Anna Niroomand
George Emilian Nita
Sandra Lindstedt- 1Department of Clinical Sciences, Lund University, Lund, Sweden
- 2Lund Stem Cell Center, Lund University, Lund, Sweden
- 3Department of Cardiothoracic Surgery and Transplantation, Skåne University Hospital, Lund, Sweden
- 4Department of Transplantation Surgery, Liverpool University Hospitals NHS Foundation Trust, Liverpool, United Kingdom
- 5Institute of Systems, Molecular and Integrative Biology, University of Liverpool, Liverpool, United Kingdom
- 6Division of Transplantation Surgery, Karolinska University Hospital, Stockholm, Sweden
- 7Wallenberg Center for Molecular Medicine, Lund University, Lund, Sweden
Solid organ transplantation has progressed rapidly over the decades from the first experimental procedures to its role in the modern era as an established treatment for end-stage organ disease. Solid organ transplantation including liver, kidney, pancreas, heart, and lung transplantation, is the definitive option for many patients, but despite the advances that have been made, there are still significant challenges in meeting the demand for viable donor grafts. Furthermore, post-operatively, the recipient faces several hurdles, including poor early outcomes like primary graft dysfunction and acute and chronic forms of graft rejection. In an effort to address these issues, innovations in organ engineering and treatment have been developed. This review covers efforts made to expand the donor pool including bioengineering techniques and the use of ex vivo graft perfusion. It also covers modifications and treatments that have been trialed, in addition to research efforts in both abdominal organs and thoracic organs. Overall, this article discusses recent innovations in machine perfusion and organ bioengineering with the aim of improving and increasing the quality of donor organs.
Introduction
Since the start of successful kidney transplantation in the 1950s followed by other solid organs, transplantation has taken on a steadfast role in medicine as the definitive treatment to improve the quality of life and prolong the survival of patients with end-stage organ disease [1–6]. In recent decades, major advances in surgical technique, and pre-and post-operative management have allowed for the prolonged survival of recipients. As the number and availability of transplants have increased, the imbalance between organ supply and demand has grown to unprecedented levels [7]. In 2024, the Organ Procurement and Transplantation Data showed over 113,000 patients waiting for a transplant on the US all-organ waiting list [8], while the NHS Blood and Transplant reported over 7,000 [9] potential recipients in the UK alone and Eurotransplant listed over 13,000 patients currently active in eight European countries [10].
To address this disparity, an increase in the number of available donor organs is needed, which could theoretically be accomplished through a variety of means, including increased organ donation, procurement, and recuperation of available grafts. Restoration and regeneration of damaged donor organs is a particularly promising path for recovering a greater number of donor grafts, considering the high percentage that are currently discarded.
Grafts considered for transplantation may be rejected for several reasons. Factors such as the age of the donor, the cause of death, and the resulting sequelae of damage to the organ are often the primary reasons for rejecting the organ for transplantation. In the evaluation of donor lungs, for example, injury from trauma, infection, and aspiration may lead to acute lung injury (ALI) and the resulting functional impairment may preclude the organ from being chosen for transplantation [11, 12].
Many of these once-rejected organs are now being reconsidered in an effort to expand the donor pool. For example, the limits of what constitutes an acceptable organ have been raised with the introduction of expanded criteria for donors, donors at increased risk, and donors with acute kidney injury [13–17]. Despite these efforts, there are still organs being discarded that could theoretically be recovered for use via advanced techniques. The utilization of bioengineered organs is an expanding and promising approach and the creation of functional bioengineered organs could contribute to an immunosuppression-free state [18]. This article reviews the use of machine perfusion in several organs, exploring both the clinical application of this technology and the bioengineering techniques leveraged to increase the quality of donor organs.
Machine Perfusion
Circulatory support of donor organs following organ retrieval has been developed for several solid organs, with the advantage of such ex vivo platforms being the ability to evaluate the functionality of organs, prolong the preservation time between explant and implant, and apply recovery therapies to the target organ.
Since Alexis Carrel and Charles Lindbergh first described ex vivo machine perfusion in 1935, reporting the successful perfusion of the feline thyroid gland [19], significant efforts have been made to assess organ function and prolong preservation. In the 1970s animal models of the lung [20] and Hardesty and Griffith’s publication on autoperfusion of combined heart and lung transplants were highlighted [21]. Various liver-assisting therapies were researched during the 1960s and 1970s (cross-circulation, hemodilution and plasmapheresis), but they were abandoned with the development of orthotopic liver transplantation and understanding of the possible immunologic implications [22]. Animal models utilizing cold perfusion, oxygenated blood or plasma for kidney [23–25] and pancreas preservation continued to develop in the early 1980s [26, 27]. In Lund, Sweden, ex vivo lung perfusion (EVLP) was brought back by Steen et al. with the first marginally viable lungs transplanted following reconditioning [28–30]. This EVLP model was further developed with the establishment of protocols for prolonged evaluation by the Toronto group [31], which led to increasing and expanding clinical application.
At their core, machine perfusion platforms consist of an organ chamber in which perfusate flows through the graft using a pump (either centrifugal or roller) via a circuit equipped with a temperature control system and an oxygenation method. In the case of lung machine perfusion called EVLP, the organ is additionally ventilated with mechanical ventilation at settings determined by the underlying protocol. In general, the flow of perfusate allows for the delivery of oxygen and nutrients as well as the removal of waste and toxins, while also providing the opportunity to deliver therapeutic agents directly to the target. Leukocyte filters may also be incorporated into the circuit, although the efficacy of such an addition is under debate [32].
Lung Perfusion
In EVLP, the organ is both perfused and ventilated, which allows for additional graft monitoring both through analysis of the perfusate as well as through bronchoscopy and bronchoalveolar lavage fluid collection. An important note about the circuit specific to the lung is the need to deoxygenate the perfusate to assess the organ’s functionality, with the ratio of arterial oxygenation to a fraction of inspired air (PaO2/FiO2) being an important clinical measure.
The two most commonly used commercially available platforms for performing EVLP are the XPS™ system made by XVIVO [33] and the portable Organ Care System™ (OCS) produced by Transmedics [34]. Among these platforms, there are also three commonly referenced protocols, including the Lund protocol [28], the Toronto protocol [35], and the OCS protocol [36]. These differ in several parameters including the target perfusate flow (often calculated based on cardiac output), whether the left atrium of the donor lung graft is open or closed, and the composition of the perfusate. Regarding the perfusate, the differences lie in the underlying base solution (STEEN vs. OCS), and the use of red blood cells vs. acellular perfusate (Lund vs. Toronto). The debate over the use of cellular versus acellular perfusate continues, with some reports finding that cellular perfusate achieves results approaching clinical standards and superior lung preservation as well as a reduced incidence of lung edema and improved compliance in a porcine model. Despite these preclinical findings, human studies using EVLP have focused on acellular perfusates with favorable results.
Pioneering investigations have established the efficacy and utility of these systems in clinical applications. Starting in 2011, the Toronto group demonstrated in the HELP trial that transplantation with high-risk donor lungs maintained on 4 h of EVLP could produce comparable results to conventional lungs using their EVLP protocol [31]. The NOVEL trial which followed a few years later in 2014 extended these findings to a multicenter trial in 6 US centers with expanded criteria donors compared to conventional lungs and again encouraged the use of EVLP [37]. In the 2016 DEVELOP-UK trial, the EVLP observational cohort was terminated early, with lower survival rates, higher rates of ECMO and early grade 3 primary graft dysfunction (PGD) in the EVLP arm which utilized lungs not suitable for standard transplantation [38]. Other reports subsequently have emerged that provide contrasting support for the continued use of EVLP, including the 2018 INSPIRE and 2019 EXPAND trials using the OCS system [36, 39]. The INSPIRE trial compared standard criteria lung transplants maintained with EVLP versus cold storage, with findings of reduced PGD grade 3 at 72 h in the EVLP arm and meeting the primary safety endpoints. The EXPAND trial was then expanded to include extended criteria donors with primary endpoints of 30-day survival and absence of grade 3 PGD. Although the performance goals of the study were not met, the study group concluded that the use of EVLP allowed the transplantation of organs that would otherwise have been rejected. Similarly, supportive data for EVLP from a single-center analysis in Vienna was published in 2017 reporting that in their use of EVLP in standard donor lungs, EVLP use provided evidence of functional results and operative outcomes comparable to standard preservation, with the cited benefit of safely extending preservation time [40].
Clinical EVLP systems have since evolved to be executed not only by a transplant service’s own team (referred to as local in-house EVLP) but also to be performed at an outside but centralized instutution (sometimes called a centralized lung evaluation system). This centralized version has recently gained traction due to the development of Lung Bioengineering’s dedicated facility in which remote EVLP was performed for seven different United States-based lung centers [41]. Such centralized EVLP may grow in popularity with the advent of the first European centralized facility coordinating EVLP between Sweden and Denmark [42].
Within these applications, the details of optimal parameters are still being discussed and guidelines for optimal execution need to be established. Ventilation methods are an additional point of interest, particularly because of the known effects of harmful ventilation causing ventilator-induced lung injury. Because excessive pressure (barotrauma), alveolar collapse (atelectrauma), and volume (volutrauma) can induce iatrogenic damage to the graft, attention to the method of ventilation is critical. The stress index which represents the rate of change in compliance during tidal inflation has been proposed as a means of assessing ventilation in EVLP, with high indices corresponding to hyperinflation and low indices corresponding to recruitment-derecruitment [43, 44]. Ventilation outside of the stress index window has been reported to correlate with cytokine-driven inflammation and longer time on mechanical ventilation, intensive care unit (ICU) stays, and hospital stays [45]. Using an auxiliary device, exhaled breath particles can also be collected from the mechanical ventilation setup on EVLP, which has shown that greater numbers of particles are correlated with higher tidal volumes and volume-controlled ventilation [46]. Higher particle flow rates using this methodology have been correlated with lung injury [47, 48] and with primary graft dysfunction in the transplant setting [49]. Other studies have investigated airway pressure release ventilation compared to conventional volume-controlled ventilation in a porcine model to try an open lung approach to ventilation, which reduced edema during EVLP and improved PaO2/FiO2 ratios following 4 h of reperfusion after left lung transplantation [50]. Negative pressure ventilation was trialed in another porcine experiment with 12 h of ex vivo perfusion and found to have lower levels of pro-inflammatory TNF-α, IL-6 and IL-8 along with less edema during EVLP [51].
Heart Perfusion
Similar to the lung, the perfusion circuit of the ex vivo machine can be altered to accommodate the specific needs of the heart, including wires that allow for defibrillation and pacing. Currently, the two systems in place for ex vivo heart perfusion (EVHP) are the Heart Box™ (XVIVO) and the portable Organ Care System (OCS) Heart™ (TransMedics). The main difference between these systems is the use of normothermic perfusion in the OCS Heart versus hypothermic preservation at 8°C in the Heart Box. The reported benefits of EVHP in these systems are notable not only for the clinical results obtained, but also for the system’s ability to assess hidden cardiac pathologies [52] and to find specific and sensitive markers of outcome [53].
