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Ann Thorac Surg 1998;66:225-230
© 1998 The Society of Thoracic Surgeons

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Original articles: general thoracic

Addition of aprotinin to organ preservation solutions decreases lung reperfusion injury

^a Division of Cardiothoracic Surgery, Department of Surgery, University of Southern California School of Medicine and Childrens Hospital Los Angeles, Los Angeles, California, USA

Accepted for publication February 20, 1998.

Address reprint requests to Dr Barr, Division of Cardiothoracic Surgery, University of Southern California, 1510 San Pablo St, Los Angeles, CA 90033

Presented at the Annual Meeting of the American Society of Transplant Surgeons, Chicago, IL, May 14–16, 1997.

	Abstract

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Background. Organ preservation injury is associated with endothelial cell damage, destabilization of mitochondrial and cell membranes, and the release of proteolytic enzymes. In addition to its well-known clinical effect of reducing perioperative blood loss, aprotinin has antiproteolytic and membrane-stabilizing properties. We hypothesized that adding aprotinin to Euro-Collins (EC) and University of Wisconsin (UW) solutions would decrease preservation injury in cultured endothelial cells and a whole organ rat lung model.

Methods. Bovine aortic endothelial cells were cultured and stored in the respective solution at 4°C for 12 or 48 hours. Endothelial cell viability after storage was assessed by dimethylthiazole tetrazolium cytotoxicity assay. In the whole organ model, rat lungs were isolated, flushed with the respective solution, and stored at 4°C for 6 or 12 hours. The lungs were ventilated with 100% O₂ and reperfused with fresh blood. Alveolar–arterial O₂ difference, O₂ tension, capillary filtration coefficient, and compliance were determined.

Results. Endothelial cell viability was optimized with the addition of aprotinin to EC and UW at a dose of 150 KIU/mL (0.02 mg/mL). In the isolated perfused lung model, after 6 hours of ischemic storage, aprotinin-enhanced (100 KIU/mL [0.014 mg/mL]) EC and UW decreased alveolar–arterial O₂ difference, increased O₂ tension, and decreased capillary filtration coefficient compared with EC and UW alone. After 12 hours of ischemic storage, aprotinin-enhanced EC and UW decreased alveolar–arterial O₂ difference, increased O₂ tension, decreased capillary filtration coefficient, and increased compliance compared with EC and UW alone.

Conclusions. The addition of aprotinin to EC and UW solutions increases endothelial cell viability in hypoxic cold storage conditions. In terms of whole organ function, aprotinin improves lung preservation as demonstrated by increased oxygenation and compliance, and decreased capillary permeability. This study is clinically applicable as there is already extensive experience with the use of aprotinin in heart and lung transplant recipients, in addition to its routine use in conventional cardiac operations.

	Introduction

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Optimizing posttransplantation graft function after cold storage provides a continuing challenge in solid organ transplantation. The transplanted lung is particularly susceptible to the deleterious effects of ischemia and reperfusion even with a storage time of only 4 to 6 hours.

Organ preservation injury is associated with lysosomal enzyme release, activation of proteolytic enzymes, and endothelial cell damage. Several studies, primarily in myocardial preservation, have shown that the loss of lysosomal membrane integrity and the activation of released lysosomal enzymes leads to many of the biochemical and structural changes associated with ischemic tissue damage [1, 2]. Aprotinin (Trasylol; Bayer Pharmaceuticals, West Haven, CT) is a serine protease inhibitor that is presently in wide clinical use for minimizing perioperative blood loss in cardiac operations and has been shown to protect against the damage of ischemia and reperfusion by suppressing the release of lysosomal enzymes and inhibiting their activities [3–6].

Previous studies with aprotinin have shown it to improve myocardial, hepatic, and renal viability after a period of ischemia followed by reperfusion [7–9]. In this study, we investigated whether aprotinin can provide protection against the adverse effects of ischemia and reperfusion in the lung.

In this study, the dose range of aprotinin providing optimal protection during cold storage and reperfusion was determined using cultured endothelial cells. That dose range was then tested in a whole organ isolated blood-perfused rat lung model. Endothelial cell viability after cold hypoxic storage in standard clinical preservation solutions—Euro-Collins (EC) and University of Wisconsin (UW)—was compared with storage in the same solutions with aprotinin. In the isolated perfused lung model, lung function as measured by oxygenation, compliance, and capillary permeability was evaluated after a period of cold storage followed by reperfusion. Again, EC and UW alone were compared with those same solutions with the addition of aprotinin.

