Once anesthetized, the animals must be intubated with 14 G catheters for rats >300g, 16 G for rats 200–300g or 22 G for mice weighing around 25g, and mechanically ventilated. For the recipient, the pressure-controlled mode is usually recommended, since the left lung will not be functional during a large part of the surgery.

The ventilator should be set to a tidal volume of 7–10ml/kg (if volume-controlled ventilation is used), or a maximum pressure of 10–12cmH2O (in the case of pressure-controlled ventilation), with a respiratory rate of 60–100breaths/minute. Moreover, a positive end-expiratory pressure of 2–5cmH2O is highly recommended for avoiding the formation of atelectasis during expiration.

Equipment specifically designed for rodents is available for recording pulse oximetry, capnography, non-invasive arterial pressure or electrocardiogram. Devices designed for humans or large animals are generally unsuitable for rodents.

It is also important to monitor body temperature using a thermometer or rectal probe and to maintain it between 37.5 and 38.5°C. Hypothermia is a very common complication in rodents and adverse events can occur if temperatures fall below 32°C, including delayed recovery from anesthesia. An electric blanket or hot water bottle can be used to maintain body temperature, although the animal must be monitored to avoid possible hyperthermia. Oxygen saturation tends to remain at levels above 99% when the inspired fraction of oxygen is increased to between 30% and 60%, compared to 21% ambient air.

Extraction of the complete heart–lung block is generally recommended. The animal is anesthetized and placed in a supine position, and the operative field is prepared by shaving the skin of the chest and applying povidone iodine or chlorhexidine in an outward movement from the center of the chest. A median sternotomy is then performed with inferior vena cava, subcutaneous or intracoronary heparin delivery (100IU/100g) to prevent clots forming in the lung for transplant. Using an optical microscope, the lung hilum is dissected to locate the aortic arch, which is then clamped. Incisions are simultaneously made in the atria and the inferior vena cava.

In most models, anterograde perfusion is performed via the right ventricle19,20 or the pulmonary trunk15,21–23although it is advisable to combine this procedure with retrograde perfusion from the left atrium22 to simulate physiological conditions. The lungs may be perfused at a constant, uniform pressure (10–15cmH2O) with cold preservation fluid (4°C), consisting of low potassium dextran glucose (LPDG).

The trachea is then ligated and dissected for extraction of the entire heart–lung block, with the lung inflated to prevent the formation of atelectasis and to improve oxygenation and the presence of surfactant in order to reduce ischemia–reperfusion injury.24,25 Recommendations for inflation range between 50%22,25 and 75%–100%26 of total lung capacity, taking care to avoid barotrauma. Inflation before preservation also reduces the expression of proinflammatory markers after reperfusion.26

For preservation, the donor lung is generally maintained at 4°C to minimize ischemic injury to the graft.27 The most commonly used preservation solution is low potassium dextran (Perfadex),16,28–30 although others are available, including Celsior,31 Euro-Collins or Wisconsin.24

When the lung block has been extracted, the left lung must be separated, clamping the bronchus to keep it inflated. The bronchus, the artery and the pulmonary veins are dissected as distally as possible to the lung to assist anastomosis (Fig. 1).

Fig. 1.

Extraction and preparation of the donor lung. Median sternotomy and dissection and extraction of the heart–lung block. Dissection of the left pulmonary artery (LPA), left pulmonary veins (LPV), and the left bronchus (LBr). AO: thoracic aorta; IVC: inferior vena cava; LA: left atrium; L SVC: left superior vena cava; LL: left left; RA: right atrium; R SVC: right superior vena cava.

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Preparation of the left lung will vary depending on the anastomosis method used: vascular and bronchial anastomosis with sutures,6,19,21,32 bronchial anastomosis with sutures and vascular anastomosis with cuffs,30,33–36 and vascular and bronchial anastomosis with cuffs20,22,23,35–39 (Fig. 2). In the first case, the lung hilum is dissected, separating each structure for anastomosis, while maintaining the lung in the low temperature preservation medium. In the latter 2 cases, rather than using sutures, the vascular and bronchial anastomoses are simplified by using cuffs designed for transplantation of lungs33 and other organs.40–42 These are ring-shaped devices made of polyvinyl chloride or Teflon®, prepared from cannulae or catheters. Sizes for rats range from 18G to 14G, the larger ones being used in the bronchus and the smaller ones in the pulmonary artery.22 Conventional cuffs consist of a main body, on which the vessels or the bronchus are joined and an extension tab to facilitate manipulation (Fig. 3). The cuff is 2–3mm in length (body 1, 1.5 or 2mm and the tab 1 or 1.5mm). A modified, tabless cuff model has been developed,32,36,42 but this makes it more complicated to manipulate. For this reason, some groups use the standard cuff, and remove the tab after anastomosis has been performed.33

Fig. 2.

