Design and Implementation of a Rat Ex Vivo Lung Perfusion Model

Summary

Ex vivo lungs are useful for a variety of experiments to collect physiological data while excluding the confounding variables of in vivo experiments. Commercial setups are often expensive and limited in the types of data they can collect. We describe a method for building a fully modular setup, adaptable for various study designs.

Abstract

Ex vivo lung preparations are a useful model that can be translated to many different fields of research, complementing corresponding in vivo and in vitro models. Laboratories wishing to use isolated lungs need to be aware of important steps and inherent challenges to establish a setup that is affordable, reliable, and that can be easily adapted to fit the topic of interest. This paper describes a DIY (do it yourself) model for ex vivo rat lung ventilation and perfusion to study drug and gas effects on pulmonary vascular tone, independent of changes in cardiac output. Creating this model includes a) the design and construction of the apparatus, and b) the lung isolation procedure. This model results in a setup that is more cost-effective than commercial alternatives and yet modular enough to adapt to changes in specific research questions. Various obstacles had to be resolved to ensure a consistent model that is capable of being used for a variety of different research topics. Once established, this model has proven to be highly adaptable to different questions and can easily be altered for different fields of study.

Introduction

Ex vivo lung perfusion (EVLP) techniques¹ have seen a rise in usage in the past decade as a means of studying lung transplantations², ischemia/reperfusion³, lung metabolism⁴, and immune responses⁵. Isolated, but intact, ventilated and perfused lungs offer the critically important ability to directly assess the response of the lungs, including the pulmonary vasculature, to potential interventions and/or therapeutics without potential confounders, such as neuronal and hormonal input or changing hemodynamics in vivo. At the same time, they maintain the physiological interplay of ventilation and perfusion, in contrast to in vitro conditions. A proposal looking at immune responses in lungs⁵, for example, needs the same quality of data as a study focused on increasing the donor pool size⁶ for lung transplantations. EVLP can be used across a variety of species, including mice³, rats^{7,8,9,10,11,12}, pigs¹³, and humans². Therefore, it is necessary to establish a model that can produce reliable data from a variety of different experimental parameters. Clinical relevance will be generated in subsequent studies using the EVLP model as a tool.

While commercial setups are available for purchase for most species, they can often be cost-prohibitive and confine researchers to a specific brand of equipment and proprietary software. Any deviation from the out-of-the-box setup (e.g., going from one species to another) requires foresight and working around the provided setup, which may prove to be difficult or impossible. In the following, a DIY (do it yourself) setup for rat isolated lungs that is both modular and cost-effective, as well as the surgical procedure for isolating the lungs, are described.

Protocol

The in vivo portion of the experiments (from general anesthesia to euthanasia) requires prior approval by the respective Institutional Animal Care and Use Committee (IACUC). All procedures described herein were approved (protocol number M1700168) by the IACUC at Vanderbilt University Medical Center, Nashville, Tennessee, and were performed in compliance with the ARRIVE guidelines¹⁴. Prior to experimentation, all the rats were housed in the institute's animal care facility, with free access to water and food. Including different studies outside the purview of this manuscript, we have used a total of 148 male Sprague Dawley rats, 7-10 weeks old, with a weight between 250 g and 400 g so far.

1. Apparatus construction

NOTE: All parts, including respective manufacturers, are listed in the Table of Materials.

1. To hold each piece of equipment in place, construct a custom-built lattice to allow for easy configuration and incorporation of the new devices (Figure 1). Attach aluminum poles (1 to 2 ft in length, 1 cm in diameter, available at any hardware store) to each other using cross clamps to create a 3D lattice, and place on a plastic tray (30 in x 21 in) to prevent spillage of fluids.
2. Mount pressure transducers at an equal height to the lungs and connect them to the lines for the pulmonary artery (PA), pulmonary vein (PV), and to the ventilation tube.
   NOTE: These transducers transmit data to their respective bioamplifiers connected to the data acquisition system (DAQ) and its software.
3. Rather than using commercially available cannulas, craft custom cannulas (Figure 2) from segments of 2 mm wide, hard plastic tubing that are flared at the end, using an open flame to allow for the secure tying-off of sutures. Bend them into a U-shape to reduce stress on the lungs during hanging and to fit in the lung chamber.
4. To make the chamber the lungs are to be ventilated and perfused in, place a 1,000 mL beaker inside a 1,500 mL beaker with a water bath between the two and inside the 1,000 mL beaker, creating a double boiler. Place these beakers on a heating plate set to 48 °C, creating a chamber for the lungs that is both humid and resistant to fluctuations in temperature.
5. Keep the buffer for the experiment in a 150 mL volumetric flask placed on a heating plate set to 37 °C. Use a magnetic stirring bar to circulate the buffer inside the beaker. Set the meniscus of the buffer at a height such that it is 4 cm above the lung, to create an innate 4 cmH₂O of pressure on the PV. During the surgery, ensure that the animal's lungs are at the same height as the buffer to reduce hydrostatic edema formation.
   NOTE: Using a flask minimizes the amount of surface area in contact with room air, which minimizes gas diffusion.
6. Use a roller pump to move the buffer through a circuit consisting of a heating coil and an air trap prior to perfusing the lung. Recycle the effluent from the PV back into the volumetric flask. Connect both the heating coil and air trap to a circulating water bath also set to 37 °C. Adjust the temperature of the water bath according to the pump speed, so the perfusate has a constant temperature of 37 °C.

