Endotracheal intubation is the process of passing a tube through the larynx into the trachea, in order to maintain a clear airway and enable breathing to be assisted if necessary. Once a tube has been positioned it will also protect the airway when the swallowing and coughing reflexes are suppressed, so that material such as saliva does not enter the trachea.
Endotracheal tubes for larger species (>1-2kg) are available from many manufacturers and are provided either as plain tubes or with an inflatable cuff that seals the gap between the wall of the tube and the trachea. The cuff can be inflated either with a syringe (2–5 ml) or with a specially designed inflator. The cuff is prevented from deflating either by means of a non-return valve (present on most disposable tubes) or by clamping with a pair of haemostats. Tubes may be reusable or be intended only for single use. Reusable tubes are generally constructed of rubber and are opaque. They deteriorate gradually, becoming brittle and easily kinked. The cuff often becomes distorted and may leak, so it is preferable to purchase single-use tubes and allow a limited amount of reuse. Clear
Endotracheal tubes suitable for use in small rodents and other small laboratory species can be purchased commercially, but can also be constructed from “over-the-needle” intravenous catheters.
Endotracheal intubation of large animals such as dogs, sheep, pigs, old-world primates and large birds (>1 kg) is relatively straightforward, provided a suitable size and shape of the laryngoscope is available. Laryngoscopes are used to obtain a clear view of the larynx so that an endotracheal tube may be passed easily and
|Species||Body weight||Endotracheal tube diameter||Laryngoscope|
|Cat||0.5–1.5 kg||2.0–3 mm O/D*||MacIntosh size 1|
|. 1.5 kg||3–4.5 mm O/D|
|Dog||0.5–5 kg||2–5 mm O/D||MacIntosh size 1–4|
|. 5 kg||4.0–15 mm O/D|
|Guinea pig||400–1000 g||16–12 gauge plastic cannula||Purpose-made laryngoscope † Otoscope|
|Hamster||120 g||1.5 mm||Purpose-made laryngoscope †|
|Mouse||25–35 g||1.0 mm||Purpose-made laryngoscope †|
|Primate||0.35–20 kg||2–8 mm O/D (or purpose-made tube for smallest animals)||MacIntosh or Wisconsin size 1–3|
|Pig||1–10 kg||2–6 mm O/D||Soper or Wisconsin size 1–4|
|10–200 kg||6–15 mm O/D|
|Rabbit||1–3 kg||2–3 mm O/D||Wisconsin size 0–1 or Otoscope|
|3–7 kg||3–6 mm O/D|
|Rat||20||18–12 gauge plastic cannula||Purpose-made laryngoscope † Otoscope|
|Sheep||10–90 kg||5–15 mm O/D||MacIntosh size 2–4|
|* O/D, Outside diameter|
|† see text|
The animal should be anaesthetised to a sufficient depth to abolish the cough and swallowing reflexes. Before intubating any animal, oxygen should be administered for approximately 2 minutes. If the larynx is inadvertently obstructed during attempted intubation, it will usually take over 60 seconds for hypoxia to develop if the animal has been breathing oxygen. If the animal has been breathing air, hypoxia will develop much more rapidly.
Intubation of Dogs, Cats and Sheep
The animal is placed in sternal recumbency, with its jaws opened as widely as possible by an assistant. The tongue is drawn forwards and the laryngoscope advanced over the tongue towards the pharynx. The larynx is usually masked by the epiglottis. Gentle upward pressure on the soft palate with the end of the endotracheal tube will disengage the epiglottis, allowing it to fall forwards, providing an unobstructed view of the larynx. In cats and sheep the larynx should be sprayed with a local anaesthetic, to prevent laryngospasm. Disposable ‘insulin’ syringes with a pre-attached 25-SWG (Standard Wire Gauge) needle, with the needle bevel cut off, are ideal. The endotracheal tube can then be advanced through the larynx into the trachea. Then the tube should be connected to the anaesthetic breathing system, the cuff (if present) inflated and the tube tied in place to the animal’s jaw, using a 1-cm-wide cotton tape. It is preferable at this stage to assist ventilation (as described earlier) and observe that there is movement of both sides of the thorax. This ensures that the tube has not been inadvertently positioned in one of the two mainstem bronchi. In addition, manual inflation of the chest will enable an appreciation of the degree of resistance to gas flow. Increased resistance may indicate twisting or kinking of the tube, or its partial obstruction due to positioning close to the bifurcation of the trachea. If any uncertainty exists about tube placement, use a stethoscope to check that breath sounds can be heard on both sides of the thorax.
