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J Am Assoc Lab Anim Sci. Mar 2009; 48(2): 166–170.
Published online Mar 2009.
PMCID: PMC2679669

Successful Management of Rabbit Anesthesia Through the Use of Nasotracheal Intubation

Abstract

Although nasotracheal intubation in the rabbit has been briefly described, scientific assessment of the procedure has not been reported. In this report we describe nasotracheal intubation performed in 38 male New Zealand White rabbits (3.0 to 5.5 kg) used for a vascular patch study. The rabbits were placed under general anesthesia twice, with 2 mo between the initial and final intubations. Rabbits were intubated by the oral or nasotracheal route and compared. Previous literature dismissed nasotracheal intubation, citing the possibility of introducing pathogens into the lungs and the necessity of high oxygen flow rates (presumably greater than 3 L/min). However, no clinical signs of respiratory disease were noted among the study animals, nor were high oxygen flow rates necessary. Several key points collectively facilitated a successful procedure. Total relaxation was essential, modification of the classic blind approach eased placement, a correct approach was necessary, and our development of a unique method of securing the tube improved tube management. The findings suggest that nasotracheal intubation can be used as an easy, less traumatic method of rabbit intubation when compared with orotracheal intubation.

The exact number of rabbits represented by various rabbit industry groups is unknown, but the 2002 US Rabbit Industry Profile8 reported that more than 250,000 rabbits were used in research facilities and that 5 million pet rabbits were owned by 2.2 million US households. Rabbits are commonly anesthetized for both biomedical research and clinical veterinary medicine. Procedures that require anesthesia typically also require a patent airway, which can be difficult to obtain in rabbits. Many references7,10,14,20,21,24-26,29 describe rabbit intubation as being technically difficult due to specific anatomical characteristics including a long, narrow oropharynx, large tongue and incisors, and the limited mobility of the temperomandibular joint. Other references21,25,26,29 note the tendency of the rabbit larynx to spasm, and even successful endotracheal intubation can result in injury.9,21 Alternative methods of maintaining a patent airway in rabbits include the laryngeal mask airway, laryngeal tube, and airway device.6,11,13,25,31 Although the laryngeal mask airway and laryngeal tube may be easy to place, there is considerable potential of causing damage to their cuffs during placement. Additional laryngeal mask airway concerns are increased emission of anesthetic gas, lingual cyanosis, and the lack of a seal, which prevents its use during intermittent positive-pressure ventilation.13 The laryngeal tube was used during ventilation with and without neuromuscular blockade, but anesthetic gas emissions were not evaluated.31 The airway device, a prototype appliance, is similar to the laryngeal mask airway but designed specifically for small laboratory animals. This device was tested in 6 New Zealand White rabbits, and although the authors felt the device had many advantages, no firm conclusions were drawn because of the limited number of animals involved in the study.11

The difficulty perceived in intubating rabbits has spawned many methods of endotracheal intubation. Numerous methods have been described, ranging from the classic ‘blind’ method to the modified blind method, various surgical methods, videoendoscopic methods, and various laryngoscopic and otoscopic methods.1-3,6,7,10,12,13,16,18,21,24,26-30,32 The difficulty with orotracheal intubation may actually benefit nasotracheal intubation. Nasotracheal intubation takes advantage of the fact that the rabbit is an obligate nasal breather. Rabbits normally have their epiglottis entrapped on the dorsal surface of the soft palate, thus allowing direct passage of air from the nasopharynx into the larynx and trachea. A tube placed nasally will naturally traverse this pathway from the nasopharynx into the larynx and trachea. Although nasotracheal intubation has been described briefly as an alternative method of intubation in the rabbit, limited literature on this technique is available, and contraindications include the possibility of introducing pathogens into the lungs and need for high oxygen flow rates.10 Rabbits that received nasotracheal intubation at our facility were observed over 2 mo, and no clinical signs of respiratory disease noted. In addition, high oxygen flow rates were unnecessary. This report describes successful nasotracheal intubation of rabbits and several key points for appropriate clinical use of nasotracheal intubation.

Materials and Methods

General Procedures.

