The establishment of an ex vivo lung perfusion rat model: insights from Jiangxi, China

Introduction

Ex vivo lung perfusion (EVLP) is a technique that separates the lungs from the body and preserves, transports, reassesses, and repairs them through circulation perfusion and mechanical ventilation outside the body (1). This technique is mainly used in the field of lung transplantation (LTx), and the extensive research has been conducted on expanding the donor pool, improving the quality of marginal donor lungs, and creating more opportunities for patients on the transplantation waiting list to receive transplant surgery and obtain treatment (2). As the patients on the waiting list for transplantation are all patients with terminal lung diseases, LTx surgery represents the most effective treatment method of treatment and the best chance for survival. Therefore, the EVLP has its unique significance in promoting the development and progress of LTx (3). EVLP is not only a research frontier in the field of international LTx but also a technological frontier in China that urgently needs to be explored (4).

Over the past decade, significant progress in EVLP research has been made by researchers such as Steen et al. (3) in Sweden and Cypel et al. in Canada (5). Most of their landmark work was completed using human lungs or large animal models such as porcine lungs (6-8). In 2010, Okamoto et al. from Kyoto University in Japan, reported a porcine EVLP model with an average weight of 115 kg (9). However, due to the high cost of large-animal research and the scarcity of relevant antibody reagents for large animals such as pigs, the implementation of EVLP experiments on large animals is limited.

In 2014, Noda et al. established a reproducible rat EVLP model, successfully extending the evaluation duration of the isolated lung under EVLP (10). In 2015, Stone et al. used a mouse isolated lung perfusion system (Hugo Sachs Elektronik, March, Germany) to establish a C57BL/6 mice EVLP model and study “Ex Vivo Perfusion With Adenosine A2A Receptor Agonist Enhances Rehabilitation of Murine Donor Lungs After Circulatory Death” (11). In 2017, Noda et al. conducted research on a rat EVLP platform, finding that a perfusion fluid with an oxygen concentration of 40% caused the least lung injury compared to other concentration groups in the experimental setup (12). A hypoxic concentration of 6% perfusion fluid was previously the preferred ratio.

The use of small animals is critical to establishing reliable ex vivo perfusion systems for other parenchymatous organs such as the kidney and liver (13-20). Valuable EVLP models based on small animal such as rats, and the methodology by which they were produced, have also been reported in the past decade (21-25). Many rodent EVLP systems have been used for pharmacokinetic analysis or to simulate the LTx of vitro models (26).

In China, research into the field of small animal EVLP is rare, with the few existing studies being based on “do it yourself” (DIY) small-animal EVLP equipment or being carried out by Chinese researchers in well-equipped centers abroad (27,28). As small-animal EVLP technology remains in its infancy in China, Chinese researchers should increase their engagement with pioneering work in this field.

In Jiangxi, our team has independently mastered certain experimental procedures in establishing rat EVLP models. The main purpose of this paper is to summarize our team’s experience in the implementation of rat EVLP experiment so as to provide reference for other engaged in this research. The potential of using small-animal EVLP platforms to conduct basic research related to LTx is considerable. Establishing a stable and reproducible rat EVLP model that can extend the EVLP duration to more than 3 hours is challenging due to the numerous variables involved in the creation of the EVLP rat model. Controlling these variables within an appropriate range is a significant obstacle to achieving this goal. This article will modularly introduce how to conduct the related work in establishing the rat model and outline the experimental venues, experimental reagents, experimental equipment, instruments, consumables, experimental procedures, and data collection and organization, among other subjects. We present this article in accordance with the SUPER and ARRIVE reporting checklists (available at https://jtd.amegroups.com/article/view/10.21037/jtd-24-1754/rc).

Pre-experimental preparations and requirements

Preparation of experimental sites

Site for preparation of lung preservation solution/perfusion solution

The site for preparation of lung preservation solution/perfusion solution should be dry, ventilated, and temperature-controlled. A special reagent storage cabinet for light protection is needed, there should be a countertop for the precise weighing and dissolving of reagents, deionized water should be easily accessible, and an ultrapure table is needed for sterilizing and filtering the preservation/perfusion solution.

Site for the EVLP machine countertop

The site for machine countertop should be dry, ventilated, and temperature-controlled; with good lighting. Special gas cylinders can be placed next to the countertop, and a water source should be easily accessible for cleaning.

Site for anesthesia and the surgical microsurgical operation platform

The site for anesthesia and the surgical microsurgical operation platform should be dry, ventilated, and temperature-controlled. A special safe for storing hazardous chemicals such as anesthetics should be present, and a water source should be easily accessible for cleaning.

Site for operation room of paraffin and frozen pathological microtome

The site for paraffin and freezing procedures should be dry, ventilated, and temperature-controlled.

Site for sample storage

For sample storage, dedicated refrigerators and freezers are optimal, and environment at site should have good heat dissipation.

Site for sample testing

For sample testing, the site should be able to accommodate routine experiments such as wet-to-dry-weight ratio, blood gas analysis, osmolarity detection, inflammatory factor kit detection, and Western blotting protein detection.