Surgical techniques in mouse model lung transplantation. (A) Vascular and bronchial anastomosis with sutures. (B) Bronchial anastomosis with sutures and use of vascular cuffs in the pulmonary vein and artery. (C) Vascular and bronchial anastomosis using cuffs. LBr d: left bronchus of donor animal; LBr r: left bronchus of recipient animal; LPA d: left pulmonary artery of donor animal; LPA r: left pulmonary artery of recipient animal; LPV d: left pulmonary vein of donor animal; LPV r: left pulmonary vein of recipient animal.

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Fig. 3.

Placement of a cuff in the donor lung and anastomosis with the recipient lung. LBr d: left bronchus of donor animal; LBr r: left bronchus of recipient animal; LPA d: left pulmonary artery of donor animal; LPA r: left pulmonary artery of recipient animal; LPV d: left pulmonary vein of donor animal; LPV r: left pulmonary vein of recipient animal.

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Irrespectively of the type of cuff used, the procedure is conducted by introducing the vein, artery or bronchus of the donor lung through the hole of the cuff. It is then everted over the cuff and fixed to the cuff with a polypropylene ligature, 7/0 or 8/0 in rats22,23 and 10/0 in mice.18 The bronchus is occasionally prepared at the moment of implantation to prevent preservation solution from infiltrating the airway.18,20

The animal is anesthetized, given appropriate analgesia and placed in right lateral decubitus position. The surgical field is prepared in the same way as for the donor animal and a lateral thoractomy is made through the fourth intercostal space. The left lung is extracted and the principal structures are isolated, clamping the left pulmonary artery, vein and bronchus, as distally as possible to the lung. These structures are dissected as close as possible to the pulmonary parenchyma. To avoid the formation of thrombi, heparinized saline (0.5–1IU/ml) is administered to the artery and vein. All structures must be handled carefully to avoid vascular tears.

The vascular structures and the bronchus are then anastomosed. Cuff anastomosis is shown in the photograph in Fig. 4.

Fig. 4.

Photograph of lung transplantation performed using the cuff technique in blood vessels and bronchus. DL: donor lung; LBr r: left bronchus of recipient animal; LPA r: left bronchus of recipient animal; LPV r: left pulmonary vein of recipient animal.

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When the bronchus has been anastomosed, the donor lung must be ventilated to avoid the formation of atelectasis and to enhance alveolar recruitment. Rapid lung reperfusion indicates that the anastomosis is patent. Finally, the native lung is extracted from the recipient animal, leaving only the donor lung.

A drainage tube must be inserted so that remains of blood, fluid or air around the lungs can be eliminated and the lungs can expand normally. The chest cavity is then closed by layers, and the drainage tube is withdrawn when the animal recovers from anesthesia and begins to breathe spontaneously.

The time to complete surgery varies, depending on the group of professionals and the technique used. In general, the procedure is faster with the cuff technique than with conventional sutures, particularly when sutures are applied to vessels and the bronchus. Similarly, the survival rate is higher when anastomosis is performed with cuffs, compared to sutures (Table 2).

The most significant complications occur as a result of the surgical procedure, in the preparation of both the donor and the recipient.44 A relatively long learning curve is required before skills are acquired in the use of microsurgical equipment and success in the technique is achieved, and mortality can be high during the training period.15 Some authors set 50 transplantations as the threshold for perfecting the procedure.20 The duration of the surgery and time spent performing anastomoses impact on the appearance of complications and on the animal's survival.6,16,30,32,44

In general, the use of cuffs requires less skill and experience, and the procedure is faster than with sutures.38,45 Vascular cuffs reduce the risk of bleeding and with it the formation of granulation tissue, reducing the long-term risk of thrombosis or stenosis in the anastomosis sites.38 Bronchial leaks, a factor causing pneumothorax which compromises the functionality of the graft, are rarer with cuff anastomosis.35 However, the diameter of the cuff must be adequate to avoid reducing the size of the lumen, and moreover, a foreign-body reaction may occur.35,38,45 Modified cuffs, without extension handles or tabs, reduce the risk of twisting of the vessels or bronchi after anastomosis and injury caused by these extensions themselves. They are also smaller, so less foreign-body reaction is produced.36 Other perioperative complications include vena cava rupture, pleural empyema,34 respiratory arrest,38 and venous thrombosis.43

The most significant postoperative complications include respiratory failure, hemoptysis, thrombosis of a bronchial vein or artery, pneumothorax, respiratory infections, pleural effusion, pulmonary edema, and graft rejection.19,46

Primary graft dysfunction occurs in the first 72h and severity depends on the PaO2/FiO2 ratio and the presence of radiological infiltrates. It is a significant cause of morbidity and mortality in the immediate postoperative period.

