A Mouse Model of Neonatal Continuous Positive Airway Pressure Administration and the Precision Lung Slice Method for Measuring Airway Contractility

Summary

We demonstrate a noninvasive method to deliver varying levels of CPAP to unanesthetized neonatal mice. The technique is intended to partially mimic CPAP administration for many preterm infants to assess its effects on lung development. We also demonstrate CPAP's effects on airway contractility using the precision lung slice method.

Abstract

Mechanical forces associated with positive-pressure respiratory support used in the intensive care of preterm infants can have both positive and inadvertent consequences for lung development. Positive pressure support improves pulmonary gas exchange, limits atelectasis, and improves systemic oxygenation, but the injurious effects of invasive modalities, such as mechanical ventilation, on lung development are also well known. However, there are major gaps in our understanding of the adverse effects of less invasive forms of ventilation. Here, continuous positive airway pressure (CPAP) is one of the most common respiratory support modalities, representing the least invasive of the positive pressure spectrum. A major limitation in studying human airway development is the lack of age-appropriate models and the ability to administer CPAP to small neonatal animals. We resolved this problem by developing a mouse model of neonatal CPAP delivered daily to awake, un-anesthetized newborn mice -- a model that utilizes clinically relevant CPAP levels at a stage of mouse lung development that corresponds with preterm infants when they are likely to receive CPAP in the ICU. Here, we demonstrate our CPAP model, which includes a convenient, custom-designed system that can deliver CPAP non-invasively to the un-anesthetized newborn with easily adjustable levels. We also provide a demonstration of the precision cut lung slice method for the measurement of airway hyperreactivity ex vivo. The PCLS is described here because it provides a convenient functional readout of the pulmonary system, which can be technically challenging in small (neonatal) animal models, and also has wide applicability to other models unrelated to CPAP. We show data demonstrating that CPAP elicits airway hyperreactivity, which may have important clinical implications for the choice of respiratory support modality in the intensive care of preterm infants.

Introduction

Respiratory-related concerns, such as apnea and respiratory distress, for many preterm infants require life-saving modes of respiratory support, which, unfortunately, contribute to infant morbidity and prolonged hospital stays. Supplemental O₂ and positive pressure respiratory support, including mechanical ventilation (MV) and less-invasive CPAP, are common respiratory support modalities. However, the major short- and long-term problem for survivors of the critical period in prematurity remains chronic bronchial airway disease^1,2,3,4, manifested as life-long wheezing/asthma, poor pulmonary function, and decreased exercise capacity. Although the injurious effects of supplemental O₂ and MV have been widely implicated in the pathogenesis of bronchopulmonary dysplasia (BPD) and longer-term respiratory morbidities for decades^5,6,7,8, some evidence suggests that noninvasive forms of respiratory support, such as CPAP, could also have unintended effects on the developing lung^6,9,10,11,12. However, very little is known about the adverse effects of CPAP, largely because of the technical challenges associated with administering it non-invasively to neonatal animals.

Given the overall rising rate of pediatric asthma¹³, and increased survival rates of preterm infants beyond the NICU stay, understanding the mechanisms initiating adverse structural and functional changes of the immature tracheobronchial airways and how they contribute to life-long pulmonary disorders independently of supplemental O₂ is an important unmet clinical and research need. Emerging evidence suggests neonatal CPAP may have long-term adverse effects on airway function and contribute to wheezing disorders, which is significant since CPAP has become the preferred respiratory support modality in efforts to avoid O₂ toxicity from supplemental O₂. The technical challenges associated with administering CPAP to small and age-appropriate neonatal animals have hindered progress in our understanding of CPAP effects (whether beneficial or detrimental) on lung development. We resolved this problem by developing the first neonatal mouse model of CPAP^{10,11,12,14,15} delivered daily (with varying levels of CPAP for 3 h/day) to awake, un-anesthetized mice from birth onwards, thus permitting study of clinically relevant levels of CPAP at a lung development stage comparable to very preterm (~24-28 week fetus) infants who are likely to receive CPAP^16,17. We have performed MRI scans in the neonatal mouse model and have confirmed that CPAP results in lung inflation¹¹ and that CPAP for the first postnatal week causes a long-term (3 weeks of age) increase in airway reactivity (i.e., 2 weeks post-CPAP)^10,11,12,14, which aligns with wheezing in toddlers (3-4 years of age). Here, we provide a detailed demonstration of our custom-designed CPAP system for newborn mice and describe a convenient method of assessing airway contractility using the ex vivo precision lung slice method (PCLS).

