passages, nasal plugs, MLDB complex, and vestibular sacs has been shown to be a safe, effective clinical and research tool. When repeat evaluations are needed, animals can be trained to voluntarily accept placement of endoscopes into the upper respiratory structures and examinations can be performed with animals in or under water. Additional investigations into the normal cytology and microflora of the upper respiratory tract of cetaceans are needed to improve clinical correlations with endoscopic findings.

References

1.  Chaplin, M., T. Kamolnick, W. Van Bonn, D. Carder, S. Ridgway, and T. Cranford. 1996. Conditioning Tursiops truncatus for nasal passage endoscopy. Proceedings TA.

2.  Cranford, T.W., M. Amundin, and K. S. Norris. 1996. Functional morphology and homology in the odontocete nasal complex: Implications for sound generation. Journal of Morphology 222:223-285.

3.  Harrel, J.H., T.H. Reiderson, J. McBain, and H. Sheetz. 1996. Bronchoscopy of the bottlenose dolphin. Proceedings IAAAM 27th Annual Conference.

4.  Hawkins, E.C., F. I. Townsend, G.A. Lewbart, M.A. Stamper, V.G. Thayer, and H.L. Rhinehart. 1996. Bronchoalveolar lavage in a stranded bottlenose dolphin. Proceedings IAAAM 27th Annual Conference.

5.  Jones, B.D., (ed). 1990. Veterinary Endoscopy. The Veterinary Clinics of North America: Small Animal Practice.

6.  Lawrence, B., and W.E. Schevill. 1965. Gular musculature in delphinids. Bull. Mus. Comp. Zool. (Harvard) 133:1 pp. l-65.

7.  Reiderson, T.H., J. McBain, and J.H. Harrel. 1996. The use of bronchoscopy and fungal serology to diagnose Aspergillus fumigatus lung infection in a bottlenose dolphin. Proceedings IAAAM 27th Annual Conference.

8.  Ridgway, S.H., and D.A. Carder. 1988. Nasal pressure and sound production in an echo-locating white whale (Delphinapterus leucas). In P.E. Nachtigall and P.W.B. Moore (eds). Animal Sonar Systems: Processes and Performance. New York: Plenum Publishing Corporation, New York, Pp. 53-60.