Analysis of this phenomenon in mouse models has led to a better characterization of the causative mechanisms, in which the primary lesion of the neutrophils caused by ischemia–reperfusion appears to play a fundamental role. Thus, neutrophils isolated from the airways of transplant recipient have been seen to stimulate the donor's dendritic cells by a contact-dependent mechanism to induce IL-12 production and to expand alloantigen-specific T cells.50 This phenomenon, in turn, has been shown to be regulated by the degranulation induced by TNF-α associated with the neutrophil plasma membrane.50 Furthermore, some factors which modulate this cell-mediated response have been identified, which may be of potential interest in the design of future therapeutic approaches. The transcriptional factor Bc13 has been seen to limit granulopoiesis via a dependent NF-kB pathway.51

Mouse models provide an ideal medium for the study of acute cell-mediated rejection, as the grade of concordance or discordance of the major histocompatibility antigen can be evaluated in pure strains. This is of particular interest in the development of specific monitoring systems for this transplantation model. In addition to using bronchoalveolar lavages for the study of airway cellularity and measuring oxygenation capacity with arterial blood gases, specific imaging techniques have been developed, such as volumetric tomography53 or ultra-short magnetic resonance.54 These, together with the histological evaluation of the ultrastructural changes in acute rejection using scanning electron microscopy,55 improve the monitoring of the transplanted lung.

Complementary procedures have also been developed for the evaluation of lung function. The forced oscillation technique is very useful in mouse models, since dynamic elastance and its frequency dependence appear to have the ability to predict the grade of acute rejection,56 and can be used as non-invasive indices for detecting and monitoring disease progression. In an image-function setting, micro-positron emission tomography can be used to identify increased 18fluorodeoxyglucose uptake by neutrophils and CD8 lymphocytes,57 providing a more precise picture of acute rejection from the evaluation of T cell metabolism.

The evaluation of the pathogenic mechanisms of acute rejection in mouse models has fostered the development of interventions for prolonging graft survival, for example, by blocking anti-CD28-B7 costimulators and the CD40–CD40 ligand.58 Other authors have shown in an MHC-mismatched orthotopic mouse lung transplantation model that monotherapy with anti-CD154 antibodies may be sufficient to attenuate acute rejection59 by blocking CD154/CD40 costimulation, which fosters a shift in the graft environment toward a predominance of CD4(+) CD25(+) Foxp3(+) regulatory T cells.60 Some pulmonary nonhematopoietic cells have also been reported to play an important role in the in vivo suppression of CD4(+) T cell immune-mediated responses.61

Although it still early, these findings may be transferred to clinical practice in the form of N-acetylcysteine therapy. The administration of this compound in mouse models has been shown to diminish acute lung rejection by reducing the macrophage and T cell infiltration that is intimately related with reduction of the NF-kB-mediated inflammatory signaling pathway.

Chronic rejection usually manifests as the development of bronchiolitis obliterans, caused by lymphocyte and neutrophils cell infiltration attributable to a series of factors, such as excessive activation of the innate immune system, abnormal angiogenesis, or inadequate epithelial regeneration and remodeling of the lymphoproliferative tissue.48

Although each animal model has its advantages and limitations in the study of bronchiolitis obliterans, the mouse model of bronchiolitis obliterans associated with lung transplantation mimics more closely chronic rejection in humans than other models of bronchiolitis obliterans induced by chemical agents. Moreover, it can be used to make serial evaluations of both lung function and multiple morphological changes or altered cell function, and for testing the efficacy of different drugs. Another important advantage is the availability of models with knock-out and transgenic strains of multiple human diseases, that can be used to reproduce situations of chronic rejection relatively similar to those that occur in different end-stage respiratory diseases, and for evaluating the contribution of potential signaling pathways.

However, some anatomical and physiological factors of the mouse lung cause translational difficulties. Compared to human lungs, mouse bronchioles have fewer submucosal glands and a greater number of Clara cells. This environment protects against bronchiolar epithelial injury, primarily due the production of Clara cell secretory protein and a solution similar to lung surfactant, so more vascular injury than epithelial injury is generated in these rejection models.58 Nevertheless, significant advances have been made in recent years in the development of different bronchiolitis obliterans mouse models63–65 and DNA microarrays for evaluating chronic rejection after transplantation.66

In conclusion, techniques in mouse lung transplantation have developed significantly, contributing to a range of different surgical alternatives with excellent viability of transplanted animals. These experimental surgical models that have demonstrated their utility in generating knowledge on the pathogenic mechanisms of ischemia–reperfusion injury and rejection are now more widely accessible, and may be translated to humans in the medium term.