In the 2014 trial of normothermic heart perfusion, standard criteria donors were compared against conventional preservation techniques, looking at the survival of the recipients over a 2-year period [54]. During this time, survival rates were comparable in the early 30-day post-transplant period, and surpassed cold static storage at one and 2 years along with a reduction (although not to a statistically significant degree) in primary graft dysfunction. In the European-based PROTECT trial [55], EVHP was evaluated for safety and was followed several years later by the PROCEED II trial, a prospective, randomized non-inferiority trial in 10 different transplant centers [56]. Short-term clinical outcomes were again comparable between groups at 30 days and in a one-center follow-up to the 2-year mark, there was again no significant difference in survival despite significantly longer ischemic times in the EVHP group [57]. Separately from the EXPAND lung trial, an EXPAND heart preservation trial examined the utility of the system for extended criteria donors, with primary endpoints of survival at 30 days and absence of PGD at 24-h post-transplant [58]. In this study, the perfusion was not compared to standard preservation techniques, but did show high utilization of donor grafts, low rates of PGD, and high survival rates. To further characterize its use in unfavorable donors, Sáez et al. placed extended criteria donor hearts on machine perfusion, describing their single-center experience using the system and reporting improved short-term outcomes, although this was not directly compared to standard criteria [59]. In the same group, patients who were bridged to transplantation using left ventricular assist devices were compared to those whose donor organ was preserved with EVHP versus standard cold storage [60]. Recipients whose grafts had received machine perfusion had lower rates of mechanical circulatory support and decreased need for blood transfusion with longer out-of-body graft time, although ultimately there was no significant difference in overall survival [60]. Another examination of extended criteria transplants supported by EVHP compared to conventional storage showed that there was an effective increase in the number of transplants possible given the recovery of organs using the system, and that the survival of the donor hearts was comparable in terms of both hospital length of stay and survival [61]. A long-term follow-up report of normothermic EVHP from the Berlin group followed recipients 4 years post-transplantation and showed high survival rates, supporting the use of the platform [62]. Other notable smaller studies include the case report of prolonged out-of-body time of 10 h using machine perfusion [63], as well as a case series from St. Vincent’s in Australia demonstrating distant procurements on donation after cardiac death (DCD) with favorable results [64]. This group also demonstrated favorable lactate profiles during EVHP in their porcine DCD model, in which they also reported that following orthotopic transplantation, only those previously placed on EVHP were able to be weaned from cardiopulmonary bypass support [65].
The hypothermic alternative to EVHP has been trialed in both porcine and human cardiac transplantation. In a study of porcine DCD procurement, following normothermic regional perfusion, cold storage was compared to hypothermic machine perfusion, finding that contractility was only found with hypothermic EVHP along with a lower need for inotropic support and signs of myocardial damage [66]. A trial of hypothermic EVHP was run by the Australian St. Vincent’s group, with a demonstration of 13 transplants, in which one patient required post-operative VA-ECMO and no postoperative mortality was observed at 30 days [67]. In the first prospective trial of hypothermic EVHP by the Lund group, standard criteria donors were compared in both modalities, showing improved mortality and cardiac-related adverse events in the machine preservation arm [68]. The HOPE trial followed with a multicenter investigation of the system in donor hearts with both short and long preservation times, up to 8 h and 47 min. When compared to comparator data retrieved from the International Society of Heart and Lung Transplantation (ISHLT), the EVHP group demonstrated improved 30-day survival and low rates of PGD development [69].
Liver, Kidney, and Pancreas Perfusion
The machine perfusion landscape in liver, kidney, and pancreas transplantation has been similar to that of thoracic organ transplantation. Attempts to improve donor organ viability, functional assessment, and overall graft and patient outcomes led to the rapid expansion of ex vivo hypo- and normothermic machine perfusion (HMP, NMP) and in vivo normothermic regional perfusion (NRP).
The concept of NMP was developed in the 1960s when it was initially studied for kidney preservation. Its development over the years has expanded its application to the liver [70], significantly improving outcomes by lowering graft injury, and discarded organs and increasing the mean preservation time compared to conventional static cold storage (SCS) techniques. Similarly, HMP was conceptualized in the 1960s. Significant advances in HMP technology were made in the 1990s through the pioneering work of Belzer and Southard [71], who developed the University of Wisconsin (UW) solution for the preservation of the liver, kidney, and pancreas.
A systematic review and meta-analysis by Jakubauskas et al. compared outcomes between NMP and HMP in liver transplantation versus static cold storage (SCS). Their study demonstrated the role of machine perfusion in reducing graft injury and decreasing the rate of liver graft rejection, early allograft dysfunction (EAD) and non-anastomotic biliary strictures compared to SCS [72].
Machine perfusion in kidney transplantation showed a similar significant improvement in graft outcomes. The international randomized controlled trial by Moers et al. demonstrated a decrease in delayed graft function and an improvement in graft survival rates when utilizing HMP as opposed to SCS alone [73].
Ex vivo machine perfusion has also been utilized in the field of solid pancreas transplantation for quality assessment of discarded organs [74] and to improve preservation capacity, both the ability to resuscitate organs and to reduce delayed graft function [75].
Organ Engineering on Machine Perfusion
As previously mentioned, the discrepancy between the availability of donor organs and the number of potential recipients waiting for a graft demands the development of novel therapies and treatments to bridge this gap. While ex vivo perfusion itself can help to evaluate marginal organs that would otherwise be discarded or facilitate the transplantation of more distantly procured grafts, there are many rejected damaged donor organs that could be retrieved. The addition of different therapies to the machine perfusion circuit presents an opportunity to recover a greater number of grafts. Advantages include highly specific temporal and spatial control of treatment delivery along with the avoidance of potential systemic effects.
Gene Therapy
Following the adoption of viral-based vectors, gene therapy has been pursued for its potential in immunomodulation and mitigation of the inflammatory response often seen in the events surrounding transplantation.
In cardiac transplantation, early rat models using adenovirus vectors carrying LacZ demonstrated that transfection could occur with minimal effect on graft survival, but the results were tempered by short gene expression lasting only weeks [76, 77]. Subsequent efforts have demonstrated that the hypothermic ex vivo platform can be a reliable conduit for vector delivery, with further development of adenovirus vectors of LacZ in both porcine and rat models and a need for continuous perfusion beyond single injections [78–80]. Further testing in rabbit models of normothermic EVHP demonstrated transfection using similar vector-gene combinations [81, 82]. Notably, the Duke group produced a porcine model in which luciferase was transfected and investigated the components of the platform that promoted or hindered successful transfection [83] in a 5 day post-transplantation model. This group additionally reported on a recombinant adeno-associated virus carrying luciferase that showed high efficiency and expression for 30 days without off-target effects or signs of rejection [84].
Within the heart, targets mainly aimed at immunomodulation have been identified and successfully delivered in animal models, including constructions of interleukin-10 or transforming growth factor-beta 1 (TGF-β1), in which transplanted grafts have shown higher vector concentrations as well as prolonged graft survival and inhibition of allograft rejection [85, 86]. Other vectors have included the delivery of a CTLA4 gene with a demonstration of localized immunosuppression in rats [87] and liposome-based delivery of IL-4 and IL-10 to rabbits with significantly increased survival and reduced rejection [88].
In the lung, gene therapy has also gained momentum for its potential for immune regulation. An important decision point is the route of administration: intravenous delivery is theoretically better at targeting the endothelium while the endotracheal route can target the epithelium. Several studies have investigated these routes of delivery, with the endotracheal route often being preferred [89, 90]. In earlier studies performed on rat transplantation by the Toronto group, the feasibility of gene transfection was explored using six and 12 hour ex vivo transfusions, although importantly this was done in cold preservation prior to the advent of modern EVLP [91]. In further developments of the model, human IL-10 within this adenovirus vector was studied, again with six or 12 h of ex vivo incubation with a reported improvement in post-transplant organ function [92]. The IL-10-containing vector was further examined in porcine and human lungs, now placed on modern EVLP systems, with endotracheal administration of treatment [93]. In the porcine model, following transplantation with a 4 h follow-up, cytokines IL-6 and IL-1β were significantly decreased in the transfected group and the rejected human organs, there was no observed deterioration in function on EVLP. The group compared delivery of the viral vector using an ex vivo model with an in vivo model, and in both cases transplanted the lung to assess post-transplant function. The ex vivo group showed improved lung function compared to the in vivo group, with the authors hypothesizing that this methodology led to less inflammation associated with vector delivery to explain the improved function [94]. In a prolonged 7-day survival model, the same therapy was applied, demonstrating continued IL-10 expression through day 7 in addition to a lack of systemic toxicity, and improved lung function on day 7 in the treated group compared to the control conditions [95]. Of note, transfection with an IL-10-carrying vector has been shown in rodent models of transplantation without the use of EVLP and has shown decreased rates of histologically scored acute rejection [96], lower levels of inducible nitric oxide synthase [97], as well as improved gas exchange and decreased IL-2 expression [98].
Gene therapy in the lung has also been pursued with other targets, notably with the delivery of small interfering double-stranded RNA (siRNA) via lentiviral vectors targeting the class-II-trans-activator and β2-microglobulin, which ultimately reduced MHC-1 and II transcription in lung endothelial cells in a porcine model [99]. This mechanism leveraged by Figueiredo et al. allows for the evasion of the transfected cells from immune system activation, a valuable attribute when transplanting a donor organ into a recipient.
Gene therapy in kidney transplantation offers a novel approach to addressing common post-transplant complications such as rejection, ischemia-reperfusion injury (IRI), and the nephrotoxic effects of immunosuppressive drugs. By targeting the genetic foundations of these issues, gene therapy has the potential to significantly improve transplant outcomes. Bogacz et al. investigated the correlation between tacrolimus dose and genetic variation for IL-10 and its effect on therapeutic outcomes in kidney transplantation patients, revealing the potential influence of IL-10 polymorphism on immunosuppressive drug dosage and the risk of acute graft rejection [100]. In cold-IRI mouse models, the anti-IL-2 immune complex was found to attenuate ischemia-reperfusion injury (IRI) after kidney transplantation by increasing renal regulatory T cells [101].
In the pancreas, gene therapy can introduce encoding for immunomodulatory proteins (e.g., interleukin-10, TGF-beta) to create a local immunosuppressive environment. An example is the role of the VEGF gene in modifying isolated pancreatic islets for transplantation. VEGF has anti-apoptotic and angiogenic effects that are critical for islets under hypoxic stress during transplantation. Studies reveal that islets modified with the VEGF gene display have improved viability and more effective glucose regulation post-transplantation compared to non-modified islets [102].
Gene therapy in liver models in the context of transplantation remains limited [103–105]. Lorvellec et al. and the UCL group designed an in vitro whole liver [106].The “Bioreactor grown Artificial Liver Model” (BALM) concept is a custom-designed 3D culture of human induced pluripotent stem cell-derived hepatocyte-like cells (hiHEPs) on a decellularized mouse liver scaffold. Adenoviral and lentiviral vectors introduced by intravascular injection have demonstrated that BALM has the potential to serve as a substitute for some in vitro and in vivo therapeutic testing methods.
It is of course important to consider the effects of the therapy delivered to the target organ and whether there are any unintended consequences. Potential risks of gene therapy include the possibility of certain types of cancer in the long term, including the suspension of Bluebird Bio’s clinical trial of LentiGlobin gene therapy for sickle cell disease due to cancer diagnoses in two recipients 5 years after the treatment [107]. Although the lentiviral vector may not have been the direct cause of cancer, there are still concerns about how well we really understand the long-term sequelae of the treatments we deliver. This highlights the need for studies that not only look at the immediate consequences of gene therapy in transplantation, but also at what might occur to treated organs years down the line. Other hypothetical concerns include how gene therapy could potentially activate the immune system, which would be of great concern in immunosuppressed transplant recipients, and whether there are off-target effects, which would ideally be mitigated by the use of therapies in an ex vivo setting where the therapy was applied directly to the target organ. Of course, it is critical to note that gene therapy is not a panacea for all transplantation troubles. Often, the disease processes that emerge are complicated and multifactorial and it is unlikely that any one target identified by gene therapy will provide a universal answer to transplantation-related complications. It is likely that multimodal therapies will be needed in order to truly optimize grafts for transplantation.
Mesenchymal Stromal Cells (MSCs)
Cell therapy has emerged as a form of treatment for damage owing to ischemia-reperfusion injury (IRI) and acute lung injury due to the properties exhibited by mesenchymal stromal cells (MSCs). This cell population has been shown to act on inflammatory pathways and play a role in immunomodulation, with multi-targeted effects and has been studied for its paracrine effects leading to tissue regeneration. Bone marrow-derived MSCs have been linked to increased levels of IL-10 to promote anti-inflammation [108] as well as the secretion of immunomodulatory growth factors [109]. A further advantage is the lack of co-stimulatory CD40, CD40L, CD80, and CD86 which confer a lack of immunogenicity to MSCs [110], making them a suitable therapy for immunosuppressed and sensitive recipients.
