A Simple Method of Mouse Lung Intubation

Summary

This paper describes a striaghforward and efficient method of intubating mice for pulmonary function measurements or pulmonary instillation, that allows the mice to recover and be studied at later times. The procedure involves an inexpensive fiberoptic light source that directly illuminates the trachea.

Abstract

A simple procedure to intubate mice for pulmonary function measurements would have several advantages in longitudinal studies with limited numbers or expensive animal. One of the reasons that this is not done more routinely is that it is relatively difficult, despite there being several published studies that describe ways to achieve it. In this paper we demonstrate a procedure that eliminates one of the major hurdles associated with this intubation, that of visualizing the trachea during the entire time of intubation. The approach uses a 0.5 mm fiberoptic light source that serves as an introducer to direct the intubation cannula into the mouse trachea. We show that it is possible to use this procedure to measure lung mechanics in individual mice over a time course of at least several weeks. The technique can be set up with relatively little expense and expertise, and it can be routinely accomplished with relatively little training. This should make it possible for any laboratory to routinely carry out this intubation, thereby allowing longitudinal studies in individual mice, thereby minimizing the number of mice needed and increasing the statistical power by using each mouse as its own control.

Introduction

In 1999, Brown et al. published a paper describing a method for intubation of the mouse lung ¹. Such a technique has considerable utility in doing repeat pulmonary function or bronchoalveolar lavage in individual mice in longitudinal studies ². Since that original paper, there have been several other papers that have described different approaches to mouse intubation ^3-9. While all of these methods can be used successfully, they usually require considerable training or cost. One of the main issues with such intubation is that as the intubation cannula approaches nears the trachea pending insertion, the cannula itself blocks the light and hence the visualization of where it needs to go. Thus, the insertion becomes blind at the most critical time. In this paper we show how to simply and inexpensively eliminate this visualization problem, thereby ensuring successful intubation with relatively little training or experience.

Protocol

1. Preparing for the Procedure

One must first obtain and prepare the following items:

1. The cannula. For intubation of 20-35 g mice, we use a 1 or 1.5 inch long, 20 gauge IV catheter (BD Insylte, Sparks, MD or Jelco Optiva, Carlsbad, CA). A new sterile catheter can be used for each mouse, but catheters can also be reused after sterilization by soaking in 70% ethanol overnight. Although neither the pharynx nor trachea of the mouse is sterile proper cleanliness procedures, including the use of sterile gloves and instruments, should be followed.
2. The fiberoptic cable. We use ≈70 cm of 0.5 mm optical cable from Edmund Optics, but the length is not critical. It is important to make sure the fiber has its edge smoothed, since after cutting the cable to length with a razor, the edge is left relatively sharp, and it does not take much effort to pierce the tracheal wall. However, it is very easy to smooth this edge by holding the fiber about 2 cm from the end and then making small circles for a few seconds with the edges of the tip touching a piece of 1,000 grit emery paper (see demonstration in the video and Figure 1 of Macdonald, et al. ¹⁰). The other end is inserted through a rubber stopper. This is most easily accomplished by first pushing an 18-gauge needle through the stopper, inserting the optical fiber through the needle bore, then withdrawing the needle. The rubber stopper is connected to a 150 watt halogen light source (e.g. NCL-150, Volpi USA, or any other or light source, even less than 150 watts). It is important to make sure to use a stopper made of silicone rubber (or other heat resistant material), since ordinary rubber or cork may burn when located so close to the hot light source.

2. Performing the Intubation

1. See Figures 1 and 2. Insert fiber optic cable through a short piece of silicone rubber tubing (≈0.8 mm ID x 4 mm OD, Cole-Palmer, EW-96410-13). Tie this rubber tube fairly tight, while still allowing the fiber optic cable to be adjusted. Inserting the silicone tubing snugly in the Luer end of the cannula fixes the fiber optic cable's position inside the cannula. Adjust the position of the fiberoptic cable so that it extends through the cannula ≈4 mm in front of the cannula tip.
2. Place the anesthetized mouse on a vertical support, suspended by its upper incisors (Figure 3). Most investigators find the best visualization with the ventral side of the mouse facing themselves. Very gently pull out the tongue and hold with thumb and forefinger. The middle finger is placed between the neck and plastic support. Traction on the tongue with the index finger and thumb is used to open the mouth, and to straighten the intubation path, the angle of the head is adjusted with the middle finger behind the neck shown in Figure 3.
3. Using the fiber optic cable as a light source and introducer, push it through the visualized vocal cords. If the cords are not visible, gently pull harder on the tongue using the middle finger as support. When inserted, advance the cannula ≈5 mm further. Then, being very careful not to move the cannula, withdraw the fiber optic cable. Lie the mouse down and secure the cannula with a piece of tape and support the cannula hub on a piece of Plasticine (modeling clay), as shown in Figure 4.
4. The procedure in step 3 cannot be easily taught or even demonstrated, since it is a solo operation. However, by subtle adjustments of the traction on the tongue and the support behind the head nearly all who try this soon find the right way to position the mouse to visualize the vocal cords.

Results

As an evaluation of the method, we used four 20 week old male BALB/c mice with average weight (± SEM) of 27.7 ± 0.40 g. They were studied on five consecutive weeks, where the lung resistance was measured using a system as previously described ¹¹. Each mouse was anesthetized with ketamine (100 μg/g BW) and xylazine (15 μg /g BW) in saline via IP injection. They were then intubated as described above. If there is any doubt whether the cannula is in the trachea and not the esophagus, this can be validated using a...

Discussion

The procedure described here has several advantages. First the apparatus is simple and relatively inexpensive.. The fabrication of the apparatus does not require any special tools or costly equipment. The use of a catheter introduces that also is the light source means that one never loses sight of the tracheal opening as the introducer approaches the tracheal opening. The use of a 0.5 mm introducer also serves to minimize trauma that might occur with an initial insertion of a larger cannula. We note here that a similar ...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Intubation cannula	BD Insylte, Sparks, MD or Jelco Optiva, Carlsbad, CA		1-in.-long, 20-gauge intravenous (IV) catheter
Fiber-optic cable	Edmund Optics, Barrington, NJ	#NT02-542	Approximately 2-ft length of 0.5-mm optical fiber (Communication grade plastic fiber). The edge of the fiber end that is inserted into the trachea should be gently rounded by holding the fiber ≈2 cm from the end and then making small circles while dragging the tip for a few seconds on 1,000-grit emory paper.
Light Source	Volpi	NCL-150	Although we use a 150-W halogen light source, any equivalent light source, even with much lower wattage can be used.
Aluminum tube			One inch O.D., with 1/16 inch wall. This may need to be change to fit whichever light source is used.
Rubber stopper			A #4 rubber stopper fits the 1 inch tube.
Small silicone rubber tube	Cole-Palmer	EW-96410-13	A ≈1.5 cm piece of silicone rubber tubing (0.8 inner diameter, 4 mm outer diameter)
Angled support stand			Ours is constucted from plexiglass, but any material to which a thread or wire can be affixed to hold the mouse at an almost vertical angle can be use.