	Material and methods

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Endothelial cell studies
Bovine aortic endothelial cells were isolated from fresh calf aortas. Endothelial cells were mechanically dislodged by injection and aspiration in a closed segment of aorta using minimum essential medium supplemented with 10% fetal bovine serum (Gibco, Gaithersburg, MD). Cells were then cultured in medium 199 (Sigma, St. Louis, MO) with 10% fetal bovine serum until confluent growth was achieved. Media was replaced with preservation solution, and the cells stored for 12 or 48 hours at 4°C in a nitrogen/CO₂ environment to simulate hypoxic conditions during organ preservation. The preservation solutions tested included EC alone, EC with aprotinin (EC+A) at 50 kallikrein inactivation units (KIU)/mL (0.007 mg/mL), EC+A at 150 KIU/mL (0.02 mg/mL), EC+A at 350 KIU/mL (0.05 mg/mL), UW alone, UW with aprotinin (UW+A) at 50 KIU/mL, UW+A at 150 KIU/mL, and UW+A at 350 KIU/mL. Sixteen wells were tested for each group (n = 16). Endothelial cell viability was determined by dimethylthiazole tetrazolium assay as described previously [10]. After storage, cells were placed in Dulbecco’s modified Eagle’s medium/Ham’s nutrient mixture F-12 (Sigma) plus 16% fetal bovine serum containing 1 mg/mL dimethylthiazole tetrazolium. In viable cells, dimethylthiazole tetrazolium is converted to formazan by mitochondria. Formazan concentration was detected by changes in optical density at 490 nm as read by a microELISA spectrophotometer (Molecular Devices, Sunnyvale, CA).

Isolated perfused lung model
Donor procedure
Male Lewis rats (250 to 350 g) were anesthetized (sodium pentobarbital, 50 mg/kg, intraperitoneally) and tracheostomized with a 14-gauge angiocatheter. The lungs were ventilated (Harvard Apparatus, South Natick, MA) with 100% O₂ at a respiratory rate of 50 breaths/min, tidal volume of 10 mL/kg, and positive end-expiratory pressure of 2 cm H₂O. A median sternotomy and thymectomy were performed, exposing the heart–lung block. The animals were heparinized (1,000 U/kg) through the inferior vena cava, and the apex of the heart was amputated. The lungs were flushed through the main pulmonary artery with cold preservation solution (4°C; 50 mL/kg) from a height of 30 cm. The heart–lung block was excised, suspended from a support rod in the esophagus, and submerged with the lungs inflated in the corresponding preservation solution for storage at 4°C.

Blood harvest
Male Lewis rats (250 to 350 g) were anesthetized as described previously. A sternotomy and laparotomy were performed and the animal was heparinized (1,000 U/kg) through the inferior vena cava. Blood was obtained through a subdiaphragmatic inferior vena caval puncture and the animal was euthanized (pentobarbital 100 mg/kg, intraperitoneally).

Reperfusion
After a 6- or 12-hour storage time, the pulmonary artery and left atrium were cannulated with 14-gauge angiocatheters, and the lung block was suspended by the esophagus from a force transducer in a humidified, temperature-controlled (37°C), vented (to atmospheric pressure) chamber. The lungs were ventilated with 100% O₂ at a rate of 50 breaths/min with a tidal volume of 10 mL/kg body weight and a positive end-expiratory pressure of 2 cm H₂O. Reperfusion was initiated by connecting the pulmonary arterial cannula to a reservoir containing the previously harvested fresh syngeneic venous blood suspended at a height that generated an inflow pressure of 15 mm Hg. The left atrial cannula was then connected to a separate reservoir and the heights of the reservoirs were used to control pulmonary capillary pressure during reperfusion. The capillary pressure was adjusted to create an isogravimetric state (no weight loss or gain) and maintained in that state for 5 minutes. After the isogravimetric period, the reservoirs were raised to create a change in capillary pressure of 15 mm Hg and maintained for 15 minutes.

