Diagnostic Ultrasound Imaging of Mouse Diaphragm Function

Summary

Diagnostic ultrasound imaging has proven to be effective in diagnosing various respiratory diseases in human and animal subjects. We demonstrate a comprehensive ultrasound protocol utilized by Dr. Zuo's lab to analyze diaphragm kinetics specifically in mouse models. This is also a non-invasive research technique which can provide quantitative information on mouse respiratory muscle function.

Abstract

Function analysis of rodent respiratory skeletal muscles, particularly the diaphragm, is commonly performed by isolating muscle strips using invasive surgical procedures. Although this is an effective method of assessing in vitro diaphragm activity, it involves non-survival surgery. The application of non-invasive ultrasound imaging as an in vivo procedure is beneficial since it not only reduces the number of animals sacrificed, but is also suitable for monitoring disease progression in live mice. Thus, our ultrasound imaging method may likely assist in the development of novel therapies that alleviate muscle injury induced by various respiratory diseases. Particularly, in clinical diagnoses of obstructive lung diseases, ultrasound imaging has the potential to be used in conjunction with other standard tests to detect the early onset of diaphragm muscle fatigue. In the current protocol, we describe how to accurately evaluate diaphragm contractility in a mouse model using a diagnostic ultrasound imaging technique.

Introduction

Recently, diagnostic ultrasound imaging techniques have been applied to mouse models of renovascular hypertension and pancreatic cancer^1,2. However, these techniques have not been widely used in rodent respiratory muscle function assay. Therefore, we have developed a diagnostic ultrasound imaging method as a valuable tool for in vivo longitudinal assessments of diaphragm mobility in mice.

There are several advantages to diagnostic ultrasound imaging. For instance, it is noninvasive, safe, portable, and allows for real time measurements at a relatively low cost³. Particularly, certain low frequency ultrasound devices were able to detect air trapping, a clinical characteristic of chronic obstructive pulmonary disease (COPD) with mild to severe airflow limitation⁴. Thus, diagnostic ultrasound imaging may serve as an easily accessible and reproducible screening method for real-time monitoring of respiratory disorders.

Diagnostic ultrasound imaging techniques are frequently applied to larger animals or human subjects. However, there have been a limited number of ultrasound imaging studies on mouse models, which is likely due to the challenges of performing ultrasound on small-scale subjects. The current protocol outlines a novel procedure for measuring diaphragm function in the mouse. In addition, although there have been several rodent studies on diaphragm function, most of the results were generated by isolating muscle strips directly from the euthanized animal^5-7. In contrast, using an in vivo diagnostic ultrasound imaging method for analyzing diaphragm activity would decrease the number of animals sacrificed for experimentation. Furthermore, long-term treatments focused on enhancing diaphragm contractility may be accurately assessed via ultrasound in rodent models without sacrificing animals.

In our lab, we have developed an effective method for visualizing as well as analyzing mouse diaphragm activity using an ultrasound machine, which helps the understanding of diaphragm function in vivo, avoids invasive methods to animals, and aids in the development of therapeutic treatments for respiratory dysfunction.

Protocol

All procedures involving animal subjects were approved and completed in accordance and compliance with The Ohio State University Institutional Animal Care and Use Committee (IACUC) regulations and guidelines.

1. Mouse Anesthesia

1. Set up a clean procedure table with a heated isothermal pad wrapped in a surgical towel. The heating pad should be maintained between 30 °C and 34 °C to stabilize the animal's core temperature while reducing potential stress to the animal.
2. Place the mouse in an anesthesia induction chamber with the following parameters: oxygen flow rate set to 1.5 L/min and isoflurane vaporizer set to 3.5%. Complete sedation should take place within 1-2 min. If an induction chamber is not available, a bell jar may be used with a wire mesh positioned at the bottom to avoid direct animal contact with the isoflurane.
3. Immediately remove the mouse from the induction chamber once it is completely anesthetized (achieved when the mouse loses voluntary motor function). Apply a nose-cone to the animal for maintenance of anesthesia. The oxygen flow rate should be reduced to approximately 0.5 L/min and the isoflurane vaporizer should be set within the range of 1.5 to 2.5%.
   1. Apply a small amount of ophthalmic ointment directly to the corneas to reduce eye dryness⁸. In addition, during anesthesia, the mouse should maintain an absence of the pedal withdrawal reflex, the mucous membranes should remain a pink color, and breathing should appear steady.