In lung transplantation, EVLP has provided a platform for MSC delivery. At the University of California, San Francisco, cells applied during ex vivo perfusion were shown to restore endothelial permeability in damaged lobes of human lungs and improve alveolar fluid clearance [111]. Improved alveolar fluid clearance was also observed with the treatment of cell culture medium conditioned with MSCs without the cells themselves. Further studies of rejected human lungs within this group also showed improvement in alveolar fluid clearance with 5 × 106 cells instilled into the lungs on EVLP [112]. In studies of cells isolated from human umbilical cord tissue by the Toronto group, administration of MSCs aimed at ameliorating prolonged cold ischemia in previously healthy donor lungs was performed first with the intent of determining ideal cell concentrations following a comparison of intrabronchial vs. intravascular routes of administration [113]. As a result, intravascular administration was found to be superior given the higher airway pressures associated with intrabronchial doses, as well as the identification of a suitable dose equivalent to 5 × 106 cells per kg in the porcine model. In a longer cold storage experiment with 24 h of cold ischemia followed by 12 h of EVLP, MSCs showed decreased apoptosis and reduced pulmonary edema after transplantation consisting of 4 h of reperfusion [114]. In an effort to leverage the immunomodulatory effects of MSCs, cells were engineered to produce anti-inflammatory IL-10 and were given to human lungs rejected for transplantation under 12 h of EVLP [115]. In this study, there were no significant differences in pulmonary vascular resistance, oxygenation capacity, and compliance compared to control lungs, which was attributed to an acidic lung microenvironment and heterogeneity of injury across human organs. Umbilical cord-derived MSCs were also examined during rat EVLP by Pacienza et al., indicating that intravascular infusion showed better lung compliance in addition to a reduction in neutrophil-related markers, supporting an anti-inflammatory role of the cell treatment [116].
In the heart, ex vivo perfusion has only been studied with the application of MSC-conditioned medium, rather than the stromal cells themselves. In an investigation of hypothermic perfusion in rats, a conditioned medium was associated with the downregulation of proinflammatory cytokines following transplantation and was hypothesized to protect against myocardial IRI [117]. In a later study by the same group, the conditioned medium was tested in brain-dead donors, showing improved systolic function after preservation and reduced cell apoptosis [118]. In normothermic EVHP, the conditioned medium as studied by the Guangzhou group showed that oxidative stress, inflammation, and apoptosis were mediated by the treatment and that warm ischemic injury was improved [119]. When the same group transplanted the conditioned hearts, spontaneous contraction and left ventricular systolic diameter improved in the treated group and lower levels of plasma cytokines were detected [120]. Compared to whole-cell treatment, these studies offer an opportunity to explore the conditioned medium as a therapy in and of itself, which may have advantages over MSC administration in terms of both time and cost savings. Interestingly, MSCs have been studied in the treatment of ischemic and non-ischemic cardiomyopathy and have even been labeled as cardioprotective, implying an avenue for further investigation in the transplantation setting [121–127].
In the kidney, MSC therapy has also been studied for its potential to repair damage from ischemia, which was tested in a human kidney model of ex vivo perfusion. MSC treatment showed reduced inflammatory cytokines and increased adenosine triphosphate and growth factors [128]. In a rat model, kidney grafts were treated with MSCs or with extracellular vesicles released by the MSCs, both of which were associated with improved metabolism and ion transport and Gregorini et al concluded that both forms of treatment protected the kidneys from reperfusion damage [129]. In later work by Pool et al, porcine kidneys were treated with MSCs during normothermic machine perfusion and labeling of bone marrow-derived MSCs showed that the cells rapidly decreased in the circulating perfusate over time and that a small portion was intact and appeared within the glomeruli [130]. The authors noted that the study demonstrated that the cells can reach and reside in the kidney, but that the infusion rate must be high enough to visualize within the capillaries. The group then also reported that with normothermic perfusion, MSC treatment at these sufficiently high cell infusion rates reduced injury markers and induced the release of immunomodulatory cytokines during ischemia-reperfusion injury [131]. MSC treatment was further followed in a transplantation model by Lohmann et al. with the finding that while MSC treatment was safe and well tolerated within their autotransplant cohort, there were no significant effects of the therapy within the 14 days post-transplantation [132]. An important question within this line of work is the dose tolerance of the kidneys to large amounts of MSCs, which was investigated by the same group through their porcine model comparing doses of ten million and one hundred million MSCs injected into the renal artery [133]. The lower dose was well tolerated, while the high dose resulted in tissue inflammation and glomerular and tubular damage. A balance must be struck between administering doses of cell therapy that are both effective and not so high as to cause damage themselves.
Extending the investigation of MSC therapy to the liver, several studies have examined the efficacy of the therapy within this solid organ. In a rat model, MSCs were administered during machine perfusion following donation after cardiac death, with treatment correlating with improved bile production and histologic findings [134]. MSC treatment was shown by another group to ameliorate injury due to oxidative stress and increase mitochondrial function within their rat liver cohort [135], which was further extended in other studies to demonstrate improved histopathology, hepatocyte ultrastructure, and microcirculation-related indices [135, 136]. In a porcine model, MSCs were shown to be retained in a machine-perfused liver with the reported continuation of their paracrine activity via increased human-specific IL-6 and IL-8 levels in porcine blood [137].
Cross-Circulation
Although the system does not employ a machine, the perfusion method of cross-circulation is noteworthy for its novelty in the development of methods to regenerate damaged donor lungs and livers. Reported advantages of cross-circulation over machine perfusion include longer recovery times and an increased duration of extracorporeal support [138]. An early report of cross-circulation in the lung appeared under the work of Dr. Joel Cooper who reported cross-circulating aspirin-treated sheep with non-treated sheep to examine the mixing of the circulating blood volumes and to study the circulating elements [139]. More recent work has emerged to establish a cross-circulation platform as a means to rehabilitate damaged donor lungs. The Bacchetta group first reported the use of cross-circulation to achieve prolonged perfusion times in excess of 36 h, followed by a porcine study in which lungs damaged by the aspiration of gastric contents were then reconditioned using the platform [138, 140]. To demonstrate the system’s ability to provide prolonged support, porcine lungs were additionally maintained in cross-circulation for 4 days, with preserved lung function and decreased levels of several cytokines, including IL-1β, IL-8, and IL-12 among others [141]. Gastric aspiration-damaged recipients were also shown to help recover donor lungs subjected to prolonged cold storage in a model that leveraged injury in both the donor and recipient within cross-circulation [142]. In a step closer to clinical application, rejected human lungs were added to the xenogeneic circuit to show functional and histological recovery under the platform [143, 144] with 24 h of support. There are of course concerns about the immunogenic response that might follow when exposing human lungs to a xenogeneic recipient. Wu et al reported that in their established cross-circulation model, porcine immune cells and immunoglobulins can be found infiltrating human donor lung tissue after 24 h of xenogeneic perfusion [145]. Similarly, in an investigation by Glorion et al., immune cells from the porcine host were found in lung cell suspensions, in bronchoalveolar lavage fluid, and in histological examination of lung tissue [146]. The group also posited that cross-circulation could act as a platform to investigate the immune processes that follow transplantation without carrying out full transplantation and could serve as a means to assess anti-inflammatory and immunomodulatory drugs based on the response of the porcine host.
Cross-circulation has also been investigated in porcine livers, as early as 2001 when a group from Kyoto studied porcine liver perfusion with baboon cross-circulation [147]. The group found microthrombi formation in the perfused porcine livers and macroscopic hemolysis in 3 out of 5 cases with a maximum duration of 6 h of cross-circulation. In a publication from the same year, Nara et al. in Hirosaki, Japan cross-circulated hepatectomized dogs with donor pigs to test a semipermeable membrane in their model of double filtration plasmapheresis cross-circulation [148]. This model was further explored by the group in their later work with a canine whole liver supporting a porcine model of hepatic failure induced by lipopolysaccharide and alpha-amanitin, which demonstrated improvements compared to the porcine group not supported by the liver cross-circulation [149]. The group reported that the lack of immune-mediated reactions was the result of a semipermeable membrane blocking the movement of IgM, which they investigated in later work [150].
Bacchetta et al, who had previously investigated lung cross-circulation also followed up with work on liver cross-circulation in 2022, with a publication using a porcine host for an extracorporeal porcine liver to show improvements in synthetic function, metabolic activity, and histopathological examination [151]. This effort was then replicated using explanted human donor livers that were supported by xenogeneic cross-circulation, which also demonstrated improved histopathologic assessment at 24 h of support [152].
While other organs have not been investigated as extensively, there are reports of cross-circulation being studied in heart transplantation. In one model, standard ex vivo cardiac perfusion in a porcine graft was compared to EVHP supplemented with cross-circulation from a paracorporeal support pig [153]. This was then further compared to a third setup in which blood from the support animal was filtered into plasma during cross-circulation. Cross-circulation was reported in both set-ups to have high oxygen consumption and vascular resistance, with the conclusion that it improved preservation compared to standard EVHP. Cross-circulation was also used to support EVHP for 72 h in a study by the Extracorporeal Life Support Laboratory at the University of Michigan, which supported ovine hearts with paracorporeal sheep. All cross-circulated hearts were deemed suitable for transplantation by the end of 72 h, while the control group, which was not cross-circulated, all failed after 6–10 h of EVHP [154]. The group then further explored porcine EVHP support in a publication showing that EVHP could be maintained for 24 h with plasma exchange, which they reported eliminated the need for a paracorporeal animal [155].
Future Directions in Bioengineering Strategies
The development of bioengineered organs that could be produced in the laboratory would be the ultimate step in alleviating the problems caused by the shortage of donor organs. In the quest to develop such a product, techniques have emerged to grow or build organs. One such method uses organoids made by three-dimensional cell cultures formed from patient-derived cells designed to replicate in vitro the microarchitectural and functional characteristics of their in vivo counterparts. One advantage of building such models is the ability to further study diseases and potential drugs and therapies on a platform that closely resembles human physiology while reducing the need for animal-based research. In the lung, for example, the advances made in microfluidic organ-on-a-chip devices have been driven by the study of specific disease processes such as those arising from particulate exposure, pulmonary fibrosis, and acute respiratory distress syndrome [156–158].
The study of human organ physiology and disease occurrence through microfluidic devices can be extrapolated for use in transplantation surgery in general. In transplantation, these could represent an avenue for patient-specific study of response to immunosuppression in an effort to optimize regimens, and minimize toxicity, adverse drug reactions, and side effects, balancing these against the risk of rejection [159].
Another potential application is the detection of transplant rejection, which remains a clinical challenge given the high rates of both acute and chronic forms of immune-mediated rejection. Madhvapathy et al. used rat models in which a wireless electronics interface was connected to transplanted kidneys, allowing for real-time monitoring of ultradian rhythms, disruption of the circadian cycle, and organ temperature. These patterns change once immunosuppression is discontinued before creatinine elevation [160].
On top of leveraging advanced techniques to further study transplant-related complications, bioengineering principles could potentially be used to develop artificial organs. In decellularization-recellularization experiments, cellular material is removed during a decellularization process to then be replenished with a new repopulation of cells during the recellularization phase. Decellularization of living tissue can be accomplished by chemical, enzymatic, physical, or combined approaches to generate an extracellular matrix scaffold that provides structural support for the future regenerated organ. The most common materials used for decellularization are a combination of detergents such as sodium dodecyl sulfate (SDS) and Triton-X100 [161]. This is an important step as retaining any cell membrane epitopes would trigger a host response in vivo. [162] Once the mechanical integrity has been established and decontaminated, the acellular scaffold undergoes the process of repopulation with organ-specific cells, thus reconstituting the organ from a functional perspective.
For the liver, in vitro studies of drug-induced hepatotoxicity, liver function and regenerative potential have advanced the development of primary hepatocyte isolation and the generation of two- and three-dimensional liver models, spheroids, organoids, slice cultures and microfluidic models [163, 164]. Decellularized human liver scaffolds have been repopulated with human cell lines and resulted in liver scaffold cubes that could be successfully xenotransplanted subcutaneously into immunocompetent mice, without any resulting immunogenic responses [165]. Other investigations in animal models have shown that decellularization of the whole organ liver is possible with preservation of the macrostructure [166]. While decellularization has been demonstrated using a variety of methodologies, there is no consensus on the most advantageous approach nor have any unified criteria been accepted [167].
Similarly, studies have reported successful recellularization of kidney scaffolds with preservation of microarchitecture and ECM components such as glycosaminoglycans (GAGs) [168–170]. After preloading of the vascular matrix with vascular endothelial growth factor (VEGF) and angiopoietin 1, pluripotent stem cell-derived endothelial cells can be delivered with efficient adherence, proliferation, and survival [168].