The PCLS provides an excellent and convenient opportunity to investigate airway hyperreactivity mechanisms and their respective pathways. Using the PCLS method, individual airways are imaged in real-time to measure airway narrowing in response to a bronchoconstrictor such as methacholine after pretreating airways with specific receptor/channel inhibitors. These studies are important for understanding the pathophysiological consequences of CPAP in prematurity to provide targetable and actionable interventions that permit optimization of necessary clinical care while mitigating potential life-long consequences.

Protocol

Experiments are performed on time-pregnant mice, which are monitored daily to determine the day of birth. Mice are treated with or without CPAP in 21% O₂ for the first week of postnatal life. Typically the lungs are removed in preparation for the measurement of AW reactivity using the precision lung slice (PCLS) method 2 weeks after CPAP, although such measurements are possible in as young as P8 days (immediately after CPAP) or into adulthood. All procedures were carried out in accordance with the National Institute of Health (NIH) guidelines for care and use of laboratory animals and were approved by the Animal Care and Use Committee at Case Western Reserve University.

1. Assembly of the CPAP mask

1. Construct an individual mask from the cut end of a 5 mL syringe containing the Luer lock section for connection of tubing to enable delivery of airflow and positive pressure to the airways of the mouse (Figure 1A).
2. Predrill the syringe component of the mask to accommodate a threaded Luer-Lock connected to an elbow, Luer to hose barb fitting, and standard aquarium tubing (3/8" diameter).
3. Use the side-port tubing to allow excurrent airflow to pass through a downstream adjustable leak to set the backpressure to the lungs of the mouse and a custom-made manometer to monitor the CPAP level in real-time.

2. Collar design

1. Construct the collar from cut fingertip sections of small-sized standard lab gloves. Use it to connect the mouse to the mask unit.
2. Use a hole-punch (1/4 inch) to create a hole in the apex of the cut fingertip, which forms a uniform and consistent size opening to accommodate the face and head of the mouse.
   NOTE: The hole-punch ensures a consistent mask size, but larger hole-punches are needed around P4 to accommodate the mouse's growth.

3. Construction of the manometer

1. Construct a manometer from two disposable serological pipets connected via curved aquarium tubing (Figure 1A). Ensure the manometer is half-filled with water containing a droplet of colored dye to aid in visualization of the CPAP level.
2. Connect the manometer in series with, and downstream of, a flow controller (flow rate ~600-700 mL/min) and the mask.

4. Fitting the collar and mask on the mouse

1. To place the collar on the mouse (1-day old), first position it onto the open end of the mask, which, via careful maneuvering, needs to be stretched so that it can accommodate the face and head of the mouse.
2. Insert the face of the mouse through the opening of the collar, which is gently relaxed from its stretched position over the face/head. At ~4 days of age, cut small slits in the opening of the collar to accommodate growth and the larger head of the mouse, although a larger-sized whole-punch is also recommended.

5. Administering CPAP to multiple mice simultaneously

1. Make a system capable of delivering CPAP to multiple (up to 10) neonatal mice using a series of flow controllers and manometers (Figure 1B). Provide the upstream flow of air by a regular air tank with a standard regulator and pass it through a flow controller that provides the overall flow to all CPAP devices and mask/collar units.
2. Bubble the air in a reservoir of water for humidification before passing it directly to the CPAP units, and administer CPAP to each animal independently.
   NOTE: A control mouse receives a mask, the same airflow, but without any back pressure or CPAP.
3. Conduct all experiments on a heat pad with re-circulated water at ~38 ^oC to maintain a stable body temperature.

6. Timeline

1. Monitor pregnant mice daily to determine the date of birth and then perform the initial CPAP session the following day (P1 days of age) for 2 h (Figure 2) to minimize the duration of time the pups are away from a nursing dam.
2. For subsequent days (P2-P7), conduct each CPAP session for a 3 h duration. Perform CPAP longer than 7 days, if needed, and at any given level of CPAP, although 7 days of CPAP at 6 cmH₂O is used routinely.
3. Then, assess the animals for CPAP effects on airway reactivity either immediately following (P8) the end of CPAP (P7) or longer-term effects (e.g., P21 or adulthood) using the precision lung cut preparation (section 8).