Monitoring
Throughout the reperfusion period, specimen weight (FT 03 force-displacement transducer; Grass Instruments, Quincy, MA), inflow pressure and outflow pressure (DTX model T4812 AD-R pressure transducer; Viggo-Spectramed, Oxnard, CA), tracheal pressure (DP 45-24 differential pressure transducer; Validyne, Northridge, CA), and tidal volume (Fleisch No. 2 pneumotachograph, OEM, Richmond, VA; Validyne DP 45-14 pressure transducer) were measured. All data were transmitted to an AST Bravo (Fort Worth, TX) IBM-compatible computer through a Validyne CD 280 and MC 1-3 carrier demodulator and a Dash 16 analog-to-digital converter, and processed and displayed by a customized version of Microsoft Visual Basic Pro software. Data was collected and recorded every 10 seconds during the entire reperfusion period. Blood was collected from the left atrium at the end of reperfusion, and the oxygen tension measured using a blood gas analyzer (Ciba-Corning 288, Medfield, MA).

Experimental protocol
Isolated lungs were flushed with either modified EC (Euro-Collins; Baxter Health Care Products, Deerfield, IL; modified with 2.5 mL of 50% magnesium sulfate and 50 mL of 50% dextrose per liter) or UW (Viaspan; Dupont Merck Pharmaceutical, Wilmington, DE with 40 U insulin and 16 mg dexamethasone per liter) solution and stored in the same solution for 6 or 12 hours. Control groups were flushed and stored using standard solution, and experimental groups were flushed and stored using solution with the addition of aprotinin at one of four doses: 50 KIU/mL (0.007 mg/mL), 100 KIU/mL (0.014 mg/mL), 150 KIU/mL (0.02 mg/mL), and 350 KIU/mL (0.05 mg/mL). At each storage time there were two control and eight experimental groups consisting of 6 animals each: standard EC, standard UW, EC+A at 50, 100, 150, and 350 KIU/mL (EC+A50, A100, A150, A350), UW+A at the same doses (UW+A50, A100, A150, A350). All animals received humane care in compliance with the "Principles of Laboratory Animal Care" formulated by the National Society for Medical Research and the "Guide for the Care and Use of Laboratory Animals" prepared by the National Academy of Science and published by the National Institutes of Health (NIH publication 85-23, revised 1985).

Data analysis
Determination of capillary filtration coefficient
The capillary filtration coefficient (Kf) is a sensitive measure of capillary permeability calculated according to the method described previously by Drake and colleagues [11]. Any organ system has a measurable permeability determined by the amount of fluid that will be filtered across a capillary bed for a given imbalance in Starling forces. The Kf is a mathematical representation of this permeability. In our model, the pulmonary capillary pressure was determined by the inflow and outflow pressures according to the method of Gaar and colleagues [12]. The heights of the inflow and outflow reservoirs were adjusted such that the lungs neither gained nor lost weight (isogravimetric state). To measure Kf, we then raised the reservoirs, creating a Starling imbalance by increasing the hydrostatic pressure in the pulmonary capillaries. After a short period of vascular filling, fluid begins to extravasate into the interstitium of the lung. This extravasation causes a predictable rise in specimen weight. After a period of rapid weight gain (representing vascular filling), the lungs enter a period of slow weight gain. If the rate of slow weight gain is measured and, under a logarithmic transformation, extrapolated to the time of initiating the Starling imbalance, the initial rate of fluid filtration can be estimated. This rate, when divided by the measured change in capillary pressure and normalized to 100 g wet lung weight, gives a constant (Kf) that represents the permeability of the microvasculature of that specimen. Higher Kf values correspond to greater capillary permeability.

Blood gas analysis
A blood sample from the left atrium was obtained at the end of reperfusion from which oxygen tension (pO₂) was measured and the alveolar–arterial oxygen difference (AaDO₂) was calculated.

Compliance
Recorded tidal volume and airway pressure values were measured every 10 seconds. Dynamic lung compliance was assessed during the isogravimetric period by calculating the ratio of tidal volume divided by the corresponding change in tracheal pressure from end expiration to end inspiration.

Statistical analysis
All data are reported as the mean ± standard deviation. The data were analyzed by analysis of variance and all comparisons tested by the Student-Newmann-Keuls multiple comparisons test (p < 0.05 showing statistical significance).