2. Procedure

1. Prior to the start of the experiment, prepare the setup.
   1. Ensure that the software is running (see below) and properly collecting data.
   2. Calibrate all pressure transducers daily to ensure they are not drifting.
   3. Prepare the buffer (see Table 1 for ingredients for a physiological saline solution with 4% bovine serum albumin [BSA]) and ensure that the pH is at 7.4, using HCl to adjust accordingly. Set up the perfusion system with warmed buffer (Table 1) circulating throughout, ensuring that no air bubbles are present. Add BSA at least 30 min prior to the start of the experiment to give it sufficient time to dissolve. In the absence of an oxygenator, bubble gas into the perfusion buffer prior to the addition of BSA with a gas composition of 65% N₂, 30% O₂, and 5% CO₂ to mimic the CO₂ levels of in vivo lungs. This prevents the solution from foaming once BSA is added.
   4. Prepare the operating area with all necessary surgical tools, sutures, and tape. Incline the operating board at a 15° angle, so that the anesthetized rat can be positioned with its head elevated above the rest of the body, and the trachea and heart-lung block can be manipulated easily.
   5. Wearing proper personal protective equipment, weigh the rat and give an intraperitoneal injection of pentobarbital (65 mg/kg^-1). After 10 min, use a toe pinch to ensure that the surgical plane of general anesthesia has been reached. Administer more anesthetic if necessary.
2. Transfer the rat to the operating area and fixate it to the operating board by taping its front legs separately, followed by the hind legs together, making sure that the front legs are taped loose enough to allow a pain reflex to be visualized should anesthesia not be deep enough (Figure 3A).
3. Secure the head of the rat by taping the mouth with a long, thin strip behind the front teeth, without restricting the tongue or airflow for the still spontaneously breathing rat (Figure 3B).
4. Perform a tracheostomy by pinching the skin above the trachea using forceps and cutting with surgical scissors. Using curved forceps, bluntly dissect the muscle and tissue to reach the trachea. Ensure that there is no bleeding during this step.
   1. Pass curved forceps underneath the trachea and open them to allow room to pass 3-0 sutures underneath that can then be pre-tied into a box knot (Figure 3B). Make a small incision between the cartilage rings of the trachea and insert the tracheal cannula (modified 18 G needle with 1 mm diameter tubing glued around the tip to create a notch). Tie off the suture, ensuring that no air can escape from the tracheal incision and that the cannula is not putting any strain on the trachea.
5. Set the ventilator to run with a gas mixture of 30% O₂, 5% CO₂, and 65% N₂, with a tidal volume (VT) of 10 mL/kg, a positive end expiratory pressure (PEEP) of 0 cmH₂O, and a rate of 60 breaths/min.
6. Once the tracheal cannula is secured with the suture, begin ventilating the rat with the above settings.
   NOTE: The surgical method used was adapted from Nelson et al.⁹ and is described step-by-step.
7. Remove the fur from the abdomen of the rat using large surgical scissors and forceps.
   NOTE: Hair removal ointments can also be used prior to the start of the experiment.
8. While holding the xiphoid process with forceps, make a small horizontal incision below the ribs, ensuring that the diaphragm stays intact. Once the internal organs can safely be visualized, widen the horizontal cut to expose the entire diaphragm.
9. Taking extreme care to avoid puncturing the lungs, inject heparin (3,000 U/kg^-1) into the inferior vena cava (IVC) with a 22 G syringe.
10. Again, hold the xiphoid process with forceps and cut cranially along the sternum while constantly visualizing the lungs to prevent cutting them.
11. To properly see the heart-lung block, spread the rib cage apart using two large forceps and be sure to avoid breaking the ribs, which can result in a sharp bone fragment puncturing the lung (Figure 3C).
    NOTE: Once the rib cage has been opened, use a PEEP of 2-3 cmH₂O, set by a water lock attached to the expiratory limb of the ventilator, to avoid atelectasis.
12. At this time, cut the IVC to euthanize the rat by exsanguination. Carefully ensure that at least 1 min has passed since the injection of heparin (see step 2.9) to give it enough time to circulate and prevent microclots in the lungs.
13. When placing the perfusion cannulas, constantly check the pressures to make sure that no sudden changes occur.
    NOTE: A sudden spike in pressure while the pump is running at a constant speed indicates blockage formation, while a sudden decrease can indicate a leakage.
14. Trim any excess thymus to allow for easier visualization of the pulmonary vasculature. Locate the PA, pass small, curved forceps underneath, and again pass a 3-0 suture underneath and pre-tie into a box knot.
15. Make a small incision into the right ventricle of the heart and insert the PA cannula (made from 2 mm diameter plastic tubing flared at the end using an open flame to allow for sutures to be tied around), being gentle to avoid rupturing the adjacent artery. Secure the cannula using the suture and perfuse, starting at 1.5 mL/min^-1 (Figure 3D).
16. Immediately excise the apex of the heart to avoid a pressure buildup in the lungs. Using small, curved forceps, rupture the mitral valve and visually ensure that the forceps are able to enter the left atrium unrestricted.
17. Place a pre-tied 3-0 suture around the heart below the atrium (Figure 3E).
18. Insert the PV cannula (similar construction to the PA cannula) into the left atrium and ensure that buffer can flow out of it prior to tying off the suture.
    NOTE: It is critical that the PV cannula is placed past the mitral valve to ensure that it can be properly tied off, while not being placed too deep as to injure the PV or encumber flow (Figure 3E). Take extra care with tying the suture, as tying it too loose results in lost flow, while tying it too tight can damage the heart, weakening the placement of the cannula and potentially causing leakages.
19. Trim excess tissue between the thoracic cavity and the tracheostomy incision using blunt-tipped scissors to avoid damaging the trachea. Ensure that the entire trachea below the tracheal cannula and the entire heart-lung block are visible.
20. Remove the heart-lung block and trachea by holding the tracheal cannula and excising connective tissue behind the trachea with a pair of curved, blunt-tipped scissors, leaving the esophagus attached for extra structural support (Figure 3F).
21. Gradually increase the flow rate to a maximum flow of 40 mL/kg^-1/min^-1. Allow the first 50 mL of buffer to flow out of the system to remove any inflammatory cytokines that may be released (Figure 3G).
22. Gently move the isolated lungs to the double boiler chamber created with the two beakers. Make sure that the PA, PV, or tracheal cannula do not become twisted at any point while moving the lungs or hanging them.
    ​NOTE: Prevention of atelectasis during lung retrieval and connection to the setup is important, especially when EVLP is used for already damaged lungs or when interventions prior to EVLP are planned. This can be achieved by, for example, a small clip on the trachea after inspiration before transfer to the ventilator¹⁵.