Experimental team personnel

In the establishment of our experiment, the experimental staff consisted of surgeons with basic surgical knowledge who were mainly responsible for the surgical manipulation of the rats and the connection of the EVLP. Medical students with professional laboratory training were also included and were mainly responsible for the preparation of the relevant solutions, the anesthesia of the rats, the implementation of the subsequent experiments of the EVLP, and the collection and collation of the experimental data related to the EVLP.

Preparation of experimental reagents

The reagents required for EVLP are mentioned in Table 1. For pulmonary preservation solution and lung perfusate, we currently adopt the independent formulation based on the Perfadex solution (XVIVO, Gothenburg, Sweden) as published on the manufacturer’s website (https://www.xvivogroup.com/wp-content/uploads/2023/11/IFU-PERFADEX-Plus-US-Issued-2022.07.pdf) and Steen patent No. US7255983B2.

The following are the specific formulation details. A locally manufactured pulmonary preservation solution is formulated according to the specifications outlined in Table 2, while a locally produced lung perfusate is also prepared as stipulated in Table 3. Tris (0.408 g) can be used to titrate the locally manufactured pulmonary preservation solution to 7.3. If sodium hydroxide titration is used, this can be completed based on real-time pH monitoring.

The perfusate is filtered with a vacuum filtration device to remove any particulate impurities or bacterial contamination and stored for up to 5 days to avoid bacterial growth (24) (Figure 1).

Figure 1 The final step of preparing the perfusate. (A) The perfusate is filtered with a vacuum filtration device. (B) Preservation status of locally manufactured lung perfusate.

Preparation of laboratory consumables

The laboratory consumables are outlined in Table 4.

Preparation of experimental equipment

Instruments for the EVLP experiment are outlined in Table 5.

Surgical instruments

The instruments associated with animal surgical operations are shown in Figures 1-6.

Figure 2 Rat tracheal intubation (for use after anesthesia).

Figure 3 Surgical instruments for thoracotomy and laparotomy in rats.

Figure 4 Orthopedic bandage gauze scissors.

Figure 5 Surgical instruments for microsurgery.

Figure 6 Tracheal intubation: (I) pulmonary artery intubation; (II) and left atrial intubation; (III) provided by IPL-2. IPL, Isolated Perfused Lung.

Animal preparation

All operations in our experiment were performed according to the guidelines for institutional animal care and use established by the Institutional Animal Care and Use Committee (IACUC) of Nanchang University, and the project was approved by the Ethics Committee of the First Affiliated Hospital of Nanchang University (ethical approval No. CDYFY-IACUC-202406QR025). Specific pathogen-free (SPF)-grade adult male Sprague Dawley rats, bought from Experimental Animal Science and Technology Center of Jiangxi University of Traditional Chinese Medicine, weighing between 200 and 320 g, were housed in a ventilated cage system maintained at 22±1 ℃ with 55%±5% humidity under a 12-hour dark-light cycle and had unrestricted access to standard rat chow feed and water. A protocol was prepared before the study without registration.

Description of the procedure

Surgical operations and EVLP loading

An open surgical procedure was performed on rats to simulate the steps of donor lung procurement surgery. The use of modularized operating procedures can avoid additional man-made damage, which can shorten the procurement time for donor lungs and reduce the duration of warm ischemia. The instructions for the surgical operation and EVLP loading are as follows:

• Wear personal protective equipment before handling rats and rat tissues, including surgical masks, surgical gloves, and disposable surgical gowns.
• Weigh the rat and record its weight.
• Prepare 500 U of heparin per 100 g of rat weight [different sources (21,22,25) have reported different amounts of heparin].
• Prepare 3.5–4 mg/100 g body weight of pentobarbital in a syringe for standby use. First, prepare a 1% pentobarbital solution by dissolving 120 mg in 12 mL of normal saline (NS).
• For the induction of anesthesia, place the rats in an isoflurane gas chamber, and after inhalation of the gas, administer sodium pentobarbital to the rats via intraperitoneal injection. After the injection, allow the rats to rest for 5 minutes to ensure that they are completely unconscious.
• Assess the anesthetic effectiveness by observing the paw withdrawal response to a toe pinch. The absence of paw withdrawal indicates the rat is not experiencing pain.
• Perform tracheal intubation in rats using indwelling pins with rigid inserts (Figure 2).
• Transfer the rat onto the operating table, secure it in a supine posture, and disinfect thoroughly with alcohol spray.
• Prepare silk suture ranging from 4 to 20 cm in length (0-0 or 2-0, 3-0, or 4-0).
• Prior to initiating the procedure, reconfirm the adequate depth of anesthesia. First, make an abdominal midline incision for entry into the abdominal cavity, sever the xiphoid process (Figure 7), and then incise the diaphragm to gain access to the thoracic cavity.
• Inject heparin (500 U heparin/100 g) into the inferior vena cava according to the rat’s body weight (Figure 8).
• Extend the incision upward to the neck until the trachea is exposed.
• Fully isolate the cervical segment of the trachea and pass a 0-0 silk suture thread behind the trachea.
• Lift the anterior part of the trachea and make an inverted T-shaped incision between the tracheal cartilage rings but do not transect the membranous part of the trachea posteriorly.
• Intubate with a metal tracheal cannula and secure it with silk suture ties and ensure that the sutures are tied in the grooves of the tracheal cannula. If necessary, cross the two suture ends behind the trachea and tie them in front to prevent displacement or detachment of the cannula.
• Connect the tracheal cannula to the ventilation circuit.
• Turn on the ventilator for small animals and begin mechanical ventilation (positive pressure ventilation).
• Cut along both sides of the sternum, upward to the sternoclavicular joint, and enter the thoracic cavity, taking care to avoid injury to the lungs (Figure 9). Moreover, avoid damaging the parasternal internal mammary artery and vein as well as the bilateral subclavian arteries and veins to prevent excessive bleeding.
• Use orthopedic bandage gauze scissors (Figure 3) to cut along the midline toward the neck to fully expose the anterior trachea.
• Carefully isolate the posterior aspect of the main pulmonary artery and ascending aorta, and pass 2-0 or 3-0 silk suture threads behind the main pulmonary artery and aorta to preset the ligation lines.
• Use 0-0 silk suture threads to preset ligation lines horizontally at the atrioventricular groove posterior to the heart.
• Cut open the inferior vena cava for bloodletting, place absorbent paper in the abdominal cavity to absorb the blood, and prevent the blood from flowing onto the operating table.
• Make a horizontal incision of approximately 1.5 mm in length on the right ventricular outflow tract, insert a pulmonary artery cannula, tie the preset suture line securely, and connect the distal end of the cannula to the pulmonary preservation fluid for pulmonary perfusion.
• Cut open the apex of the heart, retrogradely insert a pulmonary vein cannula through the mitral valve, place the tip in the left atrium, and leave the distal end open.
• Begin pulmonary perfusion (perfusion pressure 20 cmH₂O) while observing whether there is perfusion fluid flowing out of the pulmonary vein cannula and whether the pulmonary artery cannula is inserted too deeply. Avoid poor perfusion on one side of the lung to ensure good perfusion of both lungs.
• Stop perfusion when the lungs turn milky white and disconnect the distal perfusion catheter of the pulmonary artery.
• Carefully isolate the posterior mediastinum to the cervical trachea, transect the membranous part above the tracheal cannula, transect the descending aorta, and remove the heart-lung tissue block.
• Connect a syringe to the tracheal cannula, inflate the lungs, and close the trachea to maintain the lungs in a semi-inflated state.
• Place the semi-inflated donor lungs in a 4 ℃ environment or other experimental settings for a specific duration (set according to experimental needs).
• Prefill the perfusion fluid in the EVLP short-circuit circulation.
• To hang the lungs, connect the tracheal cannula to the ventilation port, connect the pulmonary artery cannula to the arterial perfusion port, and leave the pulmonary vein cannula open or connect it to the pulmonary vein connection port.

Figure 7 Separation of the xiphoid process.

Figure 8 Injection of heparin into the inferior vena cava with a 1-mL syringe.

Figure 9 The sternum and part of the ribs on either side are lifted cephalad.

EVLP running

The rat EVLP platform used in our center is the Isolated Perfused Lung (IPL-2) System for Rat and Guinea Pig (Harvard Apparatus, Holliston, MA, USA; Hugo Sachs Elektronik, March, Germany) (Figure 10).

Figure 10 Panoramic image of the Isolated Perfused Lung System for Rat and Guinea Pig (IPL-2; Harvard Apparatus, Holliston, MA, USA; Hugo Sachs Elektronik, March, Germany).

The instructions for EVLP running are as follows:

• Open the EVLP data acquisition system and the software on the computer to begin using the data acquisition system to record the data, including ventilation data, and to monitor pressure.
• Ensure that the perfusion fluid storage reservoirs I and II are filled with sufficient perfusion fluid to prevent air bubbles from entering the pulmonary artery and forming an air embolism (Figure 11). To prevent excessive bubbles from forming in the perfusion fluid storage reservoir II, connect a bubble trap to the outlet of the exchange membrane to reduce the number of bubbles entering the perfusion fluid storage (as shown in the Figure 12).
• Slowly adjust the rewarming, ventilation, and perfusion settings to the desired levels for the experiment within the initial period (set according to the experimental protocol). During this time, gradually increase the perfusion flow rate to the desired rate and/or pressure. Optimally, use a ventilator with a sigh function and set it to intermittent sigh breathing to help expel fluid from the lung space and delay the occurrence of edema.
• Obtain a sample from the sample port, quickly freeze it with liquid nitrogen, and record the sample time.
• After the experiment, retain part of the lung tissue and quickly freeze it with liquid nitrogen or place it in a fixative for further research, such as pathological analysis, detection of inflammatory factors, or other related proteins after grinding.

Figure 11 Reservoir I (A) and reservoir II (B).

Figure 12 Large number of air bubbles in the perfusion solution after deoxygenation.

Collection and organization of experimental data

The experimental data recorded in chronological order are as follows:

• Osmolarity of lung preservation solution and lung perfusate [before filtration at 4 ℃: 378 mOsm/(kg·H₂O); after filtration at 4 ℃: 374 mOsm/(kg·H₂O); before filtration at room temperature: 367 mOsm/(kg·H₂O); after filtration at room temperature: 334 mOsm/(kg·H₂O)].
• A pH value of lung preservation solution and lung perfusate (before calibration) of 6.4, consistent with the pH of the commercially available Perfadex solution.
• Weight of the rats.
• Original weight of the rat cardiopulmonary tissue block and weight after EVLP.
• Wet-to-dry-weight ratio.
• Several indicators at different time points during EVLP, including airway pressure, pulmonary artery pressure (PAP), pulmonary vascular resistance (PVR), pH values at the pulmonary artery and pulmonary vein ends, and blood gas analysis at the pulmonary artery and pulmonary vein ends.
• Results of inflammatory factors.
• Collection and analysis of pathological slice images.