Male New Zealand White SPF rabbits (Oryctolagus cuniculus; n = 38; age, 6 mo; weight, 3.0 to 5.5 kg; Myrtles Rabbitry, Thompson Station, TN) were used. The rabbits were negative for bacterial (Bordetella bronchiseptica, cilia-associated respiratory bacillus, Pasteurella multocida, and Treponema cuniculi), viral (oral papilloma virus), helminthes (Passalurus ambiguus, Taenia pisiformis), protozoan (Eimeria stiedae, E. perorans, E. irresideum, and Encephalitozoon cuniculi), and arthropod (Psoroptes cuniculi, Cheyletiella parasitovorax) pathogens. All rabbits were housed individually in a stainless steel cage system under standard environmental conditions (20.2 ± 2 °C, 30% to 70% relative humidity, 12:12-h light:dark cycle, and 12 to 15 air changes hourly). Rabbits were allowed ad libitum access to a commercial rabbit diet (Lab Rabbit Breeder CS, PMI International, Brentwood, MO) and filtered water (Edstrom Industries, Waterford, WI). Research was conducted in compliance with the Animal Welfare Act and other federal statutes and regulations. This research adhered to principles stated in the Guide for the Care and Use of Laboratory Animals.19 All procedures used in this study were approved by the institutional animal care and use committee. The facility where this research was conducted is fully AAALAC-accredited.

Anesthesia.

The rabbits were placed under general anesthesia twice, with a 2-mo time span between the initial and final intubation. Rabbits were premeditated with buprenorphine (0.03 mg/kg SC; Reckitt Benckiser Pharmaceutical, Richmond, VA), ketamine hydrochloride (35 mg/kg IM; Fort Dodge Laboratories, Fort Dodge, IA), xylazine (5 mg/kg IM; Phoenix, St Joseph, MO), and glycopyrrolate (0.01 mg/kg SC; Baxter Healthcare, Deerfield, IL). After 5 to 10 min, isoflurane (3.0% with 1.5 L/min O2) anesthesia was given via facemask until the rabbit was anesthetized sufficiently, indicated by lack of muscle tone and loss of gag and pinnae reflexes. During the procedure, the rabbit's heart rate, peripheral O2 saturation, and temperature were measured with a pulse oximeter (Vet/Ox Plus 4800, Heska Corporation, Flamborough, Ontario, Canada) and recorded. The O2 saturation values were recorded at every 10 min, and the mean for each rabbit was calculated at the end of each procedure. The animals’ respiratory rates were determined manually and recorded.

Technique.

After adequate anesthetic depth was achieved, the face mask was removed, and each rabbit was intubated nasally with a clear, silicon, uncuffed, lubricated (KY Lubricating Jelly, Johnson and Johnson, Langhorne, PA) endotracheal tube (inner diameter, 2.0 to 2.5 mm; length, 14.5 cm; Mallinckrodt Medical, St Louis, MO, and Rusch, Duluth, GA). Initially the rabbit was positioned as for the classic blind technique.5,18 The rabbit's head was held by the handler's subordinate hand so that the rabbit was facing the handler. Using the index finger and thumb, the handler grasped the maxillary arches of the rabbit's dentition, and the rabbit was suspended with its head in dorsoflexion, nose pointing up, and front feet just touching the table. The operator's dominant hand then introduced the tube. However, noting that human pediatric patients typically are intubated while they are supine or dorsal, suggested that rabbit intubation in this presentation might be preferable.

Intubation was found to be relatively easy when the rabbit was dorsally recumbent. Using the index finger and thumb of the subordinate hand, the handler grasped the rabbit by the maxillary arches. The rabbit's head was still in dorsoflexion, but its body was now in dorsal recumbency, so that the head was lifted while the lower back was on the table (Figure 1), The nasal fold was lifted, and the tube was advanced medially and ventrally toward the nasal septum and hard palate.6 The tube was checked frequently during placement for condensation. If resistance was felt during insertion, the operator pulled back the tube and reinserted it while watching for condensation. After successful intubation, a small (1 to 2 cm) piece of tape was wrapped around the free end of the tube so that a small portion extended beyond each side. The tape was sutured to the dorsum of the nose by using 4-0 prolene (Ethicon, Piscataway, NJ). This process was repeated on the opposite side so that, when complete, the tube was secured from both sides of the tape to the dorsum of the rabbit's nose (Figure 2). Although the distance from nares to glottis was not measured, a 14.5-cm endotracheal tube seemed sufficient in length; the distance from upper incisor to glottis is about 9 cm in a 2.5- to 4.0-kg rabbit.14 The endotracheal tube entered the nares, traveled through the ventral nasal meatus, through the choana and nasal pharynx, and then passed into the trachea (Figure 3).