The lung is a challenging organ to bioengineer given the complexity of the cell population that constitutes it. Scaffold generation has been approached both from the angle of using human-derived acellular scaffolds and artificial scaffolds made from synthetic polymers [171–173]. Because there are over 40 different cell types that make up the lung, recellularization is a challenging task and a variety of sources including tissue-isolated progenitor cells, differentiated pluripotent stem cells, and mesenchymal stem cells are being investigated for their potential [174–177].
While these methodologies have a long way to go before they can be considered for clinical application, the progress being made is promising for the advent of engineered organs to meet the demand for transplantation.
Conclusion
As the definitive management for end-stage organ disease, transplantation remains an in-demand procedure. While surgical and perioperative techniques have progressed, there is still a notable shortcoming, notably the mismatch between the number of available organs and the need for grafts by potential recipients on the waiting list. In order to bridge the gap between supply and demand, new methods are needed for the recuperation of suitable organs. Machine perfusion provides an emerging platform for the recovery and regeneration of organs that would otherwise be discarded. Advancements made in gene therapy, applications of mesenchymal stem cells, cross-circulation methodologies, and the advent of advanced bioengineering principles have been covered in this review as promising avenues for organ recovery. While many of these approaches are in various stages of development and clinical application, they represent important advances that can be leveraged in the search for meaningful ways to increase the donor pool.
Author Contributions
All authors listed have made a substantial, direct, and intellectual contribution to the work and approved it for publication.
Funding
The authors declare that no financial support was received for the research, authorship, and/or publication of this article.
Conflict of Interest
The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.
References
1. Hariharan, S, Israni, AK, and Danovitch, G. Long-Term Survival After Kidney Transplantation. N Engl J Med (2021) 385(8):729–43. doi:10.1056/NEJMra2014530
2. Jain, A, Reyes, J, Kashyap, R, Dodson, SF, Demetris, AJ, Ruppert, K, et al. Long-Term Survival After Liver Transplantation in 4,000 Consecutive Patients at a Single Center. Ann Surg (2000) 232(4):490–500. doi:10.1097/00000658-200010000-00004
3. Venstrom, JM, McBride, MA, Rother, KI, Hirshberg, B, Orchard, TJ, and Harlan, DM. Survival After Pancreas Transplantation in Patients With Diabetes and Preserved Kidney Function. JAMA (2003) 290(21):2817–23. doi:10.1001/jama.290.21.2817
4. Hanlon, M, Cooper, M, and Abrams, P. Quality of Life After Pancreas Transplantation: Time to Look Again. Curr Opin Organ Transpl (2019) 24(4):451–5. doi:10.1097/MOT.0000000000000676
5. Dobbels, F, Bleser, LD, Geest, SD, and Fine, RN. Quality of Life After Kidney Transplantation: The Bright Side of Life? Adv Chronic Kidney Dis (2007) 14(4):370–8. doi:10.1053/j.ackd.2007.07.005
6. Onghena, L, Develtere, W, Poppe, C, Geerts, A, Troisi, R, Vanlander, A, et al. Quality of Life After Liver Transplantation: State of the Art. World J Hepatol (2016) 8(18):749–56. doi:10.4254/wjh.v8.i18.749
7. Lewis, A, Koukoura, A, Tsianos, GI, Gargavanis, AA, Nielsen, AA, and Vassiliadis, E. Organ Donation in the US and Europe: The Supply vs Demand Imbalance. Transpl Rev (2021) 35(2):100585. doi:10.1016/j.trre.2020.100585
8. Organ Procurement and Transplantation Network. National Data - OPTN. Available from: https://optn.transplant.hrsa.gov/data/view-data-reports/national-data/# (Accessed March 25, 2024).
9. NHS Blood and Transplant. Organ Donation and Transplantation. Available from: https://www.nhsbt.nhs.uk/what-we-do/transplantation-services/organ-donation-and-transplantation/(Accessed March 25, 2024).
10. Eurotransplant. Statistics. Available from: https://statistics.eurotransplant.org/index.php?search_type=waiting+list&search_organ=all+organs&search_region=&search_period=2024&search_characteristic=&search_text=&search_collection= (Accessed March 25, 2024).
11. Diamond, JM, Arcasoy, S, Kennedy, CC, Eberlein, M, Singer, JP, Patterson, GM, et al. Report of the International Society for Heart and Lung Transplantation Working Group on Primary Lung Graft Dysfunction, Part II: Epidemiology, Risk Factors, and Outcomes-A 2016 Consensus Group Statement of the International Society for Heart and Lung Transplantation. J Heart Lung Transpl Off Publ Int Soc Heart Transpl (2017) 36(10):1104–13. doi:10.1016/j.healun.2017.07.020
12. Huppert, LA, Matthay, MA, and Ware, LB. Pathogenesis of Acute Respiratory Distress Syndrome. Semin Respir Crit Care Med (2019) 40(1):31–9. doi:10.1055/s-0039-1683996
13. Schold, JD, Howard, RJ, Scicchitano, MJ, and Meier-Kriesche, HU. The Expanded Criteria Donor Policy: An Evaluation of Program Objectives and Indirect Ramifications. Am J Transpl (2006) 6(7):1689–95. doi:10.1111/j.1600-6143.2006.01390.x
14. Kizilbash, SJ, and Chavers, BM. Strategies to Expand the Deceased Donor Pool for Pediatric Kidney Transplant Recipients. Kidney360 (2020) 1(7):691–3. doi:10.34067/KID.0001772020
15. Francke, M, Wolfson, AM, Fong, MW, Nattiv, J, Pandya, K, Kawaguchi, ES, et al. New UNOS Allocation System Associated With No Added Benefit in Waitlist Outcomes and Worse Post-transplant Survival in Heart-Kidney Patients. J Heart Lung Transpl Off Publ Int Soc Heart Transpl (2023) 42(11):1529–42. doi:10.1016/j.healun.2023.06.018
16. Watson, CJE, Johnson, RJ, and Mumford, L. Overview of the Evolution of the UK Kidney Allocation Schemes. Curr Transpl Rep (2020) 7(2):140–4. doi:10.1007/s40472-020-00270-6
17. Nita, GE, Gopal, JP, Khambalia, HA, Moinuddin, Z, and van Dellen, D. Kidney Transplantation From Donors With Acute Kidney Injury: Are the Concerns Justified? A Systematic Review and Meta-Analysis. Transpl Int (2023) 36:11232. doi:10.3389/ti.2023.11232
18. Orlando, G, Wood, KJ, Soker, S, and Stratta, RJ. How Regenerative Medicine May Contribute to the Achievement of an Immunosuppression-Free State. Transplantation (2011) 92(8):e36–8. doi:10.1097/TP.0b013e31822f59d8
19. Carrel, A, and Lindbergh, CA. The Culture of Whole Organs. Science (1935) 81(2112):621–3. doi:10.1126/science.81.2112.621
20. Jirsch, DW, Fisk, RL, and Couves, CM. Ex vivo Evaluation of Stored Lungs. Ann Thorac Surg (1970) 10(2):163–8. doi:10.1016/s0003-4975(10)65582-8
21. Hardesty, RL, and Griffith, BP. Autoperfusion of the Heart and Lungs for Preservation During Distant Procurement. J Thorac Cardiovasc Surg (1987) 93(1):11–8. doi:10.1016/s0022-5223(19)36469-4
22. Stockmann, HBAC, Hiemstra, CA, Marquet, RL, and Ijzermans, JNM. Extracorporeal Perfusion for the Treatment of Acute Liver Failure. Ann Surg (2000) 231(4):460–70. doi:10.1097/00000658-200004000-00003
23. Humphries, AL, Russell, R, Stoddard, LD, and Moretz, WH. Successful Five-Day Kidney Preservation. Perfusion With Hypothermic, Diluted Plasma. Invest Urol (1968) 5(6):609–18.
24. Belzer, FO, Ashby, BS, and Dunphy, JE. 24-Hour and 72-Hour Preservation of Canine Kidneys. Lancet Lond Engl (1967) 2(7515):536–8. doi:10.1016/s0140-6736(67)90498-9
25. Collins, GM, Bravo-Shugarman, M, and Terasaki, PI. Kidney Preservation for Transportation. Initial Perfusion and 30 Hours’ Ice Storage. Lancet Lond Engl (1969) 2(7632):1219–22. doi:10.1016/s0140-6736(69)90753-3
26. Florack, G, Sutherland, DER, Heil, J, Squifflet, JP, and Najarian, JS. Preservation of Canine Segmental Pancreatic Autografts: Cold Storage Versus Pulsatile Machine Perfusion. J Surg Res (1983) 34(5):493–504. doi:10.1016/0022-4804(83)90101-4
27. Brynger, H. Twenty-Four-Hour Preservation of the Duct-Ligated Canine Pancreatic Allograft. Eur Surg Res (2008) 7(6):341–54. doi:10.1159/000127819
28. Steen, S, Sjöberg, T, Pierre, L, Liao, Q, Eriksson, L, and Algotsson, L. Transplantation of Lungs From a Non-heart-beating Donor. Lancet Lond Engl (2001) 357(9259):825–9. doi:10.1016/S0140-6736(00)04195-7
29. Steen, S, Liao, Q, Wierup, PN, Bolys, R, Pierre, L, and Sjöberg, T. Transplantation of Lungs From Non-heart-beating Donors After Functional Assessment Ex Vivo. Ann Thorac Surg (2003) 76(1):244–52. discussion 252. doi:10.1016/s0003-4975(03)00191-7
30. Steen, S, Ingemansson, R, Eriksson, L, Pierre, L, Algotsson, L, Wierup, P, et al. First Human Transplantation of a Nonacceptable Donor Lung After Reconditioning Ex Vivo. Ann Thorac Surg (2007) 83(6):2191–4. doi:10.1016/j.athoracsur.2007.01.033
31. Cypel, M, Yeung, JC, Liu, M, Anraku, M, Chen, F, Karolak, W, et al. Normothermic Ex Vivo Lung Perfusion in Clinical Lung Transplantation. N Engl J Med (2011) 364(15):1431–40. doi:10.1056/NEJMoa1014597
32. Luc, JGY, Aboelnazar, NS, Himmat, S, Hatami, S, Haromy, A, Matsumura, N, et al. A Leukocyte Filter Does Not Provide Further Benefit During Ex Vivo Lung Perfusion. ASAIO J Am Soc Artif Intern Organs (2017) 63(5):672–8. doi:10.1097/MAT.0000000000000550
33. mkmedia.se. XVIVO. Available from: https://www.xvivoperfusion.com/(Accessed March 26, 2024).
34. Home - Transmedics. (Accessed March 26, 2024). Available from: https://www.transmedics.com/
35. Cypel, M, Yeung, JC, Hirayama, S, Rubacha, M, Fischer, S, Anraku, M, et al. Technique for Prolonged Normothermic Ex Vivo Lung Perfusion. J Heart Lung Transpl Off Publ Int Soc Heart Transpl (2008) 27(12):1319–25. doi:10.1016/j.healun.2008.09.003
36. Loor, G, Warnecke, G, Villavicencio, MA, Smith, MA, Kukreja, J, Ardehali, A, et al. Portable Normothermic Ex-Vivo Lung Perfusion, Ventilation, and Functional Assessment With the Organ Care System on Donor Lung Use for Transplantation From Extended-Criteria Donors (EXPAND): A Single-Arm, Pivotal Trial. Lancet Respir Med (2019) 7(11):975–84. doi:10.1016/S2213-2600(19)30200-0
37. Sanchez, PG, Davis, RD, D’Ovidio, F, Cantu, E, Weyant, M, Camp, P, et al. The NOVEL Lung Trial One-Year Outcomes. J Heart Lung Transpl (2014) 33(4):S71–2. doi:10.1016/j.healun.2014.01.226
38. Fisher, A, Andreasson, A, Chrysos, A, Lally, J, Mamasoula, C, Exley, C, et al. An Observational Study of Donor Ex Vivo Lung Perfusion in UK Lung Transplantation: DEVELOP-UK. Health Technol Assess Winch Engl (2016) 20(85):1–276. doi:10.3310/hta20850
39. Warnecke, G, Van Raemdonck, D, Smith, MA, Massard, G, Kukreja, J, Rea, F, et al. Normothermic Ex-Vivo Preservation With the Portable Organ Care System Lung Device for Bilateral Lung Transplantation (INSPIRE): A Randomised, Open-Label, Non-Inferiority, Phase 3 Study. Lancet Respir Med (2018) 6(5):357–67. doi:10.1016/S2213-2600(18)30136-X
40. Slama, A, Schillab, L, Barta, M, Benedek, A, Mitterbauer, A, Hoetzenecker, K, et al. Standard Donor Lung Procurement With Normothermic Ex Vivo Lung Perfusion: A Prospective Randomized Clinical Trial. J Heart Lung Transpl Off Publ Int Soc Heart Transpl (2017) 36(7):744–53. doi:10.1016/j.healun.2017.02.011
41. Mallea, JM, Hartwig, MG, Keller, CA, Kon, Z, Iii, RNP, Erasmus, DB, et al. Remote Ex Vivo Lung Perfusion at a Centralized Evaluation Facility. J Heart Lung Transpl Off Publ Int Soc Heart Transpl (2022) 41(12):1700–11. doi:10.1016/j.healun.2022.09.006
42. Bridge-Lungsforlife. THE BOARD | The Bridge - Lungs for Life. Available from: https://www.bridge-lungsforlife.com/board (Accessed March 26, 2024).