Intubation of Pigs
Intubation in the pig is complicated by the difficulty of obtaining an unobstructed view of the larynx. The animal is best positioned on its back or chest, with the
The laryngoscope is advanced over the tongue and the epiglottis disengaged from the soft palate, if necessary, by pushing on the soft palate using the tip of the introducer. Once the larynx has been located, it should be sprayed with lidocaine. The introducer and the endotracheal tube can then be gently advanced into the larynx and the introducer withdrawn. The tube should then be gently advanced; at this stage, its progress is usually arrested by the laryngeal wall. If this occurs, the tube should be withdrawn very slightly, rotated through 90 degrees and reinserted. This should be repeated as necessary until no resistance is experienced. Under no circumstances should attempts be made to pass the tube forcibly through the larynx, as this is likely to result in severe trauma, oedema, haemorrhage and consequent asphyxiation.
Intubation of Rabbits
Visualisation of the larynx in the rabbit is difficult, and it is necessary to use a purpose-designed laryngoscope blade (Flecknell blade), a Wisconsin laryngoscope blade (2- to 5-kg animal, size 1; 1- to 2-kg animal, size 0) or an otoscope if intubation is to be carried out under direct vision.
Intubation Using an Otoscope or Laryngoscope
The rabbit is positioned on its back. To view the larynx, the tongue is gently grasped and pulled forwards and to one side, taking care to avoid the sharp edges of the incisor teeth. The otoscope or laryngoscope is introduced into the mouth and advanced until the larynx is visible. It is possible to advance the instrument into the oesophagus if the tip of the epiglottis is positioned on the nasal aspect of the soft palate. To avoid this, the soft palate can be pushed with the otoscope or laryngoscope tip, or the introducer can be passed down the otoscope and pushed against the soft palate to reposition it and provide a clear view of the larynx.
As the speculum is advanced, the paler triangle of the epiglottis can often be seen through the end of the soft palate, alerting the anaesthetist to the need to manipulate the structure. In many cases, the larynx is immediately clearly visible. At this point, the larynx can be sprayed with lidocaine, although this is often unnecessary. An introducer can now be passed through the otoscope into the larynx and on into the trachea. If a purpose-made introducer is not available, then a bitch or cat urinary catheter can be used, depending upon the size of the rabbit. If a catheter is used, then the Luer fitting should be removed before use, since this will not pass through the tip of the otoscope. After placing the introducer, the otoscope or laryngoscope is removed, taking care not to change the position of the introducer. An endotracheal tube (2.5–3 mm for 2- to 3-kg rabbits) is then threaded onto the end of the introducer and advanced into the trachea. When the endotracheal tube reaches the larynx, some resistance is often felt. Gently rotating the tube as it is advanced may ease its passage into the trachea. Prior application of lubricating gel (e.g. lidocaine gel) can also aid the passage of the tube. Take care at this stage not to remove the introducer until the tube is in the trachea, or intubation will be unsuccessful. As the tube is advanced further into the trachea, the introducer is removed and the tube tied in place.
An alternative technique for intubation does not require visualization of the larynx. The rabbit is placed in sternal recumbency, and the head gripped firmly and extended and the animal lifted so that its forelegs are just touching the operating table. The endotracheal tube is advanced through the gap between the incisors and the premolars, over the tongue and towards the larynx. The operator listens for breath sounds at the end of the tube or alternatively, if a clear polyethylene tube is used, looks for the presence of condensation. A loud breath sound or condensation indicates that the tube tip is close to the larynx. As the rabbit breathes in, the tube is gently advanced. If it fails to enter the larynx, as indicated by cessation of breath sounds and loss of condensation, then the tube is withdrawn, the head repositioned either by tilting it further backwards or slightly forwards and another attempt made. Giving a quarter turn to the endotracheal tube as it enters the larynx can help its passage.
In some instances, intubation can be eased by the use of a local anaesthetic spray. This can be delivered onto the larynx by positioning the endotracheal tube at the point of maximal breath sounds, and then spraying lidocaine into the end of the tube, or injecting a small (0.1 ml) quantity of lidocaine into the end of the tube. The local anaesthetic is drawn down the tube as the rabbit inhales, and some reaches the larynx. After waiting a minute or two to allow the drug to act, another attempt at intubation can be made. If problems arise, oxygen should be administered every 2–3 minutes to ensure the animal does not become hypoxic.
Although this technique sounds challenging, it is relatively easy to become proficient and has the advantage of requiring no additional equipment. In small rabbits (<1 kg), it is not always possible to hear breath sounds or observe condensation in the small endotracheal tube (2–2.5 mm) that is needed. For this reason, it is best to intubate larger rabbits when first attempting this technique.