Postexperimental considerations and tasks

EVLP shutdown and equipment cleanup

The cleaning of the EVLP circulation pipeline after the experiment is completed is a critical step.

The instructions for EVLP shutdown and equipment cleanup are as follows:

• After shutting down the ventilator, pause the peristaltic pump, remove the cardiopulmonary tissue block, and use a short connecting tube to connect the pulmonary artery and pulmonary vein intubation to form a short circuit.
• Quickly add 500 mL of deionized water to the perfusion fluid storage reservoir and start the peristaltic pump at 50 mL/min, and begin perfusion.
• Collect and discard the perfusion fluid from the recovery end directly. During the entire EVLP operation, avoid spillage of the perfusion fluid on the table or other places, as the perfusion fluid contains high-concentration dextran, which is highly viscous and difficult to clean if spilled on the table. If the perfusion fluid is accidentally spilled, wiped it clean with a damp cloth immediately to avoid the formation of stubborn stains over time.
• Clean and wipe the tabletop, avoiding the use of solvents such as alcohol that may corrode plexiglass.
• Routinely clean, dry, disinfect, and return other items to their proper places. Power off the equipment, and properly dustproof the data acquisition system and computer.

Experimental data analysis

The acquisition of research parameters is a complex task and includes the management of time and space. In order to properly record and analyze the overall experimental data, it is critical to become proficient at time and space management. The instructions for data analysis are as follows:

• Measured the infusate osmolarity at different time points according to experimental needs, which include the immediate value after preparation, the value after filtration, the value at the pulmonary venous end, the value at the pulmonary venous end after different durations of EVLP, and the values at the pulmonary arterial end and pulmonary venous end at different time points of EVLP, such as at half an hour or one hour under the scheme of pulmonary venous infusion recovery.
• For inflammatory factor data analysis, collect pulmonary venous infusion fluid at different EVLP time points for detection and analysis and then conduct horizontal and vertical comparative analysis of the results.
• For the PAP value, collect PAP values at different EVLP time points and analyze them and then conduct horizontal and vertical comparative analysis of the results.
• For the pulmonary venous pressure (PVP) value, collect PVP values at different EVLP time points and analyze them; then conduct horizontal and vertical comparative analysis of the results.
• For blood gas analysis results, collect pulmonary venous blood at different EVLP time points for blood gas analysis and conduct horizontal and vertical comparative analysis of the results. In addition, there is a real-time oxygen partial pressure probe on the venous side of the isolated lung for data recording and continuous monitoring. The pulmonary venous oxygen partial pressure at different EVLP time points can be analyzed later; there is only a need to check whether its function is normal at the beginning of EVLP operation.
• There are real-time pH probes on both the arterial and venous sides of the isolated lung for data recording and continuous monitoring. The pH values at different EVLP time points can be analyzed later; there is only a need to verify the function of the pH probe at the beginning of EVLP operation.

Tips and pearls

According to our experience, it is best for beginners who have just begun using EVLP on the IPL-2 platform to be competent at the following steps before meaningful research can be conducted:

• Ensure accurate donor heparinization and that the heparin enters the blood when injecting heparin into the inferior vena cava.
• Perform surgical operations gently and carefully, especially during thoracotomy, to avoid causing damage or contusion to the donor lung; avoid causing massive hemorrhage that leads to premature heart stoppage and prolonged thermal ischemia.
• Complete pulmonary artery intubation under the condition that the heart is still beating. After the pulmonary artery intubation is completed and fixed, quickly cut the superior and inferior vena cava to exsanguinate the animal, immediately cut open the apex of the heart to open the left ventricle, and then immediately flush the lungs with lung preservation fluid to shorten the thermal ischemia time of the donor lung.
• When flushing the donor lung with lung preservation fluid, ensure even perfusion of both lungs to avoid blood residue.
• Adopt a perfusion fluid with high osmolarity, preferably greater than 300 mOsm/(kg·H₂O).
• During the cold ischemic preservation period after the donor lung is obtained, it is necessary to maintain the lung in a semi-inflated state.
• Prevent the donor lung from undergoing cold ischemia for an extended period before starting EVLP and control it within 1 hour.
• Employ continuous fresh perfusion fluid and do not recycle or reuse the perfusion fluid at the venous end.
• Adopt a preservation ventilation strategy: beginners can start with positive pressure ventilation and attempt negative pressure ventilation strategies after gaining proficiency.
• Adopt a preservation perfusion flow rate strategy.

If the abovementioned measures are implemented, acute pulmonary edema is less likely to occur during the EVLP process, and the duration of EVLP extend past 1 hour, which will greatly enhance beginners’ confidence in continuing the experiment. Otherwise, if pulmonary edema occurs in a short time, with a large increase in lung water in the trachea, the experiment cannot be carried out. This results in a waste of experimental resources and frustration to the experimenter, which may even cause them to lose confidence in continuing the experiment.

