Evaluation of Lung Function in an Ex vivo Murine Model

The present protocol describes a stepwise method for analyzing the respiratory mechanics of an ex vivo murine model using the forced oscillation technique (FOT).

Respiratory mechanics are a key area of study in defining and treating lung pathologies by assessing functional lung capacity. Lung mechanics can be evaluated through various lung maneuvers that involve different oscillatory waveforms. When applied to the lungs, these maneuvers measure multiple variables, such as pressure, volume, and flow, based on the response to the waveforms. These signals are then computed and analyzed to determine parameters such as hysterisivity, resistance, compliance, tissue damping, and tissue elastance, providing a detailed assessment of overall lung function. The analysis of respiratory mechanics is particularly important in evaluating donor lungs for lung transplantation. The present protocol is the first of its kind, offering a comprehensive and reproducible stepwise method for assessing respiratory mechanics using an ex vivo murine model. It includes details on the selected animal model, lung recovery, storage and preservation, and experimentation using a forced oscillation technique-based system. Additionally, it outlines data analysis, clinical significance, and the applications of the forced oscillation technique in studying an ex vivo model.

Lung transplant represents the only durable treatment for end-stage lung diseases. Approximately 4,600 people receive lung transplants each year worldwide, but almost 600 patients die on the waitlist secondary to the shortage of suitable donor lungs^1,2. In efforts to increase the pool of available lungs, donor allocation systems are continuously adjusted, which has led to surgeons traveling farther distances to secure donor organs³. The increased distances invariably increase the cold ischemic time, presenting a need for additional methods of organ preservation.

The current standard for donor organ preservation of lung transplantation is cold static preservation at 4 °C, limiting preservation time to 6-8 h - a small window of viability for transplantation⁴. However, with longer travel distances and resultant increased ischemic times, the assessment of lung function prior to transplantation is critically important⁴. With evolving policies for lung transplantation, novel research has been conducted to address this need. Recently, studies have suggested that cold static preservation at 10 °C is a more optimal storage temperature for lung preservation with resultant improvement in lung function, resistance to injury, and comparable rates of primary graft dysfunction when implanted^4,5,6,7,8. Furthermore, research centered on ex vivo lung perfusion (EVLP) has shown significant improvement in donor lung utilization and transplantations without detriment to recipients⁹. While the use of EVLP for expanding the donor pool for lung transplantation and extending the preservation time is well documented, this technology is expensive, time-intensive, and requires specialized training to perform¹⁰. As such, there is a need for additional methods to study ex vivo lung function that are comprehensive, inexpensive, and reproducible.

Traditional measures of pulmonary mechanics, e.g., compliance, resistance, elastance, and pressure-volume curves, can be reliably determined using body plethysmography or with ventilator techniques using a single-compartment model. More detailed mechanics can be obtained using the forced oscillation model to fit the constant phase model, which can partition airway mechanics into central and peripheral compartments (Newtonian resistance, tissue damping/elastance, hysteresivity)^11. While the application of these techniques is reproducible and comprehensive, a limitation thus far has been the requirement of performing such measures in an in-vivo model, presumably as the exsanguinated lung loses structure at the alveolar entrance ring¹². This study used a commercially available forced oscillation technique-based small rodent ventilator with the aim of developing an ex vivo model to better characterize lung mechanics for lung transplant applications.

This study was approved by the Committee on Animal Research in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals. C57Bl/6 wild-type mice, aged 6-8 weeks and weighing between 18-28 g, were used. Details of the reagents and equipment are provided in the Table of Materials.

1. Preparation

1. Use an operating microscope with up to 20x magnification for all surgical procedures.
2. Clean and sterilize all surgical instruments before beginning the procedure. Autoclave instruments or use a suitable sterilizing solution to maintain aseptic conditions.