43. Grasso, S, Terragni, P, Mascia, L, Fanelli, V, Quintel, M, Herrmann, P, et al. Airway Pressure-Time Curve Profile (Stress Index) Detects Tidal Recruitment/hyperinflation in Experimental Acute Lung Injury. Crit Care Med (2004) 32(4):1018–27. doi:10.1097/01.ccm.0000120059.94009.ad
44. Terragni, PP, Filippini, C, Slutsky, AS, Birocco, A, Tenaglia, T, Grasso, S, et al. Accuracy of Plateau Pressure and Stress Index to Identify Injurious Ventilation in Patients With Acute Respiratory Distress Syndrome. Anesthesiology (2013) 119(4):880–9. doi:10.1097/ALN.0b013e3182a05bb8
45. Terragni, PP, Fanelli, V, Boffini, M, Filippini, C, Cappello, P, Ricci, D, et al. Ventilatory Management During Normothermic Ex Vivo Lung Perfusion: Effects on Clinical Outcomes. Transplantation (2016) 100(5):1128–35. doi:10.1097/TP.0000000000000929
46. Broberg, E, Wlosinska, M, Algotsson, L, Olin, AC, Wagner, D, Pierre, L, et al. A New Way of Monitoring Mechanical Ventilation by Measurement of Particle Flow From the Airways Using Pexa Method In Vivo and during Ex Vivo Lung Perfusion in DCD Lung Transplantation. Intensive Care Med Exp (2018) 6(1):18. doi:10.1186/s40635-018-0188-z
47. Stenlo, M, Hyllén, S, Silva, IAN, Bölükbas, DA, Pierre, L, Hallgren, O, et al. Increased Particle Flow Rate from Airways Precedes Clinical Signs of ARDS in a Porcine Model of LPS-Induced Acute Lung Injury. Am J Physiol Lung Cell Mol Physiol (2020) 318(3):L510-L517–7. doi:10.1152/ajplung.00524.2019
48. Stenlo, M, Silva, IAN, Hyllén, S, Bölükbas, DA, Niroomand, A, Grins, E, et al. Monitoring Lung Injury With Particle Flow Rate in LPS- and COVID-19-Induced ARDS. Physiol Rep (2021) 9(13):e14802. doi:10.14814/phy2.14802
49. Broberg, E, Hyllén, S, Algotsson, L, Wagner, DE, and Lindstedt, S. Particle Flow Profiles From the Airways Measured by PExA Differ in Lung Transplant Recipients Who Develop Primary Graft Dysfunction. Exp Clin Transpl Off J Middle East Soc Organ Transpl (2019) 17(6):803–12. doi:10.6002/ect.2019.0187
50. Mehaffey, JH, Charles, EJ, Sharma, AK, Money, DT, Zhao, Y, Stoler, MH, et al. Airway Pressure Release Ventilation during Ex Vivo Lung Perfusion Attenuates Injury. J Thorac Cardiovasc Surg (2017) 153(1):197–204. doi:10.1016/j.jtcvs.2016.09.029
51. Aboelnazar, NS, Himmat, S, Hatami, S, White, CW, Burhani, MS, Dromparis, P, et al. Negative Pressure Ventilation Decreases Inflammation and Lung Edema During Normothermic Ex-Vivo Lung Perfusion. J Heart Lung Transpl Off Publ Int Soc Heart Transpl (2018) 37(4):520–30. doi:10.1016/j.healun.2017.09.007
52. Tsui, S, Dhital, K, Eisenring, C, and Ardehali, A. 387: Ex-Vivo Assessment of Donor Hearts With the OCS to Detect Hidden Pathologies. J Heart Lung Transpl (2008) 27(2):S200. doi:10.1016/j.healun.2007.11.398
53. Hamed, A, Tsui, S, Huber, J, Lin, R, Poggio, EC, and Ardehali, A. 19: Serum Lactate Is a Highly Sensitive and Specific Predictor of Post Cardiac Transplant Outcomes Using the Organ Care System. J Heart Lung Transpl (2009) 28(2):S71. doi:10.1016/j.healun.2008.11.025
54. Koerner, MM, Ghodsizad, A, Schulz, U, Banayosy, AE, Koerfer, R, and Tenderich, G. Normothermic Ex Vivo Allograft Blood Perfusion in Clinical Heart Transplantation. Heart Surg Forum (2014) 17(3):E141–5. doi:10.1532/HSF98.2014332
55. Tenderich, G, Tsui, S, El-Banayosy, A, Dhital, K, Schulte-Eistrup, S, Schulz, U, et al. 293: The 1-Year Follow-Up Results of the PROTECT Patient Population Using the Organ Care System. J Heart Lung Transpl (2008) 27(2):S166. doi:10.1016/j.healun.2007.11.302
56. Ardehali, A, Esmailian, F, Deng, M, Soltesz, E, Hsich, E, Naka, Y, et al. Ex-vivo Perfusion of Donor Hearts for Human Heart Transplantation (PROCEED II): A Prospective, Open-Label, Multicentre, Randomised Non-Inferiority Trial. The Lancet (2015) 385(9987):2577–84. doi:10.1016/S0140-6736(15)60261-6
57. Chan, JL, Kobashigawa, JA, Reich, HJ, Ramzy, D, Thottam, MM, Yu, Z, et al. Intermediate Outcomes With Ex-Vivo Allograft Perfusion for Heart Transplantation. J Heart Lung Transpl Off Publ Int Soc Heart Transpl (2017) 36(3):258–63. doi:10.1016/j.healun.2016.08.015
58. Schroder, JN, D’Alessandro, D, Esmailian, F, Boeve, T, Tang, P, Liao, K, et al. Successful Utilization of Extended Criteria Donor (ECD) Hearts for Transplantation - Results of the OCSTM Heart EXPAND Trial to Evaluate the Effectiveness and Safety of the OCS Heart System to Preserve and Assess ECD Hearts for Transplantation. J Heart Lung Transpl (2019) 38(4):S42. doi:10.1016/j.healun.2019.01.088
59. García Sáez, D, Zych, B, Sabashnikov, A, Bowles, CT, De Robertis, F, Mohite, PN, et al. Evaluation of the Organ Care System in Heart Transplantation With an Adverse Donor/Recipient Profile. Ann Thorac Surg (2014) 98(6):2099–106. doi:10.1016/j.athoracsur.2014.06.098
60. Sáez, DG, Zych, B, Mohite, PN, Sabashnikov, A, Patil, NP, Popov, A, et al. LVAD Bridging to Heart Transplantation With Ex Vivo Allograft Preservation Shows Significantly Improved: Outcomes: A New Standard of Care? J Heart Lung Transpl (2015) 34(4):S95. doi:10.1016/j.healun.2015.01.252
61. Real World Experience with Transmedics Organ Care System in Cardiac Transplantation With Donor Organs Associated With Marginal Risk Factors - UQ eSpace. (Accessed March 26, 2024). Available from: https://espace.library.uq.edu.au/view/UQ:cab11e1
62. Yeter, R, Pasic, M, Hübler, M, Dandel, M, Hiemann, N, Kemper, D, et al. Extended Donor Criteria in Heart Transplantation: 4-Year Results of the Experience with the Organ Care System. Thorac Cardiovasc Surg (2014) 62(S 01):SC44. doi:10.1055/s-0034-1367305
63. Stamp, NL, Shah, A, Vincent, V, Wright, B, Wood, C, Pavey, W, et al. Successful Heart Transplant After Ten Hours Out-Of-Body Time Using the TransMedics Organ Care System. Heart Lung Circ (2015) 24(6):611–3. doi:10.1016/j.hlc.2015.01.005
64. Dhital, KK, Iyer, A, Connellan, M, Chew, HC, Gao, L, Doyle, A, et al. Adult Heart Transplantation With Distant Procurement and Ex-Vivo Preservation of Donor Hearts After Circulatory Death: A Case Series. Lancet Lond Engl (2015) 385(9987):2585–91. doi:10.1016/S0140-6736(15)60038-1
65. Iyer, A, Gao, L, Doyle, A, Rao, P, Cropper, JR, Soto, C, et al. Normothermic Ex Vivo Perfusion Provides Superior Organ Preservation and Enables Viability Assessment of Hearts From DCD Donors. Am J Transpl Off J Am Soc Transpl Am Soc Transpl Surg (2015) 15(2):371–80. doi:10.1111/ajt.12994
66. Moeslund, N, Ertugrul, IA, Hu, MA, Dalsgaard, FF, Ilkjaer, LB, Ryhammer, P, et al. Ex-Situ Oxygenated Hypothermic Machine Perfusion in Donation After Circulatory Death Heart Transplantation Following Either Direct Procurement or In-Situ Normothermic Regional Perfusion. J Heart Lung Transpl (2023) 42(6):730–40. doi:10.1016/j.healun.2023.01.014
67. Emmanuel, S, MacDonald, P, Hayward, C, Watson, A, Iyer, A, Connellan, M, et al. Initial Australian Experience With the Xvivo Non-Ischaemic Hypothermic Perfusion Device for Heart Preservation. J Heart Lung Transpl (2023) 42(4):S61–2. 117.
