Instillation and Fixation Methods Useful in Mouse Lung Cancer Research

Summary

The goal of this paper is to describe simple methods that will greatly aid in the setup and analysis of mouse lungs with lung cancer or other pathologies. We present 3 protocols to simply and reliably carry out lung instillations, fixation, and lung volume measurements.

Abstract

The ability to instill live agents, cells, or chemicals directly into the lung without injuring or killing the mice is an important tool in lung cancer research. Although there are a number of methods that have been published showing how to intubate mice for pulmonary function measurements, none are without potential problems for rapid tracheal instillation in large cohorts of mice. In the present paper, a simple and quick method is described that enables an investigator to carry out such instillations in an efficient manner. The method does not require any special tools or lighting and can be learned with very little practice. It involves anesthetizing a mouse, making a small incision in the neck to visualize the trachea, and then inserting an intravenous catheter directly. The small incision is quickly closed with tissue adhesive, and the mice are allowed to recover. A skilled student or technician can do instillations at an average rate of 2 min/mouse. Once the cancer is established, there is frequently a need for quantitative histologic analysis of the lungs. Traditionally pathologists usually do not bother to standardize lung inflation during fixation, and analyses are often based on a scoring system that can be quite subjective. While this may sometime be sufficiently adequate for gross estimates of the size of a lung tumor, any proper stereological quantification of lung structure or cells requires a reproducible fixation procedure and subsequent lung volume measurement. Here we describe simple reliable procedures for both fixing the lungs under pressure and then accurately measuring the fixed lung volume. The only requirement is a laboratory balance that is accurate over a range of 1 mg–300 g. The procedures presented here thus could greatly improve the ability to create, treat, and analyze lung cancers in mice.

Introduction

For a number of reasons, lung cancer has not been widely studied in the mouse. One reason for this is that access to the lung is very difficult in vivo, and quantitative analysis of fixed lungs is not commonly done. The methods described in this paper are designed to remedy this situation. The goals herein are to describe simple methods that will greatly aid in the setup and analysis of mouse lungs with lung cancer or other pathologies. While none of these approaches are entirely new, they have not been presented together as stand-alone methods in the simplified manner as described here.

There have been a number of manuscripts that have described methods for intubation of the mouse lung primarily for the purpose of doing repeat pulmonary function or bronchoalveolar lavage in individual mice in longitudinal studies. Since that original paper, there have been several other papers that have described different approaches to mouse intubation^1-9. While all of these methods can be used successfully, they usually require considerable training, and are often not without a nontrivial failure rate. In addition, in order to carry out pulmonary function measurements, the cannula needs to fit the trachea tightly enough so that there is no air leakage. However, another practical use for intubation is to deliver specific agents (cancer cells or other insults) or therapeutic drugs directly to the lung. Such a procedure does not require a tight fitting cannula nor any sophisticated pulmonary function equipment. The novel feature of this method shown here involves a minor surgical procedure that allows the intubation without any possibility of the cannula entering the esophagus. This simple approach enables successful intubation with relatively little training or experience. As many as 30 mice/hr can be treated using this approach with a failure rate approaching zero.

Once the mice are ready to be sacrificed, the injured or cancerous lungs can then be removed for histologic and pathologic analysis. However, in order to properly quantify any histologic variables for comparison with other lungs, it is essential to standardize the fixation procedures and properly quantify the fixed lung volume¹⁰. This paper describes in detail the simple procedures that allow standardized fixation procedures as well as a way to measure the fixed lung volume. The volume is an essential metric in the quantification of the histology, since without such a volume determination, only relative densities can be measured¹⁰. Once the lung volume is known, however, absolute measurements of cells and other structural measurements in the lung can then be quantified.

Protocol

The following protocol describes a system that works well in 20–35 g mice. The method could easily be adaptable to larger or smaller mice simply by changing the catheter size. All animal protocols were approved by the Johns Hopkins University Animal Care and Use Committee.

1. Lung Instillation

1. Select a commercial one-inch long 20 g intravenous cannula to use for the intubation.
2. Modify the catheter tip manually bending it to generate a slight curvature at the tip as illustrated in Figure 1.
3. Anesthetize the mouse with a mixture of ketamine (100 mg/kg) and xylazine (15 mg/kg) injected IP, and confirm anesthesia by the absence of reflex motion. Apply veterinary ointment on the eyes immediately after anesthesia. Immediately after anesthesia apply veterinary ointment on the eyes and give carprofen (5-10 mg/kg SQ) for post surgical and instillation analgesia.
4. Place the mouse supine on a sloped platform. As shown in Figure 2, a large office binder with suture loops taped on works perfectly well.
5. Shave the ventral part of the neck and clean and disinfect the neck area with 70% alcohol.  With new latex- and powder- free gloves, use surgical instruments disinfected with 70% alcohol.
6. Using sharp scissors make a small surgical incision in the neck approximately 12 mm below the lower incisor.
7. With a forceps gently pull the skin in the neck caudally until the ventral wall of the trachea can be seen.
8. Gently retract the tongue and insert the cannula with the bent tip tilted toward the ventral surface of the mouse. As in 1.4, pull gently on the skin in the neck, and insert the cannula into the trachea.
   NOTE: With a little practice, the catheter will be visible moving down the trachea. If it goes in the esophagus, then there will be no visual sighting of the catheter’s movement. No incisions are made in the trachea.
9. Once the catheter is seen in the trachea in the neck, advanced it about 5 mm to be reliably passed the vocal cords but still well above the carina.
10. Prepare to instill up to 50 µl of liquid by injecting through the catheter with a gel loading pipette tip. Place the tip in the luer hub, but before injecting look carefully to observe movement of the fluid in the tip synchronous with the mouse’s breathing. Then inject the instillate.
11. With a 1 ml syringe, immediately do a relatively rapid inflation of 0.6 ml of air into the lungs through the catheter to help distribute the liquid deep in the lungs. Remove the cannula.
12. Remove the cannula. 
13. Use a small amount of cyanoacrylate adhesive to close the small surgical wound as per package insert instructions for VetBond. Place mice in individual cages and visually monitor them until they wake up and behave normally without any indication of discomfort.