2. Extraction of donor lungs

1. Perform all operations under sterile conditions.
   NOTE: This step is performed in a dedicated surgical space and under sterile conditions.
2. Place the mouse into an anesthetic induction chamber and induce anesthesia with 5% isoflurane in oxygen and maintain anesthesia with 3.5% isoflurane in oxygen throughout the procedure (following institutionally approved protocols).
3. Record mouse weight prior to the first incision.
4. Secure the mouse on the operating table and ensure proper depth of anesthesia by applying pressure to a toe. If the mouse withdraws in pain, increase anesthesia asneeded.
5. Remove the lungs from the donor following standard procedure¹³.
   NOTE: The lungs and heart were extracted en block to perfuse the lungs ex vivo, and a portion of the trachea was removed for later intubation.

3. Lung storage and preservation

1. Once the lungs are removed from the donor, intubate the trachea with an 18 G intravenous angio-catheter by carefully advancing the catheter, ensuring it does not puncture the trachea.
2. Secure the trachea over the cannula using a 3-0 silk suture, ensuring a tight seal. Advance the 18 G intravenous angio-catheter until it is within the right ventricular outflow tract (RVOT), just past the pulmonary valve.
3. Record the time lungs were removed from the body.
4. Store the lungs in a 50 mL conical tube containing the commercially available preservation solution at 4 °C or 10 °C overnight.

4. Setup and calibration

1. Begin by powering on the system and/or software.
2. Initiate the program, click on Create a new study button, and follow the on-screen prompts to define the protocol and assign subjects.
3. Click on Experimentation Session and sign in using the first and last initials of the user.
4. Begin labeling subjects by a consistent naming convention.
5. Enter the sex, birthdate, and weight information of each subject.
6. Select and assign subjects to the measurement site and confirm weights.
7. Continue setup and calibration of the software by following on-screen prompts.
   NOTE: During calibration of the tubing, the same type of cannula used in the intubation of the donor lungs was attached to the Y-tubing of the system to ensure consistency.
8. Repeat calibration if calibration values fall outside of the accepted ranges.
9. Cancel prompts to begin ventilation until ready. Experimental sessions with ventilation and data recording can be initiated later.

5. Lung ventilation and data acquisition

1. Remove lungs from the 50 mL conical tube. Flush the right ventricular outflow tract (RVOT) using the previously placed 18 G intravenous angio-catheter with additional preservation solution at a dose of 60 mL/kg to flush lung capillaries.
   NOTE: Great care must be taken to avoid dehydration of donor lungs as this is a known confounder in studying lung mechanics.
2. Secure donor lungs to the ventilator machine and begin the experiment.
3. Click on Start Ventilation, and when ready, press Start Recording to begin the experimental session.
   NOTE: The ventilator utilized for this study was set to have a respiratory rate of 150 breaths/min, a tidal volume of 10 mL/kg, and a PEEP of 3.
4. Run each task by double-clicking on the task for lung mechanics assessments from the task list on the right side of the page.
5. Run the Deep Inflation task located in the task list to further ensure that atelectatic regions are recruited and lung volumes have been standardized. For details, see the Results section.
6. Proceed with the sequence of tasks.
   NOTE: Lung function can be assessed through multiple tasks evaluating mechanical properties. Deep Inflation standardizes lung volume, while Prime-8 stabilizes lung mechanics. PV-P and PV-V measure static and dynamic compliance, airway resistance, and conductance. Snapshot 150 provides a rapid assessment of resistance, compliance, and elastance, whereas QuickPrime evaluates airway and tissue resistance along with lung viscoelasticity. These tasks collectively ensure a comprehensive analysis of lung function. In this article, the data for the tasks performed in Deep Inflation, Snapshot 150, and QuickPrime (using commercially available equipment) are provided as representative results. Lung volumes were standardized between perturbations using the Deep Inflation task in order to minimize confounding variables when interpreting the data.
7. Run the chosen sequence of tasks in triplicate.
8. Click on Stop Recording and then Stop Ventilation.
9. Export data individually or continue to the next subject and repeat steps 5.1-5.9.
   NOTE: For data derived from the Snapshot 150 and QuickPrime perturbations, a COD of >0.95 is required for reliable analysis.