68. Nilsson, J, Jernryd, V, Qin, G, Paskevicius, A, Metzsch, C, Sjöberg, T, et al. A Nonrandomized Open-Label Phase 2 Trial of Nonischemic Heart Preservation for Human Heart Transplantation. Nat Commun (2020) 11(1):2976. doi:10.1038/s41467-020-16782-9
69. McGiffin, DC, Kure, CE, Macdonald, PS, Jansz, PC, Emmanuel, S, Marasco, SF, et al. Hypothermic Oxygenated Perfusion (HOPE) Safely and Effectively Extends Acceptable Donor Heart Preservation Times: Results of the Australian and New Zealand Trial. J Heart Lung Transpl Off Publ Int Soc Heart Transpl (2024) 43(3):485–95. doi:10.1016/j.healun.2023.10.020
70. Nasralla, D, Coussios, CC, Mergental, H, Akhtar, MZ, Butler, AJ, Ceresa, CDL, et al. A Randomized Trial of Normothermic Preservation in Liver Transplantation. Nature (2018) 557(7703):50–6. doi:10.1038/s41586-018-0047-9
71. Southard, JH, van Gulik, TM, Ametani, MS, Vreugdenhil, PK, Lindell, SL, Pienaar, BL, et al. Important Components of the UW Solution. Transplantation (1990) 49(2):251–7. doi:10.1097/00007890-199002000-00004
72. Jakubauskas, M, Jakubauskiene, L, Leber, B, Strupas, K, Stiegler, P, and Schemmer, P. Machine Perfusion in Liver Transplantation: A Systematic Review and Meta-Analysis. Visc Med (2021) 38(4):243–54. doi:10.1159/000519788
73. Moers, C, Smits, JM, Maathuis, MHJ, Treckmann, J, van Gelder, F, Napieralski, BP, et al. Machine Perfusion or Cold Storage in Deceased-Donor Kidney Transplantation. N Engl J Med (2009) 360(1):7–19. doi:10.1056/NEJMoa0802289
74. Barlow, AD, Hamed, MO, Mallon, DH, Brais, RJ, Gribble, FM, Scott, MA, et al. Use of Ex Vivo Normothermic Perfusion for Quality Assessment of Discarded Human Donor Pancreases. Am J Transpl Off J Am Soc Transpl Am Soc Transpl Surg (2015) 15(9):2475–82. doi:10.1111/ajt.13303
75. Hamaoui, K, and Papalois, V. Machine Perfusion and the Pancreas: Will It Increase the Donor Pool? Curr Diab Rep (2019) 19(8):56. doi:10.1007/s11892-019-1165-y
76. Gojo, S, Niwaya, K, Taniguchi, S, Nishizaki, K, and Kitamura, S. Gene Transfer into the Donor Heart During Cold Preservation for Heart Transplantation. Ann Thorac Surg (1998) 65(3):647–52. doi:10.1016/s0003-4975(97)01295-2
77. Shiraishi, M, Kusano, T, Hara, J, Hiroyasu, S, Shao-Ping, M, Makino, Y, et al. Adenovirus-Mediated Gene Transfer Using Ex Vivo Perfusion of the Heart Graft. Surg Today (1996) 26(8):624–8. doi:10.1007/BF00311668
78. Pellegrini, C, Jeppsson, A, Taner, CB, O’Brien, T, Miller, VM, Tazelaar, HD, et al. Highly Efficient Ex Vivo Gene Transfer to the Transplanted Heart by Means of Hypothermic Perfusion With a Low Dose of Adenoviral Vector. J Thorac Cardiovasc Surg (2000) 119(3):493–500. doi:10.1016/s0022-5223(00)70128-0
79. Rao, VP, Branzoli, SE, Ricci, D, Miyagi, N, O’Brien, T, Tazelaar, HD, et al. Recombinant Adenoviral Gene Transfer Does Not Affect Cardiac Allograft Vasculopathy. J Heart Lung Transpl Off Publ Int Soc Heart Transpl (2007) 26(12):1281–5. doi:10.1016/j.healun.2007.09.018
80. Oi, K, Davies, WR, Tazelaar, HD, Bailey, KR, Federspiel, MJ, Russell, SJ, et al. Ex vivo Hypothermic Recirculatory Adenoviral Gene Transfer to the Transplanted Pig Heart. J Gene Med (2006) 8(7):795–803. doi:10.1002/jgm.913
81. Donahue, JK, Kikkawa, K, Thomas, AD, Marban, E, and Lawrence, JH. Acceleration of Widespread Adenoviral Gene Transfer to Intact Rabbit Hearts by Coronary Perfusion With Low Calcium and Serotonin. Gene Ther (1998) 5(5):630–4. doi:10.1038/sj.gt.3300649
82. Nagata, K, Marbán, E, Lawrence, JH, and Donahue, JK. Phosphodiesterase Inhibitor-Mediated Potentiation of Adenovirus Delivery to Myocardium. J Mol Cell Cardiol (2001) 33(3):575–80. doi:10.1006/jmcc.2000.1322
83. Bishawi, M, Roan, JN, Milano, CA, Daneshmand, MA, Schroder, JN, Chiang, Y, et al. A Normothermic Ex Vivo Organ Perfusion Delivery Method for Cardiac Transplantation Gene Therapy. Sci Rep (2019) 9(1):8029. doi:10.1038/s41598-019-43737-y
84. Mendiola Pla, M, Chiang, Y, Roki, A, Wang, C, Lee, FH, Smith, MF, et al. Ex Vivo Gene Delivery to Porcine Cardiac Allografts Using a Myocardial-Enhanced Adeno-Associated Viral Vector. Hum Gene Ther (2023) 34(7–8):303–13. doi:10.1089/hum.2022.241
85. Brauner, R, Wu, L, Laks, H, Nonoyama, M, Scholl, F, Shvarts, O, et al. Intracoronary Gene Transfer of Immunosuppressive Cytokines to Cardiac Allografts: Method and Efficacy of Adenovirus-Mediated Transduction. J Thorac Cardiovasc Surg (1997) 113(6):1059–66. discussion 1066-1067. doi:10.1016/S0022-5223(97)70293-9
86. Brauner, R, Nonoyama, M, Laks, H, Drinkwater, DC, McCaffery, S, Drake, T, et al. Intracoronary Adenovirus-Mediated Transfer of Immunosuppressive Cytokine Genes Prolongs Allograft Survival. J Thorac Cardiovasc Surg (1997) 114(6):923–33. doi:10.1016/S0022-5223(97)70006-0
87. Yang, Z, Rostami, S, Koeberlein, B, Barker, CF, and Naji, A. Cardiac Allograft Tolerance Induced by Intra-Arterial Infusion of Recombinant Adenoviral CTLA4Ig. Transplantation (1999) 67(12):1517–23. doi:10.1097/00007890-199906270-00004
88. Furukawa, H, Oshima, K, Tung, T, Cui, G, Laks, H, and Sen, L. Liposome-Mediated Combinatorial Cytokine Gene Therapy Induces Localized Synergistic Immunosuppression and Promotes Long-Term Survival of Cardiac Allografts. J Immunol Baltim Md (2005) 174(11):6983–92. doi:10.4049/jimmunol.174.11.6983
89. Kanaan, SA, Kozower, BD, Suda, T, Daddi, N, Tagawa, T, Ritter, JH, et al. Intratracheal Adenovirus-Mediated Gene Transfer Is Optimal in Experimental Lung Transplantation. J Thorac Cardiovasc Surg (2002) 124(6):1130–6. doi:10.1067/mtc.2002.123702
90. Chapelier, A, Danel, C, Mazmanian, M, Bacha, EA, Sellak, H, Gilbert, MA, et al. Gene Therapy in Lung Transplantation: Feasibility of Ex Vivo Adenovirus-Mediated Gene Transfer to the Graft. Hum Gene Ther (1996) 7(15):1837–45. doi:10.1089/hum.1996.7.15-1837
91. Cassivi, SD, Cardella, JA, Fischer, S, Liu, M, Slutsky, AS, and Keshavjee, S. Transtracheal Gene Transfection of Donor Lungs Prior to Organ Procurement Increases Transgene Levels at Reperfusion and Following Transplantation. J Heart Lung Transpl Off Publ Int Soc Heart Transpl (1999) 18(12):1181–8. doi:10.1016/s1053-2498(99)00095-9
92. de Perrot, M, Fischer, S, Liu, M, Imai, Y, Martins, S, Sakiyama, S, et al. Impact of Human Interleukin-10 on Vector-Induced Inflammation and Early Graft Function in Rat Lung Transplantation. Am J Respir Cell Mol Biol (2003) 28(5):616–25. doi:10.1165/rcmb.2002-0109OC
93. Cypel, M, Liu, M, Rubacha, M, Yeung, JC, Hirayama, S, Anraku, M, et al. Functional Repair of Human Donor Lungs by IL-10 Gene Therapy. Sci Transl Med (2009) 1(4):4ra9. doi:10.1126/scitranslmed.3000266
94. Yeung, JC, Wagnetz, D, Cypel, M, Rubacha, M, Koike, T, Chun, YM, et al. Ex Vivo Adenoviral Vector Gene Delivery Results in Decreased Vector-Associated Inflammation Pre- and Post–Lung Transplantation in the Pig. Mol Ther (2012) 20(6):1204–11. doi:10.1038/mt.2012.57
95. Machuca, TN, Cypel, M, Bonato, R, Yeung, JC, Chun, YM, Juvet, S, et al. Safety and Efficacy of Ex Vivo Donor Lung Adenoviral IL-10 Gene Therapy in a Large Animal Lung Transplant Survival Model. Hum Gene Ther (2017) 28(9):757–65. doi:10.1089/hum.2016.070
96. Oishi, H, Okada, Y, Kikuchi, T, Hoshikawa, Y, Sado, T, Noda, M, et al. Transbronchial Human Interleukin-10 Gene Transfer Reduces Acute Inflammation Associated With Allograft Rejection and Intragraft Interleukin-2 and Tumor Necrosis Factor-α Gene Expression in a Rat Model of Lung Transplantation. J Heart Lung Transpl (2010) 29(3):360–7. doi:10.1016/j.healun.2009.10.002
97. Itano, H, Zhang, W, Ritter, JH, McCarthy, TJ, Yew, NS, Mohnanakumar, T, et al. Endobronchial Transfection of Naked Viral Interleukin-10 Gene in Rat Lung Allotransplantation. Ann Thorac Surg (2001) 71(4):1126–33. doi:10.1016/s0003-4975(00)02651-5
98. Itano, H, Mora, BN, Zhang, W, Ritter, JH, McCarthy, TJ, Yew, NS, et al. Lipid-Mediated Ex Vivo Gene Transfer of Viral Interleukin 10 in Rat Lung Allotransplantation. J Thorac Cardiovasc Surg (2001) 122(1):29–38. doi:10.1067/mtc.2001.114636
99. Figueiredo, C, Carvalho Oliveira, M, Chen-Wacker, C, Jansson, K, Höffler, K, Yuzefovych, Y, et al. Immunoengineering of the Vascular Endothelium to Silence MHC Expression during Normothermic Ex Vivo Lung Perfusion. Hum Gene Ther (2019) 30(4):485–96. doi:10.1089/hum.2018.117
100. Bogacz, A, Polaszewska, A, Bartkowiak-Wieczorek, J, Tejchman, K, Dziewanowski, K, Ostrowski, M, et al. The Effect of Genetic Variations for Interleukin-10 (IL-10) on the Efficacy of Immunosuppressive Therapy in Patients after Kidney Transplantation. Int Immunopharmacol (2020) 89:107059. doi:10.1016/j.intimp.2020.107059
101. Jang, JY, Kim, HW, Yan, J, Kang, TK, Lee, W, Kim, BS, et al. Interleukin-2/Anti-Interleukin-2 Immune Complex Attenuates Cold Ischemia-Reperfusion Injury After Kidney Transplantation by Increasing Renal Regulatory T Cells. Clin Transl Med (2024) 14(3):e1631. doi:10.1002/ctm2.1631
102. Lee, M, Kim, MJ, Oh, J, Piao, C, Park, YW, and Lee, DY. Gene Delivery to Pancreatic Islets for Effective Transplantation in Diabetic Animal. J Ind Eng Chem (2017) 56:45–54. doi:10.1016/j.jiec.2017.07.038
103. Nguyen, TH, Mainot, S, Lainas, P, Groyer-Picard, MT, Franco, D, Dagher, I, et al. Ex Vivo Liver-Directed Gene Therapy for the Treatment of Metabolic Diseases: Advances in Hepatocyte Transplantation and Retroviral Vectors. Curr Gene Ther (2009) 9(2):136–49. doi:10.2174/156652309787909481
104. Figueiredo, C, Oldhafer, F, Wittauer, EM, Carvalho-Oliveira, M, Akhdar, A, Beetz, O, et al. Silencing of HLA Class I on Primary Human Hepatocytes as a Novel Strategy for Reduction in Alloreactivity. J Cell Mol Med (2019) 23(8):5705–14. doi:10.1111/jcmm.14484