Figure 1.
Positioning of the rabbit for nasotracheal intubation.
Figure 2.
Tube secured and sutured to the rabbit. Gauze was used to stabilize and secure the head position.
Figure 3.
Illustration of nasotracheal intubation in the rabbit. The endotracheal tube enters the nares; travels through the ventral nasal meatus, choana, and nasal pharynx; and then passes into the trachea. Anatomic features were adapted from reference 23.

During surgery, the rabbits were maintained under general anesthesia (3.0% isoflurane with 1.5 L/min O2) with spontaneous respiration and were positioned in dorsal recumbency on a circulating water blanket with their heads in dorsoflexion to expose the ventral neck region. One of 2 surgical procedures was performed. The initial surgical procedure was to place a vascular patch on the carotid artery; the terminal surgical procedure was to harvest the patch and surrounding carotid artery 2 mo after patch placement. The anesthesia times for the surgical procedures were approximately2.0 and 0.5 h, respectively.

After completion of the initial carotid patch procedure, animals were recovered from general anesthesia. Isoflurane administration was terminated, and supplemental oxygen was administered for 5 m. The rabbit was moved from the surgery suite to the recovery area, the sutures were removed from the nares, and the rabbit was placed in sternal recumbency for recovery. Recovery was rapid and unremarkable. The tube was removed as the rabbit became responsive. Once the rabbit was able to make purposeful movement, it was returned to its cage. In the terminal procedure, the rabbits were euthanized (0.22 ml/kg IV; 390 mg pentobarbital sodium and 50 mg phenytoin sodium per milliliter; Euthasol, Virbac Corporation, Fort Worth, TX) immediately after harvest while they were still under general anesthesia.

Data analysis.

Statistical analysis was conducted by using Systat software (SSI, San Jose, CA). The t test was used to assess whether mean SpO2 differed significantly between the orotracheal and nasotracheal routes (P > 0.050).

.

Results

During the first group of intubations, 7 of the 38 rabbits were intubated orally. However, because difficulties with orotracheal intubation were encountered in numerous animals, intubations of the remaining 31 animals were performed by using a nasotracheal approach. Of the 75 successful intubation attempts in the 38 rabbits, 68 were accomplished nasally. During the 2-mo time span between procedures, no clinical signs of respiratory disease in the rabbits were noted.

Of the 38 rabbits, 1 (2.6%) was found dead in its cage 15 d after surgery. The rabbit had been orally intubated; clinical signs of respiratory disease were not observed prior to death, and gross indications of respiratory disease were not detected during necropsy. The cause of death is unknown but appeared unrelated to the intubation technique.

The O2 values were reviewed after completion of the protocol, and the mean for nasotracheal intubation was compared with that for orotracheal intubation. The mean O2 saturation of the 68 nasotracheal intubations (94.2 mm Hg) did not differ significantly (P = 0.250) from that of the 7 orotracheal intubations (94.2 mm Hg). During the second (terminal) procedure, nasotracheal intubation was performed in the 6 animals that initially had undergone orotracheal intubation. The mean O2 saturation of these 6 nasotracheal intubations (95.0 mm Hg) did not differ significantly (P = 0.250) from that of orotracheal intubation in these animals (96.1 mm Hg). The data passed tests for normality (P > 0.050) and equal variance (P = 0.825).

Discussion

The results from the current study suggest that nasotracheal intubation can be used as a humane, clinically preferred method of rabbit intubation. The first 7 rabbits were intubated by using the oral route, but after numerous difficulties were encountered, nasotracheal intubation was performed on the remaining subjects. Complications of orotracheal intubation included difficult placement by the oral route, trauma to the oropharangeal soft tissue, and assuring consistent placement and maintenance during a carotid artery procedure. Oral placement required using surgical gauze (and sometimes surgical tape) to tie the tube around the back of the rabbit's head. The surgeon, who was seated during the procedure, sometimes leaned into the patient, causing tube movement and dislodgement. These difficulties led to nasotracheal intubation as a method to provide an improved clinical solution for humane patient management.