7. Tissue harvest and airway contractility

1. Harvest lungs for molecular or physiological techniques (e.g., polymerase chain reaction (PCR), western blots, immunohistochemistry, RNASeq, proteomics, metabolomics, etc.).
   NOTE: The primary focus is on CPAP effects on airway contractility, which can be assessed in vivo (anesthetized and mechanically ventilated, P21 days or older) for respiratory system mechanics or ex vivo using the PCLS (P8 days or older). The latter has major advantages for investigating compounds or agents that target specific pathways that may not be suitable for use in vivo studies.

8. Whole-Lung extraction/preparation for the precision lung slice (PCLS) method

1. Euthanize the mouse via anesthetic overdose (intraperitoneal injection of a ketamine/xylazine mix; 100 mg/kg/10 mg/kg, respectively) to prepare the lungs for ex vivo measurements of airway reactivity to bath-applied methacholine.
2. Place the mouse supine for cannulation of the trachea using an agarose-filled syringe. Insert the cannula through a small ventral neck incision and advance it ~3 mm before being secured with suture.
3. Gently inject liquefied agarose (40 °C) to inflate the lungs (0.4 mL for P8 mice; 0.8 mL for P21 mice) and place the mouse in the refrigerator for 30 min to allow the agarose to cool and gel.

9. Lung sectioning for the PCLS method

1. Once the agarose has cooled, embed a lung lobe (of choice) in 2% agarose, place it on a vibratome, slice into 300 µm sections, and immerse in DMEM + Pen/Strep solution for overnight incubation (5% CO₂, 37°C).
2. The following day, rinse the lung slices in HBSS and mount them in a recording chamber for live imaging of airway responses to methacholine challenge.
   NOTE: Results presented represent naïve (control, i.e., no CPAP) and CPAP treatment only, and no agonists or antagonists/inhibitors are used.
   1. Perform airway imaging.
      1. Place the lung slices on a custom-made perfusion chamber and cover them with a thin, lightweight sheet of mesh and a coverslip, which are held in place with silicone grease.
      2. Then, mount the recording chamber containing the slice on a microscope and perfuse (7 mL/min) continuously with HBSS at room temperature using a perfusion pump.
      3. Use a microscope mounted with a camera to identify individual airways. After an initial 3 min period of baseline recording, perfuse the chamber with increasing concentrations of methacholine (2 min each concentration) and record changes in airway lumen area continuously.
   2. Quantify airway size.
      1. Identify a single image (ImageJ software) of an airway for a given concentration of methacholine (and baseline) and quantify the lumen area by selecting the adjust threshold to ensure the lumen area is fully captured (Figure 3A).
      2. Then, apply these settings to subsequent images of the same airway at different concentrations (i.e., the same threshold is used for a given airway).
      3. Then, select the lumen area for measurement using the wand-tracing tool (Figure 3B), and press Ctrl + M (Figure 3C) to automatically quantify the lumen area in pixels. Repeat this for each methacholine concentration; the data are numerically available for graphical representation with airway reactivity expressed as a fraction of baseline lumen size (Figure 3).
   3. Sample size
      1. Choose airways at random, and perform the response to methacholine on one AW/lung section, although typically multiple sections per animal can be assessed. Thus, treatment groups can consist of multiple airways/animals from multiple litters.
      2. Categorize airways into large (>0.05 mm²) or small (<0.05 mm²), since it has been previously determined that CPAP affects airways of different sizes.

10. Drug delivery

1. Analyze using sections exposed to (or incubated with) siRNA's targeting specific pathways for 48 h after being excised and sectioned.
   NOTE: Agonists and antagonists specific for various receptors can also conveniently be applied to the lung sections acutely, which is an advantage if such compounds are not appropriate or safe for in vivo administration.
2. Alternatively, lungs can be Inflation fixed (formalin) at standardized inflation pressures (25 cmH₂O), preserved, and prepare for immunohistochemistry and staining for later analysis using confocal microscopy.

Results

MRI analysis of lung volume during neonatal CPAP
MRI analysis was used to visualize changes in lung volume in 3-day-old, un-anesthetized, spontaneously breathing mice (n = 4 mice). Lung volume increased with increasing inflation pressures (i.e., CPAP) up to 9 cmH₂O (Figure 4A). Representative images of a 3-day-old mouse at 0 and 6 cmH₂O CPAP are also shown (Figure 4B,C). The purpose of this analysis was...