Discussion

Currently, there are two forms of rat EVLP platforms: self-assembled and commercialized. Both have roughly the same basic structure, including a small-animal ventilator (ventilate the isolated lung), a centrifugal pump or roller pump (provide perfusion power to the isolated lung), a heater or cooler (adjust the EVLP temperature), tubing (the path through which the perfusion travels), a rigid fluid reservoir (perfusate storage container) and a lung chamber (in which the lung is placed within a specially designed plastic or glass chamber to maintain its stability in a fixed position during ventilation, and to ensure a warm, humid environment). There are also rat EVLP devices containing a hollow fiber oxygenator, a leukocyte filter, an inline gas analyzer, a saturation probe, and a pressure sensor, weight sensor, data acquisition system atomizing drug feeder, micro-pump drug feeder, etc.

Leveraging the small animal EVLP model, a multitude of fundamental research avenues can be pursued. These include investigations into lung preservation solutions and lung perfusion solutions, studies pertaining to pharmaceuticals, studies on the mode of administration, explorations of EVLP perfusion and ventilation parameters, as well as other operational conditions, research into anti-infection strategies, studies combining immune rejection with rat LTx models, research on stem cell repair, gene therapy-related investigations, research into lung injury, and EVLP hardware and software related research, among others.

The clinically used lung transplant preservation solutions that are intracellular fluids include the Euro-Collins solution (EC solution) and University of Wisconsin solution (UW solution); meanwhile, those that are extracellular fluids include compound sodium chloride solution, lactated Ringer’s albumin solution (Hartmann’s solution), Collins solution and its four modified solutions (C1, C2, C3, C4), low-potassium dextran (LPD) solution, Celsior solution, histidine-tryptophan-ketoglutarate solution (HTK solution), extracellular-type trehalose-containing Kyoto solution (ET-Kyoto solution), Perfadex solution, extracellular phosphate-buffered solution (EP-TU solution), Papworth solution, and Plegisol solution, among others.

The success of EVLP is significantly influenced by the type of perfusate utilized, with acellular and cellular perfusates being the two primary options, and both have distinct advantages and disadvantages (29). Acellular perfusates, such as Steen’s solution, contain no blood components and mimic the extracellular environment by providing essential electrolytes, nutrients and antioxidants. The advantages of this perfusate are its simplicity, lower cost and ability to reduce the risks associated with blood transfusions (30). However, it may not fully mimic the oxygenation and metabolic environment in vivo, which can be a limiting factor when assessing donor lung function (31). In contrast to the acellular perfusate, the cell-containing perfusate incorporates washed red blood cells and specific concentrations are added in accordance with the requirements of EVLP and resuspended. By emulating the composition of blood in the body, this perfusion solution provides the donor lung with a gas-blood exchange environment that is more closely aligned with normal physiological conditions (32), facilitating the maintenance of oxygen supply to the lung tissue throughout the perfusion process.

Some components of several preservation/perfusion solutions are listed in Table 6. High concentrations of K⁺ and Mg²⁺ can prevent the loss of K⁺ and accumulation of Na⁺ in cells of ex vivo organs, maintaining the resting potential of cell membranes. Mg²⁺ can antagonize the contractile effects of Ca²⁺, dilate blood vessels, and also antagonize the entry of calcium ions into cells, preventing intracellular calcium overload. Among the solutions available, Collins solution has a simple composition and low price, but it can produce magnesium phosphate precipitation and is high in potassium content, which can significantly damage vascular endothelial cells. EC solution does not contain Mg²⁺ and does not produce crystals, solving the problem of blockage of small blood vessels in the lungs, but it still has the issue of damage to cells from high potassium content and the use of glucose to maintain osmotic pressure, which can lead to significant acidosis in tissues. UW uses lactose and xylose to replace chlorides in the modified protective solution, obviating the cellular edema and intracellular acidosis caused by cellular hypoxia. Allopurinol is used to prevent the release of oxygen-derived free radicals mediated by xanthine oxidase, glutathione serves as a reducing agent, and adenosine serves as a precursor for the synthesis of adenine nucleotides (33); LPD and Perfadex are more effective than UW. In theory, the use of hyperosmotic protective solutions containing colloids can prevent the occurrence of pulmonary edema. However, in reality, due to factors such as increased capillary construction, lung vascular compliance, and extravasation of perfusion fluid, macromolecules may enter the tissues over time, causing pulmonary edema (34). It has been reported that Perfadex, Papworth, and EC do not differ significantly in terms of their effect on LTx (35).

Except for some studies on lung protective fluid and lung perfusion solution, many other studies on EVLP in rats used Perfadex and Steen solution as pulmonary protective fluid and pulmonary perfusion fluid.

The comprehensive evaluation index system of EVLP for donor lungs includes the following: PAP, PVR, peak inspiratory pressure (Ppeak), plateau airway pressure (Pplat), dynamic compliance (Cdyn), and static compliance (Csta) of the donor lungs, as well as the assessment of oxygenation capacity through blood gas analysis (including PaO₂/FiO₂ ratio, power of hydrogen, lactic acid level, hemoglobin level, glucose levels). The donor lung function is evaluated by measuring the difference in oxygen partial pressure before and after perfusion. Additionally, after EVLP bypass, chest X-ray examination is performed to observe the clarity of lung texture, measure changes in lung weight before and after perfusion, and assess inflammatory cytokines and cell count levels to clarify the status of lung inflammation and edema.