	Results

Top Abstract Introduction Material and methods Results Comment Acknowledgments References

 
Endothelial cell studies
Endothelial cell viability as determined by dimethylthiazole tetrazolium assay optical density is shown for 12 and 48 hours of ischemic storage in Figure 1. A dose titration of aprotinin (0, 50, 150, and 350 KIU/mL) added to EC and UW is shown. Higher optical density values correlate with better dye absorption indicating better cell viability. At 12 hours of storage, viability was maximal at a dose of 150 KIU/mL for both EC and UW. At 48 hours of storage, viability was maximal at a dose of 150 KIU/mL for UW and 50 KIU/mL for EC.

View larger version (35K): [in this window] [in a new window]   Fig 1. Dimethylthiazole tetrazolium assay optical density for a dose titration of aprotinin (A) at 12 and 48 hours of ischemic storage. Results for both Euro-Collins (EC) and University of Wisconsin (UW) solutions are shown for both storage times. Aprotinin doses tested included 0, 50, 150, and 350 kallikrein inactivation units per milliliter (KIU/mL). Significant comparisons of an optimal dose of 150 KIU/mL to the solutions alone include EC+A versus EC at 12 hours, p < 0.01; UW+A versus UW at 12 hours, p < 0.05; EC+A versus EC at 48 hours, p < 0.05; and UW+A versus UW at 48 hours, p < 0.001.

Isolated perfused lung
Dose titration
The outflow oxygen tension results for a dose titration of aprotinin (50, 100, 150, and 350 KIU/mL) added to EC and UW are shown in Table 1. Adding 100 KIU/mL aprotinin to EC gives maximal O₂ tension at both 6 and 12 hours. For UW, 150 KIU/mL maximizes O₂ tension at 6 hours, whereas 50 KIU/mL maximizes O₂ tension at 12 hours. Capillary filtration coefficient results for the same dose titration of aprotinin added to EC and UW are shown in Table 2. For both solutions and at both 6 and 12 hours of storage, an aprotinin dose of 100 KIU/mL gave the lowest (optimal) Kf values. Based on these results, an "optimal" aprotinin dose of 100 KIU/mL was added to EC and UW and compared to the preservation solutions alone for all of the measured parameters.

View larger version (33K): [in this window] [in a new window]   Fig 2. Oxygen tension (pO2) in mm Hg for all groups at 6 and 12 hours of storage. Significant comparisons at 6 hours of storage were EC+A versus EC, p < 0.001; UW+A versus UW, p < 0.001. Significant comparisons at 12 hours were EC+A versus EC, p < 0.001; UW+A versus UW, p < 0.01; and UW versus EC, p < 0.05. (A = aprotinin; EC = Euro-Collins; UW = University of Wisconsin.)

View larger version (33K): [in this window] [in a new window]   Fig 3. Alveolar–arterial oxygen difference (AaDO2) in mm Hg for all groups at 6 and 12 hours of storage. Significant comparisons at 6 hours were EC+A versus EC, p < 0.001; UW+A versus UW, p < 0.001. Significant comparisons at 12 hours were EC+A versus EC, p < 0.001; UW+A versus UW, p < 0.001; and UW versus EC, p < 0.01. (A = aprotinin; EC = Euro-Collins; UW = University of Wisconsin.)

View larger version (33K): [in this window] [in a new window]   Fig 4. Compliance for all groups at 6 and 12 hours of storage. Compliance was measured as tidal volume/change in airway pressure and expressed in mL/cm H2O. The only significant comparison at 6 hours of storage was UW+A versus EC+A, p < 0.05. Significant comparisons at 12 hours were EC+A versus EC, p < 0.001; UW+A versus UW, p < 0.01; UW versus EC, p < 0.001; and UW+A versus EC+A, p < 0.001. (A = aprotinin; EC = Euro-Collins; UW = University of Wisconsin.)