With both techniques, the confirmation of successful placement is based on observing condensation of breath on a cold surface (e.g. the end of the otoscope handle), or movement of a piece of tissue paper placed at the end of the tube. Alternatively, as in other species, a capnograph can be attached to confirm the tube is in the trachea.
Intubation of Rats
Intubation of the rat is possible using a number of different purpose-made intubation devices or using an otoscope. The rat is positioned on its back, and the tongue pulled gently forward and to one side. The laryngoscope or otoscope is then inserted until the larynx can be visualized. The animal can then be intubated using a suitably sized (12- to 16-gauge) arterial cannula (e.g. Abbocath, Abbott Laboratories). Some modification of the Luer fitting is needed to provide connections to an appropriate anaesthetic breathing system, and care must be taken to ensure that these connectors introduce only a minimum of dead space into the breathing system. To avoid inadvertent intubation of one bronchus, and to provide a seal around the larynx, a small piece of rubber tubing can be positioned around the catheter, about 0.75–1 cm from the tip. Alternatively, some ‘Micropore’ tape (3M) can be applied to make a similar cuff. This will reduce the leakage of gas around the tube, making ventilation more effective, and will also improve the efficacy of positive
When using an otoscope, it is necessary to use an introducer, since the cannula will not pass through the lumen of the otoscope. A guide wire from a Seldinger catheter makes an ideal introducer since its tip is soft and flexible. The wire is passed through the otoscope and through the larynx under direct vision, the otoscope carefully removed and the endotracheal tube threaded over the wire into the trachea. These wires can be purchased separately, and a 0.7-mm-diameter wire will fit through both 16- and 18-gauge catheters. Alternatively, the neck may be transilluminated using a powerful light source and the mouth opened using a small gag. The tongue is pulled forwards and a bright spot of light seen, which flashes as the rat breathes; this indicates the opening of the larnyx.
One of the simplest techniques to master is to purchase one of the commercially available systems that usually combine a small table for positioning the animal with a system for visualizing the larynx. The apparatus can be used for intubation of both rats and mice. An alternative approach using a fibre optic system has also been described and a commercially produced instrument is also available.
Intubation of Guinea Pigs, Mice, Gerbils and Hamsters
Intubation of the mouse, gerbil and hamster is more difficult than in larger species and requires especial skill and purpose-made apparatus. A suitable set of laryngoscope blades has been described and can be used in mice and other small rodents. The guinea pig can also be intubated using a purpose-designed laryngoscope blade, or the technique employing an otoscope, as described above for the rat, can be used. As with the rat, the use of an otoscope in combination with transillumination of the neck provides optimal conditions for intubation. Positioning of the otoscope is more difficult in the guinea pig than in the rat, and a narrow speculum is needed to pass between the cheek teeth. The pharynx narrows markedly at the junction with the larynx and the oesophagus, and considerable care must be taken to avoid inserting the speculum too far and occluding the larynx. As with the rat, intubation is achieved by passing a Seldinger guide wire through the larynx, removing the otoscope and then passing a 12- to 16-gauge catheter over the wire into the trachea.
Intubation of Birds
Intubation of birds is relatively
Placement of a catheter in one nostril allows delivery of oxygen or anaesthetic gases, and can be a useful alternative to a face mask, for example, when using a stereotaxic frame. A variety of catheters can be used, including vascular catheters, nasogastric feeding tubes, flexible oral-dosing catheters and urinary bladder catheters. The catheter should be lubricated before insertion and, in most species, should be directed medially and ventrally. The nostrils of most small animals are surrounded by muscle, and this restricts the diameter of the nasal opening, but gentle pressure from the catheter tip will usually dilate the nostril slightly, allowing passage of the catheter. Slight rotation of the catheter can aid insertion. The diameter of the passage through the nasal chamber is often significantly larger than the external nasal opening. Occasionally, there may be slight trauma to the nasal mucosa, resulting in a small amount of haemorrhage, but this is usually minor and stops rapidly.
The fresh gas flows needed are similar to those when using a face mask, approximately three times the animals’ minute volume. Some air will be drawn in through the other nostril, and this will dilute the supplied gas. However, increasing the vaporizer setting (e.g. by 0.5–1% when using isoflurane) will compensate for this. Gas scavenging can be accomplished using a down-draft table. The catheter can be connected to the anaesthetic machine using a Luer adapter and oxygen bubble tubing. This tubing is extremely useful for adapting anaesthetic breathing systems, as its internal diameter varies along its length from 3 to 8 mm, allowing it to be cut at a convenient point to connect different-sized connectors.