Repeated Orotracheal Intubation in Mice

Medicine

Your institution must subscribe to JoVE's Medicine section to access this content.

Fill out the form below to receive a free trial or learn more about access:

 

Summary

The goal of this article is to describe a refined method of intubation of the laboratory mouse. The method is noninvasive and, therefore, ideal for studies that require serial monitoring of respiratory function and/or instillation of treatments into the lung.

Cite this Article

Copy Citation | Download Citations | Reprints and Permissions

Nelson, A. M., Nolan, K. E., Davis, I. C. Repeated Orotracheal Intubation in Mice. J. Vis. Exp. (157), e60844, doi:10.3791/60844 (2020).

Abstract

The literature describes several methods for mouse intubation that either require visualization of the glottis through the oral cavity or incision in the ventral neck for direct confirmation of cannula placement in the trachea. The relative difficulty or the tissue trauma induced to the subject by such procedures can be an impediment to an investigator’s ability to perform longitudinal studies. This article illustrates a technique in which physical manipulation of the mouse following the use of a depilatory to remove hair from the ventral neck permits transcutaneous visualization of the trachea for orotracheal intubation regardless of degree of skin pigmentation. This method is innocuous to the subject and easily achieved with a limited understanding of murine anatomy. This refined approach facilitates repeated intubation, which may be necessary for monitoring progression of disease or instillation of treatments. Using this method may result in a reduction of the number of animals and technical skill required to measure lung function in mouse models of respiratory disease.

Introduction

The laboratory mouse is a common animal model for human respiratory disease. Thus, there are several published methods for mouse intubation for the purpose of both instillation of treatments and measurement of respiratory mechanics. Most of the described procedures require visualization of the glottis through the oral cavity with specialized equipment such as a laryngoscope or fiber-optic light source1,2,3,4,5,6,7. However, this can be difficult when a relatively large cannula is required, as it can obscure the view of the researcher. Limjunyawong et al.8 have addressed this concern with a method of intubation in which a small cutaneous incision is made along the midline of the ventral neck allowing for visualization of the trachea. Following the procedure, the incision is closed with tissue adhesive.

For studies requiring frequent repeated intubations, successive incising and closure of this site requires debridement of the skin margins and tissue trauma to the ventral neck. The purpose of the transcutaneous tracheal visualization approach to oral intubation is to provide a refined, noninvasive technique specifically suitable for repeated intubation studies as well as single intubation events in mice.

Subscription Required. Please recommend JoVE to your librarian.

Protocol

All animal activities described here have been approved by the Institutional Animal Care and Use Committee (IACUC) of The Ohio State University and were conducted in AAALAC-accredited facilities.

1. Procedure Preparation

  1. Construct the intubation platform. To achieve the appropriate platform slope, use a three-inch (7.6 cm) 3-ring binder. Fold a 15−20 cm length of 3-0 silk or other thread material in half and adhere the ends of the thread to the top of the inclined platform with tape to create a suspension loop (Figure 1).
  2. Select a cannula of the appropriate size and length.
    NOTE: For a 20−30 g mouse, a 1−1.5 inch (2.5−3.8 cm) long catheter up to 18 G can be used. For this study, 18 week-old female BALB/c and 10-week-old C57BL/6 mice (n = 3 of each strain) were used. An opaque white catheter sheath provides the best transcutaneous visualization.
  3. Cut a bevel at the distal tip of the catheter and smooth the cut surface with abrasive paper to create a rounded bevel tip. Gently create a slight bend in the cannula approximately 1 cm from the bevel (Figure 2).
    NOTE: A new catheter should be used for each mouse.
  4. Anesthetize the mouse with ketamine (5.4 mg/g body weight) and xylazine (16 µg/g body weight) administered intraperitoneally.  Apply sterile ophthalimic ointment to the eyes.
    NOTE: Proper anesthetic depth is achieved by lack of response of the mouse to a firm toe pinch.
  5. Suspend the mouse in a supine position on the intubation platform by hooking the upper incisors around the silk thread at the top of the angled surface (Figure 3). Once the mouse is squarely positioned in dorsal recumbency, gently grasp the base of the tail and retract the tail towards the table. Place a piece of tape over the base of tail to secure the mouse.
  6. Apply depilatory cream (Table of Materials) to the ventral cervical region for 30−45 s then remove all depilatory cream from the cervical region using a dry gauze. Repeat application process if needed. Thoroughly rinse the skin with saline or distilled water to remove any residue then wipe dry.