A previous report in literature dismissed the utility of nasal intubation, citing the possibility of introducing pathogens into the lungs and the necessity of high oxygen flow rates (presumably greater than 3 L/min).10 Clinically, neither item was an issue during this study. Of the 38 rabbits, 1 (2.6%) was found dead in its cage 15 d postoperatively. That rabbit had been orally intubated, but no clinical signs of respiratory disease were detected prior to death and no gross indications of respiratory disease were detected during necropsy. During the 2-mo time span between procedures, none of the remaining 37 rabbits (including the 31 animals that had been nasally intubated) showed any clinical signs of respiratory disease. A leading cause of upper respiratory disease in the rabbit is Pasteurella multocida. Although pasteurellosis can cause disease in various organ systems, the respiratory form in rabbits tends to present as rhinitis, sinusitis, and pneumonia. Rhinitis (‘snuffles’) is the most common clinical manifestation and may present as serous to mucopurulent nasal discharge, sneezing, and coughing.4,26 Our SPF rabbits were free of Pasteurella, Bordetella, and cilia-associated respiratory bacillus on receipt from the vendor, but infected rabbits can be clinically asymptomatic, and even deep nasal swabs may fail to detect the organisms in all rabbits.22 Chronic pneumonia in rabbits also can be asymptomatic. In a research setting, rabbits may exhibit few clinical signs because of the minimal respiratory demands of being in a cage. Even though our rabbits remained in their cages the majority of the time, they were intubated and did undergo a surgical procedure. Although this stress might have been sufficient to exacerbate an existing asymptomatic infection (if present), nasal and lung cultures and histology were not obtained from these animals. Future research should focus on the pathology of nasotracheal intubation, whether pathogens can be seeded from the nasal passages, and whether nasotracheal intubation causes damage to the nasal passages.

One report10 discussed the use of small (inner diameter, 1.0 to 1.5 mm) nasal tubes positioned in the nasal passages of rabbits. The authors of that study felt that the technique required high flow rates (presumably in excess of 3 L/min) to create positive pressure and force the anesthetic mixture into the nasopharynx.10 In the current study, the rabbits were intubated easily nasally with uncuffed, lubricated, endotracheal tubes measuring 14.5 cm in length and 2.0 to 2.5 mm in inner diameter. After intubation the rabbits were maintained under general anesthesia (3.0% isoflurane with 1.5 L/min O2) with spontaneous respiration for 0.5or 2.0 h. This O2 rate was well within the range of fresh gas flow rates (1 to 3 L/min) reported for the maintenance of spontaneously breathing rabbits.15 The ability to maintain the 1.5 L/min O2 flow rate may have been due to the size of the endotracheal tube and the location of the tube in the tracheal passage instead of the nasal passages. A review of the O2 saturation levels in the study animals indicated that the percentages were in the mid-90s and were similar for both procedures. A t test of the mean nasal and oral O2 saturation levels (nasal, 95.0 mm Hg; oral, 96.1 mm Hg) within the same subgroup of rabbits showed no statistically significant difference between the procedures. Invasive monitoring with arterial blood gas analysis would have been an absolute way to effectively judge the adequacy of ventilation and oxygenation, but because the associated surgical procedure did not require invasive monitoring of oxygen saturation, pulse oximeters, a standard noninvasive method, were used. Pulse oximeters indicate the level of oxygen saturation of hemoglobin in the blood and display the saturation of peripheral oxygen. Adequate arterial oxygenation requires a value of at least 60 mm Hg, which corresponds to 90% saturation of the hemoglobin in the arterial blood.17 Another noninvasive method to judge the adequacy of ventilation is capnography, the measurement of the concentration of CO2 in inspired and expired gas. Whereas inspired air contains virtually no CO2, exhaled air has CO2 from alveolar emptying, thus providing information on pulmonary perfusion and alveolar ventilation.17 The anesthesia monitoring system for the present study was not set up to measure CO2; however this information may be captured in future procedures to help ensure proper tube placement.

During development of the nasotracheal intubation technique, several key criteria emerged as being necessary for safe and effective nasotracheal intubation. Total relaxation of the rabbit was essential, modification of the classic blind position eased placement, a correct approach was necessary, and a unique method of securing the tube improved tube management. Total relaxation of the rabbit was achieved by premedication with ketamine and xylazine and by administering isoflurane in O2 by face mask until the rabbit lacked muscle tone and had lost the gag and pinnae reflexes. In addition, modifying the rabbit's position improved the ease of nasotracheal intubation. The rabbit's head was held in dorsoflexion during dorsal recumbency. The correct approach entailed gentle insertion of the tube into the nares medially and ventrally toward the nasal septum and hard palate. Finally, the shape of the rabbit's head and the dorsoflexion position of the tube discouraged using gauze to secure the tube. In contrast, the modified tape and suture method for securing the tube did not result in a single tube displacement.