2. Lung Fixation

NOTE: Once all experimental procedures are done in a mouse, the lungs can be readied for histologic processing by fixation with formaldehyde (or any other desired fixative).

1. Sacrifice the mouse with an IACUC acceptable procedure. For the representative mouse shown in the video, cervical dislocation of an anesthetized mouse is used.
2. Perform a tracheostomy (if not already done) by surgically exposing the ventral side of trachea, making a small cut, and inserting an 18 G stub needle tip into the trachea, and tying it with thread.
3. Carefully open the thorax with a midline incision, cut away the diaphragm, and remove the lateral chest walls to expose the lungs.
4. Connect the luer end of the needle to a reservoir on a ring stand containing formaldehyde. See Figure 3.
5. Set the top surface of the formaldehyde 25 cm above the level of the mouse. See Figure 3.  Next make sure there is no air in the fixation tubing by running fluid out the end of a stopcock. Connect the luer end of the tracheal cannula to the reservoir tubing. Open the stopcock to inflate the lungs with the formaldehyde. Leave the lungs under pressure for at least 20 min.
6. Open the stopcock to inflate the lungs with the formaldehyde. Leave the lungs under pressure for at least 20 min.
7. Next, tie off the trachea beyond the end of the stub needle.  It may help to pull back slowly on the needle to expose more of the uncannulated trachea.  When tied securely, remove the stopcock.
8. Carefully dissect out the lungs.
9. Place the lungs in formaldehyde overnight.  Longer times are fine, and some stains or procedures may specify specific times.  Also any other liquid fixatives, such as z-fix can be used for the instillation and immersion.
10. Before further histologic processing, measure the fixed lung volume as described next.

3. Measurement of the Fixed Lung Volume

1. Measure the lung volume using Archimedes principle as illustrated in Figure 4. Remove the fixed lung from the formaldehyde and dissect the heart and any other non-lung tissue.
2. Use a previously constructed simple homemade wire support device that is used to keep the lungs fully under water.
   NOTE: This device needs to be made compatible with whichever balance is being used. A typical device shown in Figure 5 is made from plastic pipettes and thin (20 G) wire. This system works well with the balance used in the video, but could easily be adapted to most laboratory balances.
3. Place a beaker with ≈200 ml of water on the balance and tare with the supporting cage in place in the water. See Figure 6. Remove the metal cage; place the lung on the water surface and press under water with the cage.
4. Record the weight on the balance. This number reflects the volume of water displaced and is thus a direct measure of the lung volume. Be careful to make sure the lung or suture or any part of the wire cage does not touch the sides or bottom of the beaker.
5. For accuracy, repeat this measurement. Remove the lung from the water, and dry on a tissue. Tare the beaker with cage in place again and repeat the lung volume measurement. The two volume measurements should then be averaged.
   NOTE: If the lungs are left in the formalin for more than about a week, the air in the lungs will be dissolved in the liquid. When this happens, the lung will sink, so it is no longer necessary to use any device as in Figure 5 to keep the lung submerged. In such case the volume can be measured by simply by holding the lung by one of the suture strings until it is completely submerged as illustrated in Figure 4.

Results

The procedure describe in the first protocol does not by itself lead to any generalized results. It only describes a very reliable means to instill substances directly into the trachea. Figure 7 shows an example of a lung in which trypan blue was instilled with the method described here. There is widespread distribution of the dye, similar to what has been seen with other dyes or tracers given directly into the trachea or mice^11,12. We have also used this method to deliver either bleomycin or ...

Discussion

The procedures described here have several advantages. First the required equipment is simple and inexpensive. Second, the intubation can be quickly done with few errors. Third, the ability to fix the lungs at a constant pressure, and then measure the fixed lung volume allows proper quantification of structures or cells in the lung¹⁰.

One possible downside to the intubation is the minor surgery. This may limit the ability to repeat the procedure if a 2^nd instillation is r...

Materials

Name	Company	Catalog Number	Comments
Laboratory Balance	Ohaus	Adventurer Pro Model AV 313	Other balances can be used if they have a range of 1-300 g
20 g Luer intravenous catheter	Insylte		Several other possible vendors, e.g., Jelco Optiva
500 ml laboratory bottle	Various		Several other possible vendors