3. Data acquisition

1. For data collection, use any commercially available analog-to-digital converter and DAQ system.
2. Pay attention that the sampling frequency is sufficient for the given purpose (200 Hz here) and that all relevant dependent parameters from bioamplifiers and other input devices are collected concurrently (airway pressure, PA, and PV pressure chosen for this protocol)¹⁶.
3. Record independent parameters (e.g., perfusion speed, ventilation rate, ventilated gas composition, buffer electrolyte composition, and lung weight).

Results

Following 10 min of stabilization and baseline readings, we randomized a first set of 10 male Sprague Dawley rats into five small groups: global no-flow ischemia for 5, 7.5, 8, 9, or 10 min (n = 2 per group) followed by reperfusion; these limited preliminary dose-finding experiments were conducted to identify the longest possible ischemia time to still allow sufficient ventilation and reperfusion before the eventual development of a precipitous and irreversible increase in airway pressure and edema formation. Importantly...

Discussion

More than 100 experiments have been successfully performed in our lab using this setup. The modular design of this customized setup gave great flexibility to potential changes in experimental requirements. While other setups utilize a deoxygenator¹⁸ to mimic constant oxygen consumption and CO₂ production by end organs, this simplified model did not employ this feature, due to the focus on studying the effects of different gas compositions on pulmonary vascular tone. This approach, in wh...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
1,000 mL Glass Beaker	Pyrex, Chicago, IL
1,500 mL Glass Beaker	Pyrex, Chicago, IL
Air Trap Compliance Chamber	Radnoti	130149
Bioamplifiers	CWE Inc	BPM-832
Clamps	Fisher Scientific	S02626
DAQ (Data Acquisition)	National Instruments, Austin, TX	NI USB-6343
Gas Mixer	CWE Inc, Ardmore, PA	GSM-4
Heating Coil	Radnoti, Covina, CA	158822
Heating Plate	Thermo Fisher Scientific, Waltham, MA	11-100-49SH
Heparin	Pfizer	W63422
LabVIEW Full Development System 2014	National Instruments
Pentobarbital	Diamondback Drugs	G2270-0235-50
pH700 Probe	OAKTON, Vernon Hills, IL	EW-35419-10
Polystat Water Bath	Cole-Parmer	EW-12121-02
Rodent Ventilator	Harvard Apparatus, Holliston, MA	Model 683
Roller Pump	Cole-Parmer, Wertheim, Germany	Ismatec REGLO Digital MS 2/8
Sprague Dawley Rat	Charles River, Wilmington, MA	Strain code 001
VetScan i-STAT	Abraxis, Chicago, IL	i-STAT 1