2. Intubation Procedure

  1. Use straight, flat forceps in the nondominant hand to gently retract the tongue in a manner that sufficiently opens the mouth for introduction of the cannula.
    NOTE: Rat tooth forceps should not be used as this will damage the tongue.
  2. With the dominant hand, advance the cannula into the mouth such that the end that is distal to the slight bend is against the roof of the subject’s mouth.
  3. Release the tongue and slide the flat edge of the closed forceps caudally along the ventral neck until the manubrium is reached. This motion laterally displaces the salivary glands and flattens the muscle covering the trachea. The trachea appears transcutaneously as a white line (Figure 3A). If necessary, rotate the forceps in a craniodorsal direction while maintaining tension on the skin in a caudal direction to cause the laterally displaced salivary glands to peak. This maneuver creates more contrast around the trachea (Figure 3B).
    NOTE: Avoid excessive force on the ventral neck as it can collapse the trachea and impair breathing.
  4. Advance the cannula while simultaneously angling the distal tip of the cannula ventrally by supination of the dominant hand with simultaneous flexion of the wrist.
  5. The proper placement of the cannula is indicated by visualization of the opaque cannula in the trachea (Figure 4B,D). If the cannula has been advanced past the level of the origin of the masseter muscle and visualization of the cannula in the trachea has not been confirmed, retract the cannula and reattempt the maneuver.
  6. Confirm proper cannula placement by connecting a lung inflation bulb to the cannula and observing thoracic expansion with concurrent depression of the device.
  7. Without displacing the cannula, carefully unhook the incisors of the mouse from the intubation platform. Move the mouse to a horizontal platform (Table of Materials) and insert the cannula to the adaptor on the ventilator. Following the deep inflation, ventilate the mouse for 60 s then measure respiratory resistance.

3. Recovery

  1. Once the procedure is complete, move the mouse to a warmed platform. Provide constant stimulation via light toe or tail pinches to encourage spontaneous respiration.
  2. Extubation can occur when the mouse just begins to chew. Grasp the cannula at the level of the hub and gently pull the tube cranially and away from the mouse until the cannula is completely removed from the subject’s mouth.
    NOTE: It is preferable to provide airway support with the rigid cannula for as long as possible during the recovery process.
  3. Once extubated, transfer the mouse to a clean recovery cage with heat support. Continuously monitor the mouse until it is fully ambulatory, and recovery is complete.

Subscription Required. Please recommend JoVE to your librarian.

Representative Results

Serial monitoring of baseline pulmonary function
Eighteen-week-old female BALB/c and 10-week-old C57BL/6 mice (n = 3 of each strain) were intubated using the described method on day 0, 3, 10, and 17. Following intubation on each day, the subject was connected to a mechanical ventilator supplied with 100% oxygen (Table of Materials). Respiratory resistance (Rrs) was measured using the forced oscillation technique for 60 s following a deep inflation to 25 cm H2O held for 5 s. No software errors associated with this sustained breath hold along with Rrs values within physiological range provide additional support for proper placement of the cannula. Data revealed no significant differences of measured Rrs observed between time points within each strain (Figure 5). It is assumed that the absence of an increase in Rrs over time indicates lack to trauma-associated inflammation in the respiratory system over four successive time points.

Statistical analysis
Descriptive statistics (mean and standard error) were calculated using statistical analysis software (Table of Materials). The Kolmogorov-Smirnov method was used to verify the Gaussian data distribution. Statistical analyses of datasets were made by unpaired ANOVA, with a post hoc Tukey-Kramer multiple comparison post-test. All data are presented as mean ± SEM. P < 0.05 was considered statistically significant.

Figure 1
Figure 1: Intubation platform. The intubation platform consists of a three-ring binder with a loop of silk thread adhered to the top of the binder to create a suspension loop. Please click here to view a larger version of this figure.

Figure 2
Figure 2: Cannula preparation. (A) Lateral view of the prepared cannula. Note the gentle angle created approximately 1 cm from the rounded bevel at the distal end of the catheter and the orientation of the cannula angle in relation to the bevel. (B) Dorsoventral view of the prepared cannula. Note the rounded and smoothed edge of the bevel. Please click here to view a larger version of this figure.

Figure 3
Figure 3: Tracheal visualization. (A) Forceps are placed on the ventral neck and the skin is gently retracted caudally to laterally displace the salivary glands and provide visualization of the trachea as a white structure on ventral midline (black arrow). (B) Craniodorsal rotation of the forceps on the ventral neck creates a protrusion of the salivary glands (*). The trachea is visualized as the white linear structure on ventral midline between the salivary glands (black arrow). Please click here to view a larger version of this figure.