105. Goldaracena, N, Spetzler, VN, Echeverri, J, Kaths, JM, Cherepanov, V, Persson, R, et al. Inducing Hepatitis C Virus Resistance After Pig Liver Transplantation-A Proof of Concept of Liver Graft Modification Using Warm Ex Vivo Perfusion. Am J Transpl Off J Am Soc Transpl Am Soc Transpl Surg (2017) 17(4):970–8. doi:10.1111/ajt.14100
106. Lorvellec, M, Pellegata, AF, Maestri, A, Turchetta, C, Alvarez Mediavilla, E, Shibuya, S, et al. An In Vitro Whole-Organ Liver Engineering for Testing of Genetic Therapies. iScience (2020) 23(12):101808. doi:10.1016/j.isci.2020.101808
107. Gene Therapy Needs a Long-Term Approach. Nat Med. 2021;27(4):563. doi:10.1038/s41591-021-01333-6
108. Islam, MN, Das, SR, Emin, MT, Wei, M, Sun, L, Westphalen, K, et al. Mitochondrial Transfer From Bone-Marrow-Derived Stromal Cells to Pulmonary Alveoli Protects against Acute Lung Injury. Nat Med (2012) 18(5):759–65. doi:10.1038/nm.2736
109. Lee, JW, Krasnodembskaya, A, McKenna, DH, Song, Y, Abbott, J, and Matthay, MA. Therapeutic Effects of Human Mesenchymal Stem Cells in Ex Vivo Human Lungs Injured With Live Bacteria. Am J Respir Crit Care Med (2013) 187(7):751–60. doi:10.1164/rccm.201206-0990OC
110. Ryan, JM, Barry, FP, Murphy, JM, and Mahon, BP. Mesenchymal Stem Cells Avoid Allogeneic Rejection. J Inflamm Lond Engl (2005) 2:8. doi:10.1186/1476-9255-2-8
111. Lee, JW, Fang, X, Gupta, N, Serikov, V, and Matthay, MA. Allogeneic Human Mesenchymal Stem Cells for Treatment of E. coli Endotoxin-Induced Acute Lung Injury in the Ex Vivo Perfused Human Lung. Proc Natl Acad Sci U S A (2009) 106(38):16357–62. doi:10.1073/pnas.0907996106
112. McAuley, DF, Curley, GF, Hamid, UI, Laffey, JG, Abbott, J, McKenna, DH, et al. Clinical Grade Allogeneic Human Mesenchymal Stem Cells Restore Alveolar Fluid Clearance in Human Lungs Rejected for Transplantation. Am J Physiol Lung Cell Mol Physiol (2014) 306(9):L809–15. doi:10.1152/ajplung.00358.2013
113. Mordant, P, Nakajima, D, Kalaf, R, Iskender, I, Maahs, L, Behrens, P, et al. Mesenchymal Stem Cell Treatment Is Associated With Decreased Perfusate Concentration of Interleukin-8 During Ex Vivo Perfusion of Donor Lungs After 18-Hour Preservation. J Heart Lung Transpl Off Publ Int Soc Heart Transpl (2016) 35(10):1245–54. doi:10.1016/j.healun.2016.04.017
114. Nakajima, D, Watanabe, Y, Ohsumi, A, Pipkin, M, Chen, M, Mordant, P, et al. Mesenchymal Stromal Cell Therapy during Ex Vivo Lung Perfusion Ameliorates Ischemia-Reperfusion Injury in Lung Transplantation. J Heart Lung Transpl Off Publ Int Soc Heart Transpl (2019) 38(11):1214–23. doi:10.1016/j.healun.2019.07.006
115. Nykänen, AI, Mariscal, A, Duong, A, Estrada, C, Ali, A, Hough, O, et al. Engineered Mesenchymal Stromal Cell Therapy During Human Lung Ex Vivo Lung Perfusion Is Compromised by Acidic Lung Microenvironment. Mol Ther Methods Clin Dev (2021) 23:184–97. doi:10.1016/j.omtm.2021.05.018
116. Pacienza, N, Santa-Cruz, D, Malvicini, R, Robledo, O, Lemus-Larralde, G, Bertolotti, A, et al. Mesenchymal Stem Cell Therapy Facilitates Donor Lung Preservation by Reducing Oxidative Damage During Ischemia. Stem Cells Int. (2019) 2019:8089215. doi:10.1155/2019/8089215
117. Korkmaz-Icöz, S, Li, S, Hüttner, R, Ruppert, M, Radovits, T, Loganathan, S, et al. Hypothermic Perfusion of Donor Heart With a Preservation Solution Supplemented by Mesenchymal Stem Cells. J Heart Lung Transpl Off Publ Int Soc Heart Transpl (2019) 38(3):315–26. doi:10.1016/j.healun.2018.12.003
118. Korkmaz-Icöz, S, Li, K, Loganathan, S, Ding, Q, Ruppert, M, Radovits, T, et al. Brain-Dead Donor Heart Conservation With a Preservation Solution Supplemented by a Conditioned Medium From Mesenchymal Stem Cells Improves Graft Contractility After Transplantation. Am J Transpl Off J Am Soc Transpl Am Soc Transpl Surg (2020) 20(10):2847–56. doi:10.1111/ajt.15843
119. Zeng, Z, Xu, L, Xu, Y, Ruan, Y, Liu, D, Li, J, et al. Normothermic Ex Vivo Heart Perfusion With Mesenchymal Stem Cell-Derived Conditioned Medium Improves Myocardial Tissue Protection in Rat Donation After Circulatory Death Hearts. Stem Cells Int. (2022) 2022:8513812. doi:10.1155/2022/8513812
120. Zhou, P, Liu, H, Liu, X, Ling, X, Xiao, Z, Zhu, P, et al. Donor Heart Preservation With Hypoxic-Conditioned Medium-Derived From Bone Marrow Mesenchymal Stem Cells Improves Cardiac Function in a Heart Transplantation Model. Stem Cell Res Ther (2021) 12(1):56. doi:10.1186/s13287-020-02114-7
121. Bartunek, J, Terzic, A, Davison, BA, Filippatos, GS, Radovanovic, S, Beleslin, B, et al. Cardiopoietic Cell Therapy for Advanced Ischaemic Heart Failure: Results at 39 Weeks of the Prospective, Randomized, Double Blind, Sham-Controlled CHART-1 Clinical Trial. Eur Heart J (2017) 38(9):648–60. doi:10.1093/eurheartj/ehw543
122. Butler, J, Epstein, SE, Greene, SJ, Quyyumi, AA, Sikora, S, Kim, RJ, et al. Intravenous Allogeneic Mesenchymal Stem Cells for Nonischemic Cardiomyopathy. Circ Res (2017) 120(2):332–40. doi:10.1161/CIRCRESAHA.116.30971
123. Menasché, P, Vanneaux, V, Hagège, A, Bel, A, Cholley, B, Parouchev, A, et al. Transplantation of Human Embryonic Stem Cell-Derived Cardiovascular Progenitors for Severe Ischemic Left Ventricular Dysfunction. J Am Coll Cardiol (2018) 71(4):429–38. doi:10.1016/j.jacc.2017.11.047
124. Hare, JM, DiFede, DL, Rieger, AC, Florea, V, Landin, AM, El-Khorazaty, J, et al. Randomized Comparison of Allogeneic versus Autologous Mesenchymal Stem Cells for Nonischemic Dilated Cardiomyopathy: POSEIDON-DCM Trial. J Am Coll Cardiol (2017) 69(5):526–37. doi:10.1016/j.jacc.2016.11.009
125. Xiao, W, Guo, S, Gao, C, Dai, G, Gao, Y, Li, M, et al. A Randomized Comparative Study on the Efficacy of Intracoronary Infusion of Autologous Bone Marrow Mononuclear Cells and Mesenchymal Stem Cells in Patients With Dilated Cardiomyopathy. Int Heart J (2017) 58(2):238–44. doi:10.1536/ihj.16-328
126. Florea, V, Rieger, AC, DiFede, DL, El-Khorazaty, J, Natsumeda, M, Banerjee, MN, et al. Dose Comparison Study of Allogeneic Mesenchymal Stem Cells in Patients With Ischemic Cardiomyopathy (The TRIDENT Study). Circ Res (2017) 121(11):1279–90. doi:10.1161/CIRCRESAHA.117.311827
127. Huang, A, Liu, Y, Qi, X, Chen, S, Huang, H, Zhang, J, et al. Intravenously Transplanted Mesenchymal Stromal Cells: A New Endocrine Reservoir for Cardioprotection. Stem Cell Res Ther (2022) 13(1):253. doi:10.1186/s13287-022-02922-z
128. Brasile, L, Henry, N, Orlando, G, and Stubenitsky, B. Potentiating Renal Regeneration Using Mesenchymal Stem Cells. Transplantation (2019) 103(2):307–13. doi:10.1097/TP.0000000000002455
129. Gregorini, M, Corradetti, V, Pattonieri, EF, Rocca, C, Milanesi, S, Peloso, A, et al. Perfusion of Isolated Rat Kidney with Mesenchymal Stromal Cells/Extracellular Vesicles Prevents Ischaemic Injury. J Cell Mol Med (2017) 21(12):3381–93. doi:10.1111/jcmm.13249
130. Pool, M, Eertman, T, Sierra Parraga, J, ’t Hart, N, Roemeling-van Rhijn, M, Eijken, M, et al. Infusing Mesenchymal Stromal Cells Into Porcine Kidneys during Normothermic Machine Perfusion: Intact MSCs Can Be Traced and Localised to Glomeruli. Int J Mol Sci (2019) 20(14):3607. doi:10.3390/ijms20143607
131. Pool, MBF, Vos, J, Eijken, M, van Pel, M, Reinders, MEJ, Ploeg, RJ, et al. Treating Ischemically Damaged Porcine Kidneys With Human Bone Marrow- and Adipose Tissue-Derived Mesenchymal Stromal Cells During Ex Vivo Normothermic Machine Perfusion. Stem Cells Dev (2020) 29(20):1320–30. doi:10.1089/scd.2020.0024
132. Lohmann, S, Pool, MBF, Rozenberg, KM, Keller, AK, Moers, C, Møldrup, U, et al. Mesenchymal Stromal Cell Treatment of Donor Kidneys During Ex Vivo Normothermic Machine Perfusion: A Porcine Renal Autotransplantation Study. Am J Transpl Off J Am Soc Transpl Am Soc Transpl Surg (2021) 21(7):2348–59. doi:10.1111/ajt.16473
133. Munk, A, Duvald, CS, Pedersen, M, Lohmann, S, Keller, AK, Møller, BK, et al. Dosing Limitation for Intra-Renal Arterial Infusion of Mesenchymal Stromal Cells. Int J Mol Sci (2022) 23(15):8268. doi:10.3390/ijms23158268
134. Sasajima, H, Miyagi, S, Kakizaki, Y, Kamei, T, Unno, M, Satomi, S, et al. Cytoprotective Effects of Mesenchymal Stem Cells During Liver Transplantation From Donors After Cardiac Death in Rats. Transpl Proc (2018) 50(9):2815–20. doi:10.1016/j.transproceed.2018.02.180
135. Yang, L, Cao, H, Sun, D, Lin, L, Zheng, WP, Shen, ZY, et al. Normothermic Machine Perfusion Combined with Bone Marrow Mesenchymal Stem Cells Improves the Oxidative Stress Response and Mitochondrial Function in Rat Donation After Circulatory Death Livers. Stem Cells Dev. (2020) 29(13):835–52. doi:10.1089/scd.2019.0301
136. Yang, L, Cao, H, Sun, D, Hou, B, Lin, L, Shen, ZY, et al. Bone Marrow Mesenchymal Stem Cells Combine With Normothermic Machine Perfusion to Improve Rat Donor Liver Quality-The Important Role of Hepatic Microcirculation in Donation After Circulatory Death. Cell Tissue Res (2020) 381(2):239–54. doi:10.1007/s00441-020-03202-z
137. Verstegen, MMA, Mezzanotte, L, Ridwan, RY, Wang, K, de Haan, J, Schurink, IJ, et al. First Report on Ex Vivo Delivery of Paracrine Active Human Mesenchymal Stromal Cells to Liver Grafts During Machine Perfusion. Transplantation (2020) 104(1):e5–7. doi:10.1097/TP.0000000000002986
138. O’Neill, JD, Guenthart, BA, Kim, J, Chicotka, S, Queen, D, Fung, K, et al. Cross-circulation for Extracorporeal Support and Recovery of the Lung. Nat Biomed Eng (2017) 1(3):0037–15. doi:10.1038/s41551-017-0037
139. Egan, TM, Saunders, NR, Dubois, P, Choiniere, L, McDonald, JW, and Cooper, JD. Contribution of Circulating Formed Elements to Prostanoid Production in Complement-Mediated Lung Injury in Sheep. Surgery (1985) 98(2):350–8.
