INTRODUCTION

TOP ABSTRACT INTRODUCTION METHODS AND RESULTS DISCUSSION REFERENCES

THE ABILITY TO SUCCESSFULLY INTUBATE the trachea of mice and control their ventilation is important for longitudinal studies requiring recovery from anesthesia and repeated pulmonary-function measurements or other physiological or anatomic measurements such as the use of radiological imaging (e.g., computed tomography or magnetic resonance imaging). There have been some reports in the literature regarding the intubation and ventilation of mice, but most have not provided sufficient detail that would enable one to duplicate this procedure (2, 3, 7).

In this communication, we present a simple method of intubation and ventilation in mice by using a smoothed metal blade as a laryngoscope to facilitate oropharyngeal exposure, transillumination of the neck to facilitate visualization of the trachea through the oropharynx, readily available polyethylene (PE) tubing to intubate the trachea, and a simple solenoid ventilator to maintain physiological ventilation. This method proved useful for repeated pulmonary-function measurements and agonist challenge in individual mice.

	METHODS AND RESULTS

TOP ABSTRACT INTRODUCTION METHODS AND RESULTS DISCUSSION REFERENCES

For our studies, we used three species of common inbred mice (A/J, C3H, and C57B/6), at 6-10 wk of age (Jackson Laboratories, Bar Harbor, ME). Anesthesia was induced with etomidate (2 mg/ml), an ultra-short-acting nonbarbiturate hypnotic agent, and fentanyl (50 µg/ml), a short-acting narcotic agent in a 3:1 mixture (0.2-0.3 ml ip). This dose can keep 20- to 25-g mice of these strains fully anesthetized for 20-30 min. The mouse was then suspended at an ~45° angle by the two front upper teeth by a rubber band attached to a Plexiglas support (Fig. 1). A 150-W halogen light source (World Precision Instruments, Sarasota, FL) with two 24-in. flexible fiber-optic arms allowed transillumination of the trachea just below the vocal cords (Fig. 1). The positions of the fiber-optic arms were adjusted in each mouse to provide the best visualization of the trachea. A metal laryngoscope fabricated from 3/32-in. sheet brass (Fig. 2) was used to lift the lower jaw of the mouse and keep the mouth open and the tongue displaced to maximize oropharyngeal exposure. This provided a clear view of the tracheal opening (Fig. 3). During this direct visualization, a 2-cm-long PE-90 catheter attached to the hub of a needle (Fig. 2) was inserted ~3 mm into the trachea. This PE tubing had a tip beveled at ~45°, with the bevel on the convex side of the natural curve of the tubing. To prevent tissue damage from the bevel point, the sharp tip end was carefully rounded off with small dissecting scissors. The mouse was then removed from the Plexiglas support and attached to the mouse ventilator. The mouse was ventilated at 120 breaths/min with a tidal volume of 200 µl. These values have been previously shown to provide adequate ventilation in mice of these sizes and strains (6).

View larger version (126K): [in this window] [in a new window]   Fig. 1.   Illustration of a mouse suspended on plastic support for visualization of trachea through the oropharynx. Fiber-optic light source is adjusted just below vocal cords to provide the best view of trachea.

View larger version (148K): [in this window] [in a new window]   Fig. 2.   Illustration of custom-made mouse "laryngoscope," and the PE-90 catheter attached to the needle hub used for endotracheal intubation.

View larger version (91K): [in this window] [in a new window]   Fig. 3.   Close-up photograph of typical view of the illuminated trachea seen through oropharynx, before placement of PE-90 cannula. Image identifies tracheal structures.

We assessed potential air leaks around the endotracheal tube by inflating the lung to 20 cmH₂O and measuring the change in pressure after the intubated lungs were sealed off. Figure 4 shows negligible air leak around the endotracheal tube at this pressure in one mouse. Thus the elastic properties of the trachea around this size PE tubing are sufficient to provide a reasonably tight functional seal. Similar observations were also made in mice from other strains and at ages up to 16 wk.

View larger version (17K): [in this window] [in a new window]   Fig. 4.   Airway pressure tracing from 1 mouse, demonstrating negligible air leak around endotracheal tube even at an air pressure of 24 cmH2O.