Figure 4
Figure 4: Proper cannula placement. (A) C57BL/6 mouse positioned on the intubation platform with the cannula introduced into the proximal oral cavity. (B) C57BL/6 mouse with the cannula properly placed in the trachea. Note the cannula can be easily visualized as the white structure within the trachea (white arrow). (C) BALB/c mouse positioned on the intubation board with the cannula introduced into the proximal oral cavity. (D) BALB/c mouse with the cannula properly placed in the trachea. The white cannula can be easily visualized within the trachea (black arrow). Please click here to view a larger version of this figure.

Figure 5
Figure 5: Serial measurement of resistance. No significant differences of measured Rrs observed between time points within each strain. Please click here to view a larger version of this figure.

Subscription Required. Please recommend JoVE to your librarian.

Discussion

Intubation using the transcutaneous tracheal visualization technique offers a refined approach to the standard skin incision method. With special attention to several key steps, intubation can be easily and quickly achieved. The animal must be placed squarely in dorsal recumbency on the intubation platform with the mouse secured in gentle retraction. This will extend the animal into vertical alignment and proper positioning for intubation. In addition, the depilatory cream should not remain in contact with the animal’s skin for longer than 30−45 s and should be thoroughly rinsed to removal all residue. Extended skin contact with the depilatory cream will cause unnecessary pain for the animal and ulcerations can obstruct the view of the trachea9. It is imperative to use the proper wrist motion as the dominant hand introduces the catheter into the glottis. The dominant wrist should flex while the hand moves in a supination motion. It is also critical to monitor the subject closely as the flat edge of the forceps are pressed on the ventral neck to visualize the trachea. Pressure from the forceps will occlude the trachea and cause hypoxia if maintained for a prolonged duration. If the patient appears cyanotic, allow a brief pause for mucus membranes to return to a pink color and for respiration to stabilize before repeating attempts.

Extensive mouse intubation experience was not necessary to perform this technique. The most common complications in inexperienced individuals include laryngeal trauma and upper airway inflammation due to multiple intubation attempts. Close monitoring is necessary during the recovery of these patients as medical intervention with nonsteroidal anti-inflammatories may be indicated. Repeated unsuccessful intubation attempts may result in tissue trauma and inflammation of the distal oral cavity, which could result in upper respiratory noise, dyspnea, hypoxemia, prolonged recovery, inability to perform repeated intubation or death.

Several modifications are recommended in the event that intubation is not successful. First, ensure the bevel of the cannula is smooth, rounded and cut to the appropriate length for the animal’s size. The bevel edge may be smoothed using abrasive paper to minimize tissue trauma and facilitate intubation7. In addition, check that the cannula exhibits a slight curve of approximately 15° at one-third distance away from the bevel and the tip of the cannula is beveled at a 45° angle as described in Brown et al.6. Always check that the catheter is in the proper orientation before and while performing this procedure.

For this study, mice were intubated for repeated lung function tests using a mechanical ventilation system to record lung function measurements. A large, 18 G cannula was used to create a tight seal. To perform repeat lung function studies on mice with a smaller tracheal diameter due to age or strain, it may be challenging to place a larger cannula. If a smaller cannula is elected for use, ensure that a proper seal can still be achieved, and that the resistance of the cannula is not higher than resistance of the test subject’s airway10. A successful deep inflation perturbation is adequate confirmation of an appropriate seal. Note that such a seal is unnecessary if only installation of treatments into the lung is desired.

Although the described method has made modifications that prevent external tissue damage, the upper limit of frequency of intubation is still a function of cumulative trauma to the glottis and trachea due to excessive introduction of the cannula. Concurrent monitoring of a control group for significant increases in airway resistance during a study is recommended since tissue trauma is accompanied by inflammation that will result in decreased luminal diameter of the trachea. Significant increases in airway resistance over the course of repeated intubation procedures were not observed in the current study. Mice remained clinically normal for the study duration and gross necropsy of upper airway structures was unremarkable at study conclusion in all animals.

In summary, the described intubation technique offers a noninvasive method for placing endotracheal cannulas with minimal equipment including an inclined surface, forceps, a polypropylene cannula and depilatory supplies. This refined method enables repeated intubation events without recurrent tissue trauma and pain associated with a cutaneous incision site on the ventral neck or a tracheotomy procedure. In addition, this method reduces the number of mice required as individual mice may be repeatedly intubated throughout the course of a study. It also eliminates the need for specially designed intubation restraint devices, scopes or transilluminating equipment for airway visualization. BALB/c and C57BL/6 strains were used in this study to demonstrate technique success in both light and dark pigmented strains and animals of a relatively young age and small size (10−20 week-old mice). This refined technique is suitable for single or repeated intratracheal instillation of compounds, bronchoalveolar lavage, imaging or lung function testing. This minimally invasive, versatile method can be implemented for virtually any procedure that requires access to the lower respiratory tract.

Subscription Required. Please recommend JoVE to your librarian.

Disclosures

The authors have nothing to disclose.