The present study described the development of a nasotracheal technique for rabbits, compared the orotracheal and nasotracheal techniques, and identified nasotracheal intubation as the preferred methodology for New Zealand White rabbits. Although nasotracheal intubation has been thought to present a risk for introducing pathogens into the lungs and to require high oxygen flow rates, neither concern was realized in the present study, but additional research is needed to determine the pathology of nasotracheal intubation. During the 2-mo time span between intubations, no clinical signs of respiratory disease were noted, and typical flow rates delivered sufficient oxygen to the rabbits.

Rabbit intubation has long been described as being technically demanding and time-consuming; numerous articles describe diverse techniques. As described in this report, the use of nasotracheal intubation and the novel method of securing the endotracheal tube have several benefits over traditional orotracheal intubation of rabbits. Nasotracheal intubation provides minimal trauma, unlike previously reported laryngeal injuries from orotracheal intubation.8,21 Placing the endotracheal tube nasally affords the surgeon increased access to the oral cavity without interference from an endotracheal tube. In addition, suturing the tube to the rabbit's nose facilitates maintaining the airway, allowing the surgeon to work without fear of dislodging the tube. After a few technical modifications and some practice attempts, operators preferred nasotracheal intubation as an easy, atraumatic method of rabbit intubation. The benefits of nasotracheal intubation were the ease of performing the procedure, lack of complicated equipment and supplies, and the lack of rabbit complications. Clinical observations of the intubated animals suggest that nasotracheal intubation can be used as a humane, atraumatic alternative method for safe and effective rabbit intubation.

Acknowledgments

The author thanks Malcolm Kling and Ron Banks for editorial support; Eugene Cauley, Leticia Simon, and David Robinson for technical assistance; John Tsai for support as the principal investigator; Jordan T Mastrodonato for help with the figures; and James McPherson for assistance with statistical analysis. Opinions, interpretations, conclusions, and recommendations are those of the author and are not necessarily endorsed by the US Army.

A technical abstract regarding nasotracheal intubation of rabbits was submitted and accepted for presentation at the 58th National Meeting of the American Association for Laboratory Animal Science and the 2009 AVMA Laboratory Animal Medicine Program.

