Endotracheal Intubation in Mice via Direct Laryngoscopy Using an Otoscope

Summary

We have developed a simple, reliable, and relatively inexpensive method for endotracheal intubation in mice via direct laryngoscopy using an otoscope with a 2.0 mm speculum. This technique is atraumatic and can be used for repeated measurements in chronic experiments. We find it superior to tracheostomy or previously reported nonsurgical techniques.

Abstract

Mice, both wildtype and transgenic, are the principal mammalian model in biomedical research currently. Intubation and mechanical ventilation are necessary for whole animal experiments that require surgery under deep anesthesia or measurements of lung function. Tracheostomy has been the standard for intubating the airway in these mice to allow mechanical ventilation. Orotracheal intubation has been reported but has not been successfully used in many studies because of the substantial technical difficulty or a requirement for highly specialized and expensive equipment. Here we report a technique of direct laryngoscopy using an otoscope fitted with a 2.0 mm speculum and using a 20 G intravenous catheter as an endotracheal tube. We have used this technique extensively and reliably to intubate and conduct accurate assessments of lung function in mice. This technique has proven safe, with essentially no animal loss in experienced hands. Moreover, this technique can be used for repeated studies of mice in chronic models.

Introduction

The laboratory mouse has supplanted virtually all species as the principal mammalian model of biology and pathobiology. The laboratory mouse is the smallest mammalian species that has been clearly and extensively shown to be of value as a model of human disease and has proven invaluable in advances of our understanding of human biology and disease. The short gestation time and substantially lower cost has allowed the development and study of null and transgenic mice as a commonplace tool in biomedical research. However, the size of the average laboratory mouse (20-25 g) has limited their study in physiologically or surgically based studies and, consequently, some investigators study larger mammalian species. An impediment to using mice in these studies is the difficulty encountered with intubation techniques that would allow physiological measurements or extensive surgical procedures under deep anesthesia. Tracheostomy ¹ has been used as a standard technique instead of intubation because of the greater ease of performing this technique and modest skill required. However, tracheostomy is not conducive to chronic or recovery surgery studies; thus, it is limited to acute experiments. Tracheostomy can also be a confounding variable in research in which inflammation or sensitive physiologic reflexes are important.

Our laboratory has tried most of the techniques described by other investigators and found them inadequate for a variety of reasons. Tracheostomy is too traumatic and induces bleeding and airway inflammation. Much more problematic is that it cannot be feasibly repeated. Many relatively noninvasive techniques that require a modest investment in equipment are not sufficiently reliable. Other techniques require expensive equipment that is difficult to justify without knowing if the equipment will work in a specific application. Thus, we sought to develop a nontraumatic technique that required no more than a modest investment in specialized equipment, could be accomplished quickly and reliably, could be repeated in chronic models, and could be used in large numbers of animals. Here we report such a technique.

Protocol

1. Animal Preparation

1. Obtain mice that are older than 8 weeks and more than 20 g (smaller mice can be intubated by an expert).
2. Anesthesia
   1. Inject mice with  20 mg/kg, each, of ketamine and xylazine intraperitoneally as a preanesthetic. (This dose is insufficient to fully anesthetize the mouse, but facilitates the safe transfer, after intubation, to mechanical ventilation.) However, adjustment of the dosages may be necessary according to the anesthetic response in consultation with the institutional veterinarian.)
   2. Induce anesthesia in mice with 3.5% isoflurane/oxygen in an induction chamber for 90-120 sec.
   3. Observe the respiratory rate carefully. It should slow progressively to not less than 45 bpm.