Acknowledgments

The authors thank Lucia Rosas, Lauren Doolittle, Lisa Joseph and Lindsey Ferguson for their technical assistance and the University Laboratory Animal Resources for their animal care support. This work is funded by NIH T35OD010977 and R01-HL102469.

Materials

Name Company Catalog Number Comments
18 G x 1 1/4" intravenous catheter, Safelet Fisher Scientific #14-841-14 Cannula for intubation
70% ethanol, 10 L Fisher Scientific 25467025 Cleaning cannula
Abrasive paper (sandpaper) Porter-Cable 74001201 Cannula preparation
AnaSed (xylazine sterile solution) injection (100 mg/mL) Akorn Animal Health NDC# 59399-111-50 Anesthesia
Blue labeling tape (0.5 in x 14 yds) Fisher Scientific 15966 Restraint on intubation platform
Braided silk suture without needle, nonsterile, (3-0) Henry Schein Item #1007842 Intubation platform
Deltaphase Isothermal Pad Braintree Scientific 39DP Mouse thermoregulation and recovery
Deltaphase Operating Board Braintree Scientific 39OP Mouse recovery (prior to extubation)
Distilled water ThermoFisher 15230253 Cleaning mouse following depilation
Eye Scissors, angled, sharp/sharp Harvard Apparatus 72-8437 Cannula preparation
FlexiVent (FX Module 2) Scireq N/A Record lung function data (not required to perform procedure, used in this study to validate procedure)
Gauze sponges Fisher scientific 13-761-52 Hair removal
Heavy-Duty 3" 3-Ring View Binders Staples 24690CT Intubation platform
Instat Software Graphpad N/A Statistical analysis software
Insulin syringe (0.5 cc, U100) Fisher Scientific 329461 Anesthesia administration
Ketamine HCl Injection, USP (100 mg/mL) Llyod Laoratories List No. 4871 Anesthesia
Lung inflation bulb Harvard Apparatus 72-9083 Confirm cannula placement
Micro Forceps, Curved, Smooth Harvard Apparatus 72-0445 Retract tongue and create tension on neck for cannula visualization
Nair (hair removal lotion), 9 oz bottle Church & Dwight 42010440 Hair removal
Sterile saline (0.9%), 10 mL Fisher Scientific NC9054335 Anesthesia, cleaning skin following hair removal

DOWNLOAD MATERIALS LIST

References

  1. Spoelstra, E. N., et al. A novel and simple method for endotracheal intubation of mice. Laboratory Animals. 41, (1), 128-135 (2007).
  2. Rivera, B., Miller, S. R., Brown, E. M., Price, R. E. A Novel Method for Endotracheal Intubation of Mice and Rats Used in Imaging Studies. Contemporary Topics in Laboratory Animal Science. 44, (2), 52-55 (2005).
  3. Sparrowe, J., Jimenez, M., Rullas, J., Martinez, A. E., Ferrer, S. Refined Intratracheal Intubation Technique in the Mouse, Complete Protocol Description for Lower Airway Models. Global Journal of Animal Scientific Research. 3, (2), 363-369 (2015).
  4. Deyo, D. J., Wei, J. A Novel Method of Intubation and Ventilation in Mice. Anesthesia & Analgesia. 88, (2), 179 (1999).
  5. Vergari, A., et al. A new method of orotracheal intubation in mice. European Review for Medical and Pharmacological Sciences. 8, (3), 103-106 (2004).
  6. Brown, R. H., Walters, D. M., Greenberg, R. S., Mitzner, W. A method of endotracheal intubation and pulmonary functional assessment for repeated studies in mice. Journal of Applied Physiology. 87, (6), 2362-2365 (1999).
  7. Das, S., MacDonald, K., Chang, H. S., Mitzner, W. A Simple Method of Mouse Lung Intubation. Journal of Visualized Experiments. (73), e50318 (2013).
  8. Limjunyawong, N., Mock, J., Mitzner, W. Instillation and Fixation Methods Useful in Mouse Lung Cancer Research. Journal of Visualized Experiments. (102), e52964 (2015).
  9. Qin, W., Baran, U., Wang, W. Lymphatic response to depilation-induced inflammation assessed with label-free optical lymphangiography. Lasers in Surgery and Medicine. 47, (8), 669-676 (2015).
  10. McGovern, T. K., Robichaud, A., Fereydoonzad, L., Schuessler, T. F., Martin, J. G. Evaluation of Respiratory System Mechanics in Mice using the Forced Oscillation Technique. Journal of Visualized Experiments. (75), e50107 (2013).

Comments

0 Comments


    Post a Question / Comment / Request

    You must be signed in to post a comment. Please or create an account.

    Usage Statistics