References

1. Bechtold SV, Abrutyn D. 1991. An improved method of endotracheal intubation in rabbits. Lab Anim Sci 41:630–631 [PubMed]
2. Corleta O, Habazettl H, Kreimeier U, Vollmar B. 1992. Modified retrograde orotracheal intubation techniques for airway access in rabbits. Eur Surg Res 24:129–132 [PubMed]
3. Davies A, Dallak M, Moores C. 1996. Oral endotracheal intubation of rabbits (Oryctolagus cuniiculus). Lab Anim 30:182–183 [PubMed]
4. Deeb BJ. Respiratory disease pasteurellosis. Queensberry KE, Carpenter JW, editors. Ferrets, rabbits, and rodents: clinical medicine and surgery. 2nd ed.Philadelphia (PA): WB Saunders; 2004. p172–182
5. Flecknell P. Anaesthesia. Flecknell P, editor. BSAVA manual of rabbit medicine and surgery, 2nd ed.Gloucester (UK): British Small Rabbit Veterinary Association Publishing; 2006. p 110
6. Flecknell P. Anaesthesia and perioperative care. Meredith A, Flecknell P, editors. BSAVA manual of rabbit medicine and surgery, 2nd ed.Gloucester (UK): British Small Rabbit Veterinary Association Publishing; 2006. p 154
7. Gografe SI, Wilson JS, Johnson BL, Rushing G, Bowser A, Parker JL, Cornelius CE. 2003. Successful management of long-term general anesthesia in rabbits used as an animal model of human disease. Contemp Top Lab Anim Sci 42:16–19 [PubMed]
8. Grannis J. US rabbit industry profile. Fort Collins (CO): Centers for Epidemiology and Rabbit Health, Center for Emerging Issues; 2002
9. Grint NJ, Sayers IR, Cecchi R, Harley R, Day MJ. 2006. Postanaesthetic tracheal strictures in three rabbits. Lab Anim 40:301–308 [PubMed]
10. Harcourt-Brown F. Anaesthesia and analgesia. A textbook of rabbit medicine. Oxford (UK): Alden Press; 2001. p 121
11. Imai A, Eisele PH, Steffey EP. 2005. A new airway device for small laboratory rabbits. Lab Anim 39:111–115 [PubMed]
12. Irazuzta J, Hopkins J, Gunnoe P, Brittain E. 1997. Simple method of multipurpose airway access through percutaneous tracheotomy in rabbits (Oryctolagus cuniculus). Lab Anim Sci 47:411–413 [PubMed]
13. Kazakos GM, Anagnostou T, Savvas I, Raptopoulos D, Psalla D, Kazakou IM. 2007. Use of the laryngeal mask airway in rabbits: placement and efficacy. Lab Anim (NY) 36:29–34 [PubMed]
14. Krüger J, Zeller W, Schottmann E. 1994. A simplified procedure for endotracheal intubation in rabbits. Lab Anim 28:176–177 [PubMed]
15. Lipman NS, Marini RP, Flecknell PA. Anesthesia and analgesia in rabbits. Kohn DF, Wixson SK, White WJ, Benson GJ, editors. Anesthesia and analgesia in laboratory animals. New York (NY): Academic Press; 1997. p 205
16. Macrae DJ, Guerreiro D. 1989. A simple laryngoscopic technique for the endotracheal intubation of rabbits. Lab Anim 23:59–61 [PubMed]
17. Mason DE, Brown MJ. Monitoring of anesthesia. Kohn DF, Wixson SK, White WJ, Benson GJ, editors. Anesthesia and analgesia in laboratory animals. New York (NY): Academic Press; 1997. p 73
18. Morgan TJ, Glowaski MM. 2007. Teaching a new method of rabbit intubation. J Am Assoc Lab Anim Sci 46:32–36 [PubMed]
19. National Research Council 1996. Guide for the care and use of laboratory animals. Washington (DC): National Academy Press
20. Patil VU, Fairbrother C, Dunham BM. 1997. Endotracheal intubation in the rabbit: a quick reliable method. Lab Anim 26:28–29
21. Phaneuf LR, Barker S, Groleau MA, Turner PV. 2006. Tracheal injury after endotracheal intubation and anesthesia in rabbits. J Am Assoc Lab Anim Sci 45:67–72 [PubMed]
22. Percy DH, Barthold SW. Pathology of laboratory rodents and rabbits, 2nd ed.Ames (IA): Iowa State Press; 2001
23. Popesko P, Rajtová V, Horák J. A colour atlas of the anatomy of small laboratory animals, plate 25, p 38 London (UK): Saunders; 2002
24. Rukavina G. 2007. Step by step: guided endotracheal intubation in rabbits. Tech Talk. 12:1–3
25. Smith JC, Robertson LD, Auhll A, March TJ, Derring C, Bolon B. 2004. Endotracheal tubes versus laryngeal mask airways in rabbit inhalation anesthesia: ease of use and waste gas emissions. Contemp Top Lab Anim Sci 43:22–25 [PubMed]
26. Suckow M, Brammer DW, Rush HG, Chrisp CE. 2002. Biology and diseases of rabbits, p 329 Fox JG, Anderson LC, Loew FM, Quimby FW, editors. Laboratory rabbit medicine, 2nd ed.San Diego (CA): Academic Press
27. Tran HS, Puc MM, Tran JL, Del Rossi AJ, Hewitt CW. 2001. A method of endoscopic endotracheal intubation in rabbits. Lab Anim 35:249–252 [PubMed]
28. Weinstein CH, Fujimoto JL, Wishner RE, Newton PO. 2000. Anesthesia of 6-week-old New Zealand White rabbits for thoracotomy. Contemp Top Lab Anim Sci 39:19–22 [PubMed]
29. Wixson SK. Rabbits and rodents: anesthesia and analgesia. Hampshire V, Gonder JC, editors. Research animal anesthesia, analgesia and surgery. Greenbelt (MD): SCAW; 2007. p 53
30. Worthley SG, Roque M, Helft G, Soundararajan K, Siddiqui M, Reis ED. 2000. Rapid oral endotracheal intubation with a fibre-optic scope in rabbits: a simple and reliable technique. Lab Anim 34:199–201 [PubMed]
31. Yamamoto Y, Inoue S, Abe R, Kawaguchi M, Furuya H. 2007. Airway management with the laryngeal tube in rabbits. Lab Anim 36:33–35 [PubMed]
32. Yurevich S. 2002. Blind intubation in rabbits. Veterinary Technician 23:291–293

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