2. Preparation for Intubation

1. Remove the mouse from the induction chamber and scruff tightly.
2. Use cushioned fine forceps to gently extend the tongue from the mouth.
3. Maintain extension of the tongue by holding the tongue between two fingers and applying gentle force.
4. Pull the mouse up on the speculum of the otoscope with a vertical motion.
5. Look through the ocular of the otoscope while scruffing tightly and pull the mouse up gently by the tongue and the scruff.
6. Look carefully for the vocal cords. They should be easily visible at this time. The laryngis aditus (the structure of the laryngeal opening) should look relatively white. There should be movement of the cords with each breath. If the cords are not moving or are poorly visualized, rotate the animal slightly and gently hyperextend the neck.
7. With the dominant hand, hold the 20 G, 1 in catheter, with a 1 cm length of the PE10 tubing extending through the tip of the catheter, like a pencil and insert it into the side of the speculum. The PE tubing serves as a stylet or bougie.
8. Direct the PE10 tubing (stylet) through the vocal cords and advance the 20 G catheter (endotracheal tube) over the PE tubing until the hub is at the level of the lower incisor. Remove this stylet quickly.
9. Gently take the animal off of the speculum and verify the tube location by placing the animal on a mechanical ventilator with continuous 2% isoflurane to maintain general anesthesia throughout the experiment. Visualize expired air (bubbles) going through a PEEP trap to confirm intubation. Although esophageal intubation may result in a few bubbles, it will not be as pronounced or as consistent with endotracheal intubation. Also, observe an airway pressure tracing for negative deflections (Figure 1). These confirm proper placement of the tube. Alternatively, place a small amount of water into IV tubing and connect it to the ET tube. Verifying movement to and from the mouse with respiration². In the case of esophageal intubation, the pressure tracing will reveal significantly higher pressures and no negative deflections. Although end-tidal CO₂ would be helpful, technically this would be very difficult given the small tidal volumes (~200 µl) and the equipment to do this would be very expensive. The simple techniques described are fully sufficient and much less expensive. If isoflurane anesthesia is not available, all procedures can be conducted using ketamine 80-120 mg/kg in conjunction with xylazine 10-20 mg/kg for general anesthesia.   

3. Alternative Technique: Direct Laryngoscopy with the Otoscope Technique is Used Readily for Other Purposes, Mainly for Direct Instillation of Study Substances into the Lung.

1. Induce anesthesia in the animals with isoflurane alone for this procedure at 3.5% for 90-120 sec in the induction chamber.
2. Scruff the animal tightly at the base of the skull and extend the tongue gently with fine forceps.
3. Holding the tongue gently, pull the animal straight up on the speculum (unmodified) until the animal cannot be pulled up further.
4. The vocal cords can be visualized most often by this technique alone, but rotate the animal and hyper-extended the neck to bring them into full view.
5. Advance a gel-loading pipette containing the instillate to the glottis and instill the fluid. Because the speculum makes a seal with the hypopharynx the mouse will aspirate any residual liquid that does not go through the cords. This is easily verified by the rhonchorous sounds the mouse now makes when breathing until the fluid is distributed fully in the lungs.
6. Alternatively, connect PE10 tubing to a 0.5 ml syringe containing 50 µl of fluid with an air bolus behind the liquid. Advance the PE tubing carefully through the vocal cords for 0.5 cm and discharge the contents into the airway slowly. However, occasional trauma to the lower airway occurs using this technique without better delivery of the sample.

Results

Intubation with the above technique is reliable and fast. The appropriate placement of the endotracheal tube is most easily verified by observing expired gas bubbling from the submerged expiratory limb of the of the ventilator circuit (usually in a PEEP trap) and negative deflections on an airway pressure tracing (Figure 1). The negative deflections on airway pressure tracing are the most reliable. Others have used movement of a small fluid droplet in intravenous tubing connected to the expiratory ventil...

Discussion

In this report we describe a simple, reliable technique to intubate mice that is nontraumatic and can be used repeatedly in the same animal. This technique can be accomplished with simple laboratory or medical equipment that can be purchased for a modest sum. The technique of direct laryngoscopy, originally reported by Hastings and colleagues⁴, can also be used for a variety of purposes, but mainly to accurately deliver test substances to the lower respiratory tract. We have found this technique superior to th...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Operating Otoscope Head	Welch Allyn	21700	$188.98
Otoscope Handle	Welch Allyn	71000	$112.20
Reuseable Speculum	Welch Allyn	22002	$3.98
Fine Forceps	Miltex	18-779	$107.18
Small clamp stand to hold otoscope
Isoflurane Vaporizer (multiple circuit with vacuum waste gas handling)	Summit Medical		$3,000
Flexivent (Animal Ventilator)	SCIREQ		$35,000
Intravenous catheter (20 G, 1 in)	BD	381233	$9.62
Polyethylene tubing (PE10) (0.011 in I.D., 0.024 in O.D.) 100 ft	Intramedic, Clay-Adams	427401	$115.70
Ketamine 100 mg/ml (10 ml bottle)	Butler	23061	$10.00
Xylazine (100 ml bottle)	Vedco	24105	$20.00
Isoflurane (250 ml bottle)			$15.00
Vecuronium bromide 10 mg/10 ml	Pfizer	NDC 0069-0094-01	$15.00