A graphical depiction of the experimental design is provided for the mouse model (Figure 1). Lungs were inflated using a commercially available forced oscillation technique-based small rodent ventilator system to assess the respiratory mechanics of the donor tissue under various conditions (Figure 2). When comparing the results between the groups of preserved donor lungs, groups of donor lungs that were stored at 10 °C were found to perform better than all groups stored at 4 °C in all parameters. In the 10 °C storage groups, the donor lungs were found to have a significant decrease in resistance and a significant increase in compliance when assessed with the Snapshot 150 perturbation. When using the QuickPrime perturbation, trends toward decreased tissue elastance, tissue damping, and hysteristivity were noted in the 10 °C groups (Figure 3). The underlying mechanism of improvement in lung function within this group is under investigation. Preliminary data demonstrates the addition of cyclic stretch leads to less lung injury and greater protection of mitochondrial health¹⁴. This preliminary data demonstrates the efficacy, efficiency, and reproducibility of FOT in studying the respiratory mechanics and lung function of ex vivo donor lungs.

Figure 1: Experimental design of functional lung assessment of an ex vivo murine transplant model. The lungs and heart were procured en block from a donor mouse (A) and stored in the preservation solution in cold static conditions for 24 h (B). The lungs and heart en block were removed from the solution and secured to the ventilator system after 24-h preservation and functional lung assessments were conducted (C). Please click here to view a larger version of this figure.

Figure 2: Donor lungs before and after deep inflation. Donor lungs before deep inflation (A) and donor lungs after deep inflation (B). Please click here to view a larger version of this figure.

Figure 3: Respiratory function measurements on donor mice lungs. Results of physiologic evaluation including resistance, compliance, tissue elastance (H), tissue damping (G), and hysterisivity after 24 h of storage in 4 °C static, 10 °C static, or 10 °C ventilated conditions. *p < 0.05; data expressed as mean ± SD. N = 4 for each storage condition. Please click here to view a larger version of this figure.

Importance and potential applications
Respiratory mechanics are routinely used in various applications to study lung pathology and lung injury. The study of respiratory mechanics has been described many times for the progression of diseases such as ARDS and in cases of assisted ventilation but has been described far less in the literature as it pertains to organ transplantation^{15,16,17,18,19}. Additionally, as lung allocation policy changes and donor lung storage techniques continue to evolve, it is critical to analyze the respiratory mechanics of lungs to optimize transplant outcomes. As such, there is a need for research on ex vivo animal models to simulate and compare donor lung storage conditions. Here, an ex vivo murine model was used to analyze functional lung capacity when donor lungs were subjected to various cold storage conditions.

The results of these findings indicate that respiratory mechanics can be reliably measured in an ex vivo setting. This has important clinical applications as this protocol describes a method for quantifying lung function that is efficient, inexpensive, reproducible, and requires little to no specialized training to perform.

Critical steps
Critical steps include securing the trachea tightly over the 18 G cannula to ensure an airtight seal. The occurrence of air leaks could drastically affect the accuracy of data collection. Additionally, calibration of the ventilator system is essential for accurate data recording. All downstream computation and analysis rely on precise calibration values that fall within accepted ranges. Finally, flushing the lungs with additional Perfadex prior to conducting experimentation is a critical step as this flushes lung capillary networks.

Modifications and troubleshooting
While most procurements can be completed without modification, repetition of experiments may be necessary if air leaks are found. Particular attention should be taken when dividing the inferior pulmonary ligament to avoid air leaks during donor procurement. Furthermore, blunt forceps or cotton tips are preferable for manipulating the lungs to avoid puncturing the lungs and/or trachea.

Limitations
This experiment was conducted on approximately 12 laboratory mice (n = 4 in each storage condition), yielding a small sample size. Further experimentation is needed to increase the generalizability of the results. Furthermore, donor lungs were not transplanted into recipient mice, and lung function after transplantation was not measured or recorded. Therefore, the functional results are purely preliminary and do not adequately compare preservation techniques on the functional capacity of lungs once transplanted.

The authors declare the research was conducted without any commercial or financial relationships that could be misconstrued as a conflict of interest.

The authors would like to thank Sophie Paczensy for the use of the ventilator system, and Colin Welsh for his assistance. Figure 1 was created using biorender.com. This research was supported by a grant from the South Carolina Clinical and Translational Institute (NIH/National Center for Advancing Translational Sciences ) under award number UL1-TR001450.