Discussion

CPAP is a necessary respiratory support modality for preterm infants with respiratory distress. CPAP is superimposed on spontaneous breathing and serves to improve systemic oxygenation by maintaining lung volume elevated and prevents alveolar collapse and atelectasis. Thus, there are major life-saving benefits to CPAP, and it has been shown to improve alveolar development, although the benefits may depend on the severity and duration of CPAP^6,18,

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
5 mL syringe	Fisher Scientific	149556458
1 mL syringe	Fisher Scientific	309659	To inject agarose through cannula
1/4" hole punch	McMaster Carr	3427A57
22 x 22 x 20 embedding molds	Fisher Scientific	22-19
23 G needles	Fisher Scientific	305193	To attach cannula to syringe, other sizes would be required for ages other than P21
2 mL serological pipettes	Fisher Scientific	13-678-12c	For making moanometer
Adjustable leak valves	any pet store		https://www.petco.com/shop/ en/petcostore/product/imagi tarium-air-control-valve
Aquarium tubing	any pet store		https://www.petsmart.com/fish/ filters-and-pumps/air-and-water -pumps/top-fin-aquarium-airline -tubing-5291863.html
Camera	Amscope	MU300
Cannula tubing	Fisher Scientific	427411	For P21 mice. Other sizes will be needed for other ages.  Tubing should fit snugly in trachea
Coverslips	Fisher Scientific	12-541-014	18 mm x 18 mm
DMEM:F12	Invitrogen	11039-021
Flow controllers	Key instruments	GS10510AVB	For airflow to individual mice
Flow meter	Avantor	MFLX68560-04	For airflow to the entire system
Gaymar T Pump	Braintree Scientific	TP-700	For mainaing body temperature during CPAP
Hanks balanced salt solution	Fisher Scientific	MT-20023cv	Dilute to 1x for use
ImageJ	NIH	free download	https://imagej.net/ij/download.html
Latex gloves size small	Fisher Scientific	mf-300-s	For making CPAP masks
Low metling point agarose	Invitrogen	165200-100
Mesh	Amazon		https://www.amazon.com/Top- Trimming-Polyester-Horsehair -selling/dp/B01AC93LJ0/ref=sr _1_fkmr2_1?keywords=dritz%2 Bpolyester%2Bhorsehair%2Bbr aid&qid=1562072725&s=gatew ay&sr=8-1-fkmr2&th=1
Methacholine	Millipore	A2251
Microscope	Leica	DMLFS
Pen/strep	Invitrogen	15140-148
Perfusion pump	Harvard Apparatus	70-2027
Recording chamber	Warner instruments	RC-27	Or similar
Reusable heating pad	Braintree Scientific	TP-R	For use with gaymar T pump
Silicone grease (Molykote111)	Amazon		https://www.amazon.com/Molykote -Lubricant-Components-Resistant- Volatility/dp/B0DYDFLXGZ/ref=sr_1 _1_sspa?crid=2D4LQC7SRX07P& dib=eyJ2IjoiMSJ9.HcWWXlt8ThxcB eGCZ6iH5jwumyClyFGkW141klcjnn OH3hc60r0hy53Ji-9HDW0TtH33nJ- 7UmalEzJGg59qB2gzsGNzFMelwG aBpsTElSrogzwQv27amBxERmXW P3eedvi9rAYS1oy_HNesZTAxjTbqku 6PxkC7t24srDUq8mHRpDFor4I0iiRM j6Ij2TWdlQf07ccTIMJ1l4n90nHxPop 9Nwh77ufMeWY2vu5B3to.0mfWIF 3_PrmciPZhO1EAE6RGiNj16v7l-v CH1cqec1Y&dib_tag=se&keyword s=molykote+111&qid=1743607899 &sprefix=molykote+111%2Caps%2 C102&sr=8-1-spons&sp_csd=d2lkZ 2V0TmFtZT1zcF9hdGY&psc=1
Sterile 6 well plates	Fisher Scientific	3516	For culturing slices
Suture thread 3-0	Fisher Scientific	sp117	To tie cannula into trachea
Tubing fittings	Harvard Apparatus	72-1406
Vibratome (pelco easislicer)	Ted Pella	11000	Or equivalent