In some animals, we also tested the airway responsivity to intravenous acetylcholine (ACh). With the solenoid ventilator, the respiratory resistance and compliance can be quickly measured with a sudden inspiratory occlusion, as previously described in detail (4). However, the agonist delivery approach used by Ewart et al. (4) of direct injection into the abdominal inferior vena cava does not allow the animal to recover. To administer bronchoconstrictor agents in the present study, we made a small incision in the neck to expose a jugular vein. Then a 31-gauge needle was inserted into the jugular vein. This needle was connected to a syringe containing ACh (0.67 µg/ml) with PE-10 tubing. The response to a 1-min ACh infusion at stepwise increasing rates is shown in Fig. 5. Figure 5 also shows the respiratory resistance and compliance at each of these infusion rates. Over this dose range shown, there was a progressive 23% fall in compliance and a 270% increase in resistance. Note that to calculate the respiratory system resistance, it is necessary to subtract the resistance of the intubation cannula, which, in our system, equals 1.7 cmH₂O · ml¹ · s¹. After the final concentration was delivered, the needle was withdrawn, and the skin wound was closed with cyanoacrylate adhesive (Future Glue, Pacer Technology, Rancho Cucamonga, CA).

View larger version (12K): [in this window] [in a new window]   Fig. 5.   Ventilatory pressure and flow records from 1 mouse, showing response to increasing concentrations of acetylcholine (ACh) delivered to jugular vein. Shown are individual breaths with an inspiratory occlusion of 0.24 s used to calculate the resistive and compliant pressures (Pr and Pc, respectively). Respiratory system resistance (Rrs; units = cmH2O · ml1 · s1) and compliance (Crs; units = ml/cmH2O) are then calculated from the flow and tidal volume as previously described (4). ACh infusion rates are in µg/min.

Mice were then removed from the ventilator and monitored until they started to breathe spontaneously. This usually occurs within 1 min after removal. The endotracheal tube was then removed, and the mice were returned to their cages when they regained their righting reflex. All animals recovered, and normal mouse behavior was evident within 2 h.

	DISCUSSION

TOP ABSTRACT INTRODUCTION METHODS AND RESULTS DISCUSSION REFERENCES

Until recently, most studies in mice that required measurement of pulmonary function have been limited to one time point. Once the animal was anesthetized, the trachea was secured by incising the neck and performing a tracheostomy, with no allowance for recovery and repeated measurements. With the current interest in the development of genetic models using mice, the ability to perform longitudinal studies has many obvious advantages. It is to this end that we have described in detail the methods that allow repeated intubation, ventilation, and agonist challenge in mice. This method can be used for repeated pulmonary-function measurements over time. Furthermore, this method can also be used for other studies in mice, where surgery and recovery are required, or other protocols, which may include radiological imaging where respiratory control is necessary.

Although many aspects of the method described here are not new and have been used by different investigators, there has not been a comprehensive description that would allow others to duplicate the procedure. In their studies of cardiac electrophysiology in the C57BL/6J mouse, Berul et al. (2) described intubation of C57BL/6J mice via direct laryngoscopy, with transillumination of the ventral neck. These authors used a rodent respirator to successfully evaluate responses to pacing and other cardiovascular alterations. Unfortunately, their paper only provides cursory details on how the mice were intubated. In an abstract, Deyo and Wei (3) also mention a method of intubation in mice by using an endotracheal tube made from an over-the-needle catheter. However, there was sufficient trauma to the trachea, such that 3 of the 20 mice in their study group died due to complications from the technique. Neither of these two studies was concerned with repeated assessments of respiratory function and reactivity. One relatively new approach has been used for repeated assessments of airway reactivity in mice, which uses barometric plethysmography (5). Unfortunately, the complex variable derived for this purpose (Penh) does not have any clear relation to conventional resistance and compliance measurements and may be dominated by inspiratory and expiratory timing. Indeed, very recent work (1) demonstrated that airway responses to methacholine could be assessed, with less noise and greater reproducibility, simply by measuring the changes in ventilatory timing. The method we describe in the present paper does allow repeated measurements of conventional indexes of pulmonary function. We have not investigated the maximal number of repeated measurements that can be obtained, nor the timing between them. However, we have successfully been able to reintubate the mice and reinject the same jugular vein with ACh 1 wk later. We did observe minor wound adhesions and dried adhesive in the neck that needed to be dissected away to expose the jugular vein, but the repeat needle insertion and injection were without problem, and this second closure healed without complication.

In summary, we have described a method for rapid and repeated intubation of mice for baseline pulmonary-function measurements, measurement of conventional respiratory responses to agonist challenge, or for other studies where controlled ventilation is required for repeated experimental procedures.