Ku et al. successfully delivered rituximab to donor lungs using the EVLP platform. Their experiments illustrated the potential of EVLP as a platform to deliver monoclonal antibody therapies to treat donor lungs pretransplant with the goal of eliminating a latent virus responsible for considerable morbidity after LTx (36). Lonati et al. tested the idea that treatment with the synthetic α-melanocyte-stimulating hormone analogue [Nle4,D-Phe7]-α-MSH (NDP-MSH) during EVLP could exert positive influences in lungs exposed to different injuries. Their findings suggest that NDP-MSH administration preserves lung function through broad positive effects on multiple pathways and suggest that exploitation of the melanocortin system during EVLP could improve reconditioning of marginal lungs before transplantation (37). Hasenauer et al. studied the effects of cold or warm ischemia and EVLP on the release of damage associated molecular patterns and inflammatory cytokines in experimental LTx. Their study found that EVLP significantly promoted an inflammatory response in both the cold (CI-E) and hot (WI-E) groups, which was not associated with cell death or damage-associated molecular pattern (DAMP) release at the end of EVLP, but with the release of S100A8 after LTx. EVLP mitigated graft damage and dysfunction in hot-ischemic lungs, but not in cold-ischemic lungs. The pathological mechanism of pulmonary aseptic inflammation is obviously dependent on conditions. The release of HMGB1 (without EVLP) and S100A8 (after EVLP) may be important factors in the pathogenesis of LTx (38). In other aspects, Amaia Ojanguren, etc., evaluated whether transient heat application during EVLP [thermal preconditioning (TP)] might recondition damaged lungs before LTx. They hypothesized that EVLP could also permit nonpharmacologic repair through the induction of a heat shock response, which confers stress adaptation via the expression of heat shock proteins (HSPs) (39). Arni et al. used K (ATP) channel modulators diazoxide or 5-hydroxydecanoic acid (5-HD) during EVLP experiments to protect against lung ischemia-reperfusion injury and inhibit the formation of reactive oxygen species (40). Wang et al. certificated rat lungs obtained after warm ischemia could be reconditioned during EVLP using the ROS/RNS scavenger Mn(III)-tetrakis (4-benzoic acid) porphyrin chloride (MnTBAP) or the PARP inhibitor 3-aminobenzamide (3-AB) (41).

The key to establishing rat EVLP is to become proficient in the above-described procedures and apply them under restricted conditions. According to relevant reports, most rat EVLP models run for 1 to 4 hours, with the longest lasting up to 6 hours (42). Once EVLP has been able to run successfully and stably for more than an hour, the above meaningful experiments in various fields can be carried out based on the model.

Conclusions

We have accumulated relevant experience on the rat EVLP platform IPL-2 and present it here to provide a reference and inspire fellow researchers in the hope of promoting technological progress and development in this field.

The operation of IPL-2 relies heavily on five core technical pillars: precise application of lung preservation solution and perfusion fluid; skilled and standardized experimental operation; scientific construction of the perfusion power system; rational setup of the ventilation system; and accurate execution of data collection, recording, and analysis. The synergistic effect of these technologies ensures the stable operation and excellent performance of IPL-2.

The modular and systematic conduct of EVLP-related procedures on rats has increased the success rate of experiments and greatly reduced costs, which is more conducive to the extensive development of basic research in the field of EVLP.

Since our team was one of the early ones to engage in research in this field domestically in China, many of the problems we faced were like crossing a river by feeling the stones. Our research achievements mentioned above might seem to our international peers as merely duplicating work and not making significant contributions, but these are very meaningful for us to further carry out in-depth related research.

In the future, we will continue to conduct in-depth research and exploration in this field, constantly deepening our understanding and knowledge in order to achieve additionally breakthroughs in this area. We will invest more time and energy to conduct experiments, with the aim of generating better solutions to the problems and challenges in this field. Moreover, we will engage more intensely in cooperation and exchanges with experts and scholars in China and abroad to promote the development and progress of this field. We believe that through continuous effort and collaboration, greater achievements in this field can be realized.

Acknowledgments

Funding: This study was funded by grants from Jiangxi Provincial Natural Science Foundation – Youth Project (No. 20202BABL216001) and the National Key R&D Program of China (No. 2023YFC2508604).

Footnote

Reporting Checklist: The authors have completed the SUPER and ARRIVE reporting checklists. Available at https://jtd.amegroups.com/article/view/10.21037/jtd-24-1754/rc

Peer Review File: Available at https://jtd.amegroups.com/article/view/10.21037/jtd-24-1754/prf

Conflicts of Interest: All authors have completed the ICMJE uniform disclosure form (available at https://jtd.amegroups.com/article/view/10.21037/jtd-24-1754/coif). The authors have no conflicts of interest to declare.

Ethical Statement: The authors are accountable for all aspects of the work in ensuring that questions related to the accuracy or integrity of any part of the work are appropriately investigated and resolved. All operations in our experiment were performed according to the guidelines for institutional animal care and use established by the Institutional Animal Care and Use Committee (IACUC) of Nanchang University, and the project was approved by the Ethics Committee of the First Affiliated Hospital of Nanchang University (ethical approval No. CDYFY-IACUC-202406QR025).