2. Preparing for Diagnostic Ultrasound Imaging Procedure

1. Restrain each leg of the mouse on the heated procedure table with a removable adhesive, such as surgical tape.
2. Using an electric razor, remove the hair on the ventral body surface between the abdomen and half way up the thoracic cavity. Apply hair removal cream to further remove the remaining hair that is not cut by the razor. Wipe off the cream with a damp gauze pad after 2-3 min.
3. Remove the excess hair using a water-moistened gauze pad and clean the shaved region with 70% alcohol or equivalent antiseptic. The ultrasound probe will be applied to this area to visualize diaphragm function. A topical analgesic may be provided to animals experiencing minor skin irritation due to hair removal.

3. Diagnostic Ultrasound Imaging Protocol

1. Turn on the ultrasound device and adjust the output power (if necessary) on the apparatus by percentage to obtain optimum resolution.
2. Set the ultrasound machine to either B (brightness)-mode, M (motion)-mode, or both before imaging, which allows for proper visualization of the mouse diaphragm contraction.
3. Apply a small amount of ultrasound gel on the mouse's upper abdomen and massage the gel toward the thoracic cavity.
4. Place the ultrasound transducer in this area and angle it upward towards the heart. Adjust the probe until an optimized resolution of the image is achieved. Note: for this protocol, a micro-convex array or linear phased array transducer is an ideal probe to use due to the small footprint and excellent axial resolution⁹; the frequency needs to be adjusted across the bandwidth and for these experiments a range of 6.5-12 MHz may be utilized.
5. Press the freeze button to temporarily save the diaphragm images and view the selected contractions.
6. Save the recording as a cine loop, which allows for later measurements of diaphragmatic excursion as well as respiration rate. Note: frames of images can be saved in the computer memory or on an external hard drive for future analysis⁹.
   1. Precisely measure the depth of diaphragm movement from relaxation to contraction using the electronic calipers that are part of the ultrasound software.
   2. Convert the cine loop file into a MPEG file and determine the respiration rate by counting the number of diaphragmatic contractions during the recording period. Alternatively, the number of contractions per min (respiration rate) may be counted from the M-mode image.

4. Post Anesthesia Animal Recovery

1. The mouse should completely recover from anesthesia within 1 hour. Do not leave the animal unattended until it has regained sufficient consciousness to maintain sternal recumbency.

Results

A typical ultrasound image of a mouse diaphragm is shown in Figure 1A. The mouse diaphragm maximal vertical displacement was recorded. This distance was calculated by precisely measuring the depth of diaphragm movement from relaxation to contraction using the electronic calipers that are part of the ultrasound software. Table 1 displays these distance measurements of diaphragmatic contractions from three different mice. After converting the cine loop file into a MPEG file, the respiratio...

Discussion

The current experimental protocol develops diagnostic ultrasound imaging techniques specific to the diaphragm activity in a mouse model via a non-invasive, in vivo approach. The anesthesia apparatus settings are approximated values, which may be slightly adjusted for each animal since individual mice may respond differently to anesthesia. To prevent improper anesthesia administration, it is important to regularly monitor the mouse's vital signs including the heart rate, respiration rate, and body temperature...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Veterinary digital ultrasonic diagnostic imaging system	Edan	DUS 3 VET	Ultrasound parameters include: frequency of 6.5 MHz, Depth of 29 mm. Note: An equivalent ultrasound machine may be used for this protocol
Micro-convex array transducer	Edan	C611	Or equivalent
GE Logiq i hand-carried unit (HCU)	GE Healthcare	GE Logiq i hand-carried unit (HCU)	Or equivalent
GE 12 MHz linear array probe	GE Healthcare	12L-RS	Or equivalent
Veterinary anesthetic vaporizer	Webster Veterinary	Serial #: W422021	Isoflurane was exclusively used with this vaporizer (or equivalent). A custom made induction chamber for anesthesia was assembled for initial anesthesia. Maintenance anesthesia was performed using a nose cone
Isothesia (Isoflurane, USP)	Butler Schein	29405 250ML PVL	Or equivalent
Enviro-pure anesthesia absorbing canister	Surgivet Smiths Medical PM, Inc.	Part #: 32373B10	Or equivalent
Ultrasound transmission gel	HM Sonic	N/A	Or equivalent
Puralube vet ointment	Puralube	NDC 17033-211-38	Or equivalent
Deltaphase isothermal pad	Braintree Scientific Inc.	39DP	Or equivalent
Hair remover	Nair	N/A	Or equivalent
Electric razor	Remington	HC-5015	Or equivalent
Surgical tape	3M Micropore	1530-1	Or equivalent
Gauze sponges	Dynarex	3262	Or equivalent