140. Guenthart, BA, O’Neill, JD, Kim, J, Queen, D, Chicotka, S, Fung, K, et al. Regeneration of Severely Damaged Lungs Using an Interventional Cross-Circulation Platform. Nat Commun (2019) 10(1):1985. doi:10.1038/s41467-019-09908-1
141. Hozain, AE, Tipograf, Y, Pinezich, MR, Cunningham, KM, Donocoff, R, Queen, D, et al. Multiday Maintenance of Extracorporeal Lungs Using Cross-Circulation With Conscious Swine. J Thorac Cardiovasc Surg (2020) 159(4):1640–53. doi:10.1016/j.jtcvs.2019.09.121
142. Chen, P, Van Hassel, J, Pinezich, MR, Diane, M, Hudock, MR, Kaslow, SR, et al. Recovery of Extracorporeal Lungs Using Cross-Circulation With Injured Recipient Swine. J Thorac Cardiovasc Surg (2024) 167(5):e106–30. doi:10.1016/j.jtcvs.2023.09.032
143. Hozain, AE, O’Neill, JD, Pinezich, MR, Tipograf, Y, Donocoff, R, Cunningham, KM, et al. Xenogeneic Cross-Circulation for Extracorporeal Recovery of Injured Human Lungs. Nat Med (2020) 26(7):1102–13. doi:10.1038/s41591-020-0971-8
144. Kelly Wu, W, Guenthart, BA, O’Neill, JD, Hozain, AE, Tipograf, Y, Ukita, R, et al. Technique for Xenogeneic Cross-Circulation to Support Human Donor Lungs Ex Vivo. J Heart Lung Transpl Off Publ Int Soc Heart Transpl (2023) 42(3):335–44. doi:10.1016/j.healun.2022.11.002
145. Wu, WK, Stier, MT, Stokes, JW, Ukita, R, Patel, YJ, Cortelli, M, et al. Immune Characterization of a Xenogeneic Human Lung Cross-Circulation Support System. Sci Adv (2023) 9(13):eade7647. doi:10.1126/sciadv.ade7647
146. Glorion, M, Pascale, F, Estephan, J, Huriet, M, Gouin, C, Urien, C, et al. A Cross-Circulatory Platform for Monitoring Innate Allo-Responses in Lung Grafts. PLoS One (2023) 18(5):e0285724. doi:10.1371/journal.pone.0285724
147. Nishitai, R, Ikai, I, Terajima, H, Kanazawa, A, Takeyama, O, Uesugi, T, et al. Influence of Extracorporeal Porcine Liver Perfusion on Nonhuman Primates: Minimizing Hemolysis Improves Subsequent Survival. Liver Transpl Off Publ Am Assoc Study Liver Dis Int Liver Transpl Soc (2001) 7(7):615–22. doi:10.1053/jlts.2001.25362
148. Nara, M, Hakamada, K, Totsuka, E, Nozaki, T, Takiguchi, M, and Sasaki, M. Efficacy of Double Filtration Plasmapheretic Cross-Circulation Using a High Permeability Membrane Between Totally Hepatectomized Dogs and Donor Pigs. Transplantation (2001) 72(11):1736–42. doi:10.1097/00007890-200112150-00006
149. Ishizawa, Y, Totsuka, E, Umehara, M, Nishimura, A, Ono, H, and Sasaki, M. Efficacy of Double-Filtration Plasmapheretic Cross-Circulation With a High-Permeability Membrane Using Canine Harvested Liver in Porcine Fulminant Hepatic Failure Model. Transpl Proc (2004) 36(8):2344–8. doi:10.1016/j.transproceed.2004.06.038
150. Umehara, M, Totsuka, E, Ishizawa, Y, Nara, M, Hakamada, K, and Sasaki, M. In vitro Evaluation of Cross-Circulation System Using Semipermeable Membrane Combined With Whole Liver Perfusion. Transpl Proc (2004) 36(8):2349–51. doi:10.1016/j.transproceed.2004.08.009
151. Wu, WK, Tumen, A, Stokes, JW, Ukita, R, Hozain, A, Pinezich, M, et al. Cross-Circulation for Extracorporeal Liver Support in a Swine Model. ASAIO J Am Soc Artif Intern Organs (2022) 68(4):561–70. doi:10.1097/mat.0000000000001543
152. Wu, WK, Ukita, R, Patel, YJ, Cortelli, M, Trinh, VQ, Ziogas, IA, et al. Xenogeneic Cross-Circulation for Physiological Support and Recovery of Ex Vivo Human Livers. Hepatol Baltim Md (2023) 78(3):820–34. doi:10.1097/HEP.0000000000000357
153. Church, JT, Alghanem, F, Deatrick, KB, Trahanas, JM, Phillips, JP, Hee Song, M, et al. Normothermic Ex Vivo Heart Perfusion: Effects of Live Animal Blood and Plasma Cross Circulation. ASAIO J Am Soc Artif Intern Organs (2017) 63(6):766–73. doi:10.1097/MAT.0000000000000583
154. McLeod, JS, Poling, C, Church, JT, Jung, J, Sarosi, E, Langley, M, et al. Ex Vivo Heart Perfusion for 72 Hours Using Plasma Cross Circulation. ASAIO J Am Soc Artif Intern Organs (2020) 66(7):753–9. doi:10.1097/MAT.0000000000001061
155. Tchouta, L, Drake, D, Hoenerhoff, M, Rojas-Pena, A, Haft, J, Owens, G, et al. Twenty-Four-Hour Normothermic Perfusion of Isolated Ex Vivo Hearts Using Plasma Exchange. J Thorac Cardiovasc Surg (2022) 164(1):128–38. doi:10.1016/j.jtcvs.2020.11.158
156. Zhang, X, Su, L, and Pan, P. Advances and Applications of Lung Organoids in the Research on Acute Respiratory Distress Syndrome (ARDS). J Clin Med (2024) 13(2):346. doi:10.3390/jcm13020346
157. Jain, N, Shashi Bhushan, BL, Natarajan, M, Mehta, R, Saini, DK, and Chatterjee, K. Advanced 3D In Vitro Lung Fibrosis Models: Contemporary Status, Clinical Uptake, and Prospective Outlooks. ACS Biomater Sci Eng (2024) 10(3):1235–61. doi:10.1021/acsbiomaterials.3c01499
158. Shah, D, Dave, B, Chorawala, MR, Prajapati, BG, Singh, S, M Elossaily, G, et al. An Insight on Microfluidic Organ-On-A-Chip Models for PM2.5-Induced Pulmonary Complications. ACS Omega (2024) 9(12):13534–55. doi:10.1021/acsomega.3c10271
159. Ma, C, Peng, Y, Li, H, and Chen, W. Organ-on-a-Chip: A New Paradigm for Drug Development. Trends Pharmacol Sci (2021) 42(2):119–33. doi:10.1016/j.tips.2020.11.009
160. Madhvapathy, SR, Wang, JJ, Wang, H, Patel, M, Chang, A, Zheng, X, et al. Implantable Bioelectronic Systems for Early Detection of Kidney Transplant Rejection. Science (2023) 381(6662):1105–12. doi:10.1126/science.adh7726
161. Guyette, JP, Gilpin, SE, Charest, JM, Tapias, LF, Ren, X, and Ott, HC. Perfusion Decellularization of Whole Organs. Nat Protoc (2014) 9(6):1451–68. doi:10.1038/nprot.2014.097
162. Aamodt, JM, and Grainger, DW. Extracellular Matrix-Based Biomaterial Scaffolds and the Host Response. Biomaterials (2016) 86:68–82. doi:10.1016/j.biomaterials.2016.02.003
163. Saxton, SH, and Stevens, KR. 2D and 3D Liver Models. J Hepatol (2023) 78(4):873–5. doi:10.1016/j.jhep.2022.06.022
164. Yang, S, Ooka, M, Margolis, RJ, and Xia, M. Liver Three-Dimensional Cellular Models for High-Throughput Chemical Testing. Cell Rep Methods (2023) 3(3):100432. doi:10.1016/j.crmeth.2023.100432Available from: https://www.cell.com/cell-reports-methods/abstract/S2667-2375(23)00047-4 (Accessed March 25, 2024)
165. Mazza, G, Rombouts, K, Rennie Hall, A, Urbani, L, Vinh Luong, T, Al-Akkad, W, et al. Decellularized Human Liver as a Natural 3D-Scaffold for Liver Bioengineering and Transplantation. Sci Rep (2015) 5(1):13079. doi:10.1038/srep13079
166. Soto-Gutierrez, A, Zhang, L, Medberry, C, Fukumitsu, K, Faulk, D, Jiang, H, et al. A Whole-Organ Regenerative Medicine Approach for Liver Replacement. Tissue Eng C Methods (2011) 17(6):677–86. doi:10.1089/ten.TEC.2010.0698
167. Wang, J, and Jin, X. Strategies for Decellularization, Re-CellularIzation and Crosslinking in Liver Bioengineering. Int J Artif Organs (2024) 47(3):129–39. doi:10.1177/03913988231218566
168. Leuning, DG, Witjas, FMR, Maanaoui, M, de Graaf, AMA, Lievers, E, Geuens, T, et al. Vascular Bioengineering of Scaffolds Derived From Human Discarded Transplant Kidneys Using Human Pluripotent Stem Cell–Derived Endothelium. Am J Transpl (2019) 19(5):1328–43. doi:10.1111/ajt.15200
169. Chani, B, Puri, V, Sobti, RC, Jha, V, and Puri, S. Decellularized Scaffold of Cryopreserved Rat Kidney Retains its Recellularization Potential. PLoS ONE (2017) 12(3):e0173040. doi:10.1371/journal.pone.0173040
170. Salti, H, Kramer, L, Nelz, SC, Lorenz, M, Breitrück, A, Hofrichter, J, et al. Decellularization of Precision-Cut Kidney Slices—Application of Physical and Chemical Methods. Biomed Mater (2023) 18(2):025004. doi:10.1088/1748-605X/acb02e
171. O’Neill, JD, Anfang, R, Anandappa, A, Costa, J, Javidfar, J, Wobma, HM, et al. Decellularization of Human and Porcine Lung Tissues for Pulmonary Tissue Engineering. Ann Thorac Surg (2013) 96(3):1046–55. discussion 1055-1056. doi:10.1016/j.athoracsur.2013.04.022
172. Petrella, F, and Spaggiari, L. Artificial Lung. J Thorac Dis (2018) 10(Suppl. 20):S2329-S2332–32. doi:10.21037/jtd.2017.12.89
173. De Santis, MM, Bölükbas, DA, Lindstedt, S, and Wagner, DE. How to Build a Lung: Latest Advances and Emerging Themes in Lung Bioengineering. Eur Respir J (2018) 52(1):1601355. doi:10.1183/13993003.01355-2016
174. Ren, X, Moser, PT, Gilpin, SE, Okamoto, T, Wu, T, Tapias, LF, et al. Engineering Pulmonary Vasculature in Decellularized Rat and Human Lungs. Nat Biotechnol (2015) 33(10):1097–102. doi:10.1038/nbt.3354
175. McCauley, KB, Hawkins, F, Serra, M, Thomas, DC, Jacob, A, and Kotton, DN. Efficient Derivation of Functional Human Airway Epithelium From Pluripotent Stem Cells via Temporal Regulation of Wnt Signaling. Cell Stem Cell (2017) 20(6):844–57. doi:10.1016/j.stem.2017.03.001
176. Huang, SXL, Green, MD, de Carvalho, AT, Mumau, M, Chen, YW, D’Souza, SL, et al. The In Vitro Generation of Lung and Airway Progenitor Cells From Human Pluripotent Stem Cells. Nat Protoc (2015) 10(3):413–25. doi:10.1038/nprot.2015.023
Keywords: transplantation, machine perfusion, bioengineering, gene therapy, cross circulation
Citation: Niroomand A, Nita GE and Lindstedt S (2024) Machine Perfusion and Bioengineering Strategies in Transplantation—Beyond the Emerging Concepts. Transpl Int 37:13215. doi: 10.3389/ti.2024.13215
Received: 01 May 2024; Accepted: 19 August 2024;
Published: 29 August 2024.
Copyright © 2024 Niroomand, Nita and Lindstedt. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.
*Correspondence: Sandra Lindstedt, sandra.lindstedt_ingemansson@med.lu.se