Open Access Statement: This is an Open Access article distributed in accordance with the Creative Commons Attribution-NonCommercial-NoDerivs 4.0 International License (CC BY-NC-ND 4.0), which permits the non-commercial replication and distribution of the article with the strict proviso that no changes or edits are made and the original work is properly cited (including links to both the formal publication through the relevant DOI and the license). See: https://creativecommons.org/licenses/by-nc-nd/4.0/.

References

1. Cypel M, Keshavjee S. The clinical potential of ex vivo lung perfusion. Expert Rev Respir Med 2012;6:27-35. [Crossref] [PubMed]
2. Cypel M, Yeung JC, Liu M, et al. Normothermic ex vivo lung perfusion in clinical lung transplantation. N Engl J Med 2011;364:1431-40. [Crossref] [PubMed]
3. Steen S, Sjöberg T, Pierre L, et al. Transplantation of lungs from a non-heart-beating donor. Lancet 2001;357:825-9. [Crossref] [PubMed]
4. Lin H. Current strategies of ex vivo lung perfusion as a platform for optimizing donor lungs. Chinese Journal of Organ Transplantation 2022;43:456-9.
5. Cypel M, Yeung JC, Hirayama S, et al. Technique for prolonged normothermic ex vivo lung perfusion. J Heart Lung Transplant 2008;27:1319-25. [Crossref] [PubMed]
6. Erasmus ME, Fernhout MH, Elstrodt JM, et al. Normothermic ex vivo lung perfusion of non-heart-beating donor lungs in pigs: from pretransplant function analysis towards a 6-h machine preservation. Transpl Int 2006;19:589-93. [Crossref] [PubMed]
7. Wiebe K, Poeling J, Meliss R, et al. Improved function of transgenic pig lungs in ex vivo lung perfusion with human blood. Transplant Proc 2001;33:773-4. [Crossref] [PubMed]
8. Okamoto T, Chen F, Zhang J, et al. Comparison of extracellular-type-Kyoto solution and Perfadex as a preservation solution in a pig ex vivo lung perfusion model: impact of potassium level. Transplant Proc 2011;43:1525-8. [Crossref] [PubMed]
9. Okamoto T, Chen F, Zhang J, et al. Establishment of an ex vivo lung perfusion model using non-heart-beating large pigs. Transplant Proc 2010;42:1598-601. [Crossref] [PubMed]
10. Noda K, Shigemura N, Tanaka Y, et al. Successful prolonged ex vivo lung perfusion for graft preservation in rats. Eur J Cardiothorac Surg 2014;45:e54-60. [Crossref] [PubMed]
11. Stone ML, Sharma AK, Mas VR, et al. Ex Vivo Perfusion With Adenosine A2A Receptor Agonist Enhances Rehabilitation of Murine Donor Lungs After Circulatory Death. Transplantation 2015;99:2494-503. [Crossref] [PubMed]
12. Noda K, Tane S, Haam SJ, et al. Optimal ex vivo lung perfusion techniques with oxygenated perfusate. J Heart Lung Transplant 2017;36:466-74. [Crossref] [PubMed]
13. Zhang P, Sun C, Mo S, et al. Salvaging donated kidneys from prolonged warm ischemia during ex vivo hypothermic oxygenated perfusion. Kidney Int 2024;106:273-90. [Crossref] [PubMed]
14. Muth V, Gassner JMGV, Moosburner S, et al. Ex Vivo Liver Machine Perfusion: Comprehensive Review of Common Animal Models. Tissue Eng Part B Rev 2023;29:10-27. [Crossref] [PubMed]
15. Bahena-Lopez JP, Rojas-Vega L, Chávez-Canales M, et al. Glucose/Fructose Delivery to the Distal Nephron Activates the Sodium-Chloride Cotransporter via the Calcium-Sensing Receptor. J Am Soc Nephrol 2023;34:55-72. [Crossref] [PubMed]
16. Valdivia E, Rother T, Yuzefovych Y, et al. Genetic Modification of Limbs Using Ex Vivo Machine Perfusion. Hum Gene Ther 2022;33:460-71. [Crossref] [PubMed]
17. Higashi Y, Homma J, Sekine H, et al. External pressure dynamics promote kidney viability and perfusate filtration during ex vivo kidney perfusion. Sci Rep 2022;12:21564. [Crossref] [PubMed]
18. Rigo F, Navarro-Tableros V, De Stefano N, et al. Ex Vivo Normothermic Hypoxic Rat Liver Perfusion Model: An Experimental Setting for Organ Recondition and Pharmacological Intervention. Methods Mol Biol 2021;2269:139-50. [Crossref] [PubMed]
19. Haque O, Pendexter CA, Cronin SEJ, et al. Twenty-four hour ex-vivo normothermic machine perfusion in rat livers. Technology 2020;8:27-36. (Singap World Sci). [Crossref] [PubMed]
20. Watanabe M, Okada T. Langendorff Perfusion Method as an Ex Vivo Model to Evaluate Heart Function in Rats. Methods Mol Biol 2018;1816:107-16. [Crossref] [PubMed]
21. Oliveira P, Yamanashi K, Wang A, et al. Establishment of an Ex Vivo Lung Perfusion Rat Model for Translational Insights in Lung Transplantation. J Vis Exp 2023;