View larger version (24K): [in this window] [in a new window]   Fig 5. Capillary filtration coefficient (Kf) for all groups at 6 and 12 hours of storage. The capillary filtration coefficient was normalized to 100 g wet lung weight and expressed in mL · min-1 · mm Hg-1 · 100 g-1. Significant comparisons at 6 hours were EC+A versus EC, p < 0.001; UW+A versus UW, p < 0.001; UW versus EC, p < 0.05; UW+A versus EC+A, p < 0.01. Significant comparisons at 12 hours were EC+A versus EC, p < 0.001; UW+A versus UW, p < 0.001; and UW versus EC, p < 0.001. (A = aprotinin; EC = Euro-Collins; UW = University of Wisconsin.)

	Comment

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Liver preservation injury has been shown to be ameliorated by aprotinin. Studies in rats and pigs have demonstrated that aprotinin treatment reduces serum levels of hepatocellular enzymes (serum glutamic-oxaloacetic transaminase and serum glutamate pyruvate transaminase) and increases safe organ storage time [8, 14]. Similarly, studies in rats and sheep subjected to kidney ischemia and reperfusion showed better glomerular preservation and better renal function with aprotinin treatment [9]. This study was designed to examine whether these protective properties of aprotinin could reduce ischemia–reperfusion injury in the lung and improve preservation, which represents a much greater clinical problem in routine lung transplantation than in other solid organs.

Previous studies with aprotinin in organ preservation have used a variety of dosing regimens and delivery methods, including systemic administration to the donor or recipient and addition to the flush or reperfusion solution. In some cases combinations of delivery methods were used. In clinical lung transplantation, the most immediately relevant means of delivering aprotinin to the graft is in the flush solution at the time of harvest. This allows delivery to the specific organ of interest in the donor, and does not require dosing the recipient unless otherwise clinically indicated. Therefore, we added aprotinin to the flush and storage solution only in our model.

To determine the optimal dose of aprotinin for preservation, endothelial cell viability studies were done using a dose titration of aprotinin. Aprotinin improved endothelial cell viability and maximal viability was noted at a dose of 150 KIU/mL. These data allowed us to perform a tighter dose titration in the isolated perfused lung model. The addition of aprotinin to EC and UW did not result in any visible particulate matter or discoloration.

The goals of pulmonary preservation efforts in transplantation are compliant lungs that oxygenate well and are resistant to edema formation. In this isolated blood-perfused rat lung model, the addition of aprotinin to EC and UW solutions improved lung function by all of these measures. Lungs in the aprotinin groups were more compliant, developed higher outflow oxygen tensions, resulted in lower AaDO₂, and demonstrated lower pulmonary Kf at an optimal dose of 100 KIU/mL.

This study did not attempt to elucidate the mechanism by which aprotinin improves lung preservation. Probably, antiproteolytic and lysosomal membrane stabilizing properties, as shown in myocardial preservation studies, are also functioning in the lung. Aprotinin has been shown to reduce cytokine-induced nitric oxide synthetase expression in airways [15] and interleukin-8-dependent neutrophil accumulation in the lungs during cardiopulmonary bypass [16]. Further studies to determine the relevant mechanisms are indicated. In addition, although our study uses a sensitive experimental model for lung function, it does not involve a complete organ transplant with all of the concomitant recipient variables. Animal studies involving orthotopic lung transplantation would provide further information as to the potential benefits of aprotinin on the function of transplant grafts. Because aprotinin is a bovine protein, the potential for hypersensitivity exists, particularly with reexposure, in regards to application of these results to clinical practice.

Currently aprotinin enjoys widespread use in patients on cardiopulmonary bypass for its hemostatic properties [17]. Aprotinin has been used extensively in lung transplant recipients and several studies have shown its beneficial effects on blood loss and transfusion requirements [18, 19]. The use of aprotinin as an adjunct to preservation into clinical practice would then be a natural extension with potentially significant impact as ischemia–reperfusion injury and the resulting primary graft dysfunction remains a persistent cause of early graft loss in lung transplantation [20].

	Acknowledgments

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This work was supported by a grant to Dr Barr from the Heart and Lung Surgery Foundation, Los Angeles, CA. We also acknowledge Carol M. Cassidy, MS, and Stan Horton, PharmD, from Bayer Pharmaceuticals Division and Stanley M. Yamashiro, PhD, from the University of Southern California School of Engineering for their assistance and support, and Erin Rachel for her assistance in preparing the manuscript.

	References

Top Abstract Introduction Material and methods Results Comment Acknowledgments References

 

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