22. Cleveland WJ, Hees JE, Balzer C, et al. Design and Implementation of a Rat Ex Vivo Lung Perfusion Model. J Vis Exp 2023; [Crossref]
23. Ohsumi A, Kanou T, Ali A, et al. A method for translational rat ex vivo lung perfusion experimentation. Am J Physiol Lung Cell Mol Physiol 2020;319:L61-70. [Crossref] [PubMed]
24. Bassani GA, Lonati C, Brambilla D, et al. Ex Vivo Lung Perfusion in the Rat: Detailed Procedure and Videos. PLoS One 2016;11:e0167898. [Crossref] [PubMed]
25. Nelson K, Bobba C, Eren E, et al. Method of isolated ex vivo lung perfusion in a rat model: lessons learned from developing a rat EVLP program. J Vis Exp 2015;52309. [Crossref] [PubMed]
26. Eriksson J, Sjögren E, Thörn H, et al. Pulmonary absorption - estimation of effective pulmonary permeability and tissue retention of ten drugs using an ex vivo rat model and computational analysis. Eur J Pharm Biopharm 2018;124:1-12. [Crossref] [PubMed]
27. Wang X, Parapanov R, Debonneville A, et al. Treatment with 3-aminobenzamide during ex vivo lung perfusion of damaged rat lungs reduces graft injury and dysfunction after transplantation. Am J Transplant 2020;20:967-76. [Crossref] [PubMed]
28. Wang W, Qian J, Zhu M, et al. Normothermic ex vivo lung perfusion outperforms conventional cold preservation in a deceased rat lung. Ann Transl Med 2022;10:99. [Crossref] [PubMed]
29. Gouchoe DA, Lee YG, Greenfield A, et al. Exploring Alternative Perfusion Solutions Using Next-Generation Polymerized Hemoglobin-Based Oxygen Carriers in a Model of Rat Ex Vivo Lung Perfusion. J Vis Exp 2024;
30. Chilvers NJS, Gilmour J, Brown ML, et al. A Split-Lung Ex Vivo Perfusion Model for Time- and Cost-Effective Evaluation of Therapeutic Interventions to the Human Donor Lung. Transpl Int 2024;37:12573. [Crossref] [PubMed]
31. Nilsson T, Gielis JF, Slama A, et al. Comparison of two strategies for ex vivo lung perfusion. J Heart Lung Transplant 2017; [Crossref]
32. Becker S, Steinmeyer J, Avsar M, et al. Evaluating acellular versus cellular perfusate composition during prolonged ex vivo lung perfusion after initial cold ischaemia for 24 hours. Transpl Int 2016;29:88-97. [Crossref] [PubMed]
33. Wahlberg JA, Southard JH, Belzer FO. Development of a cold storage solution for pancreas preservation. Cryobiology 1986;23:477-82. [Crossref] [PubMed]
34. Wittwer T, Albes JM, Fehrenbach A, et al. Experimental lung preservation with Perfadex: effect of the NO-donor nitroglycerin on postischemic outcome. J Thorac Cardiovasc Surg 2003;125:1208-16. [Crossref] [PubMed]
35. Oto T, Griffiths AP, Rosenfeldt F, et al. Early outcomes comparing Perfadex, Euro-Collins, and Papworth solutions in lung transplantation. Ann Thorac Surg 2006;82:1842-8. [Crossref] [PubMed]
36. Ku TJY, Ribeiro RVP, Ferreira VH, et al. Ex-vivo delivery of monoclonal antibody (Rituximab) to treat human donor lungs prior to transplantation. EBioMedicine 2020;60:102994. [Crossref] [PubMed]
37. Lonati C, Battistin M, Dondossola DE, et al. NDP-MSH treatment recovers marginal lungs during ex vivo lung perfusion (EVLP). Peptides 2021;141:170552. [Crossref] [PubMed]
38. Hasenauer A, Bédat B, Parapanov R, et al. Effects of cold or warm ischemia and ex-vivo lung perfusion on the release of damage associated molecular patterns and inflammatory cytokines in experimental lung transplantation. J Heart Lung Transplant 2021;40:905-16. [Crossref] [PubMed]
39. Ojanguren A, Parapanov R, Debonneville A, et al. Therapeutic reconditioning of damaged lungs by transient heat stress during ex vivo lung perfusion. Am J Transplant 2023;23:1130-44. [Crossref] [PubMed]
40. Arni S, Maeyashiki T, Latshang T, et al. Ex Vivo Lung Perfusion with K(ATP) Channel Modulators Antagonize Ischemia Reperfusion Injury. Cells 2021;10:2296. [Crossref] [PubMed]
41. Wang X, Wang Y, Parapanov R, et al. Pharmacological Reconditioning of Marginal Donor Rat Lungs Using Inhibitors of Peroxynitrite and Poly (ADP-ribose) Polymerase During Ex Vivo Lung Perfusion. Transplantation 2016;100:1465-73. [Crossref] [PubMed]
42. Becerra D, Linge H, Jeffs S, et al. Liquid Ventilation Reconditions Isolated Rat Lungs Following Ischemia-Reperfusion Injury. Tissue Eng Part A 2022;28:918-28. [Crossref] [PubMed]

(English Language Editor: J. Gray)