Name: mouse pneumonectomy model of compensatory lung growth
Uploaded: 2014-12-17T14:00:00.000+00:00
Duration: 9 min 22 s
Description: Watch this Scientific Journal Video about Mouse Pneumonectomy Model of Compensatory Lung Growth at JoVE.com

The authors would like to acknowledge the Cincinnati Children&#8217;s Hospital Division of Veterinary Services for their assistance. This manuscript was supported by the National Institutes of Health K12 HD028827. Anna Perl PhD taught the authors this surgical procedure.

NOTE: Animal use statement: All procedures in this study were conducted with approval and following the guidelines of the Institutional Animal Use and Care Committee (IACUC) at Cincinnati Children&#8217;s Hospital. Eight week-old C57BL/6J male mice were obtained from Jackson Laboratories (Bar Harbor, ME) and allowed to acclimate for one week prior to use. Up until surgery, animals were housed in a pathogen-free barrier facility and provided autoclaved chow and filtered water ad libdium. Each mouse cage was supplied with a dedicated air and water, and rooms were maintained on a 12 hr day-night cycle. Following recovery from surgery, mice were maintained in cages with filtered tops, provided autoclaved chow ad libidum, and provided filtered water from a water bottle.
1. Preparation of Instruments
<ol>
	<li>Make 6 skin retractors using paper clips and pins. Twist straightened paper clips on the shanks of paper pins, leave a 5 cm straight steel wire on one end and make a single 0.5 cm &#8220;U&#8221; shaped hook at the end of the wire.</li>
	<li>Make some 15 x 15 cm square surgical drapes using plastic wrap. Prepare one dressing per mouse. Put a paper tower in between each wrap.</li>
	<li>Sterilize all surgical tools along with a stack of 12 x 12 inch cork tiles, gauze, and cotton-tipped swabs.</li>
</ol>
2. Mouse Preparation
<ol>
	<li>Induce anesthesia with 2% isoflurane. Weigh animal.</li>
	<li>In a dedicated surgical preparation area shave left thorax and neck area with electrical shaver.</li>
	<li>Apply a drop of the artificial tear ointment to the mouse&#8217;s eyes.</li>
	<li>Decontaminate neck and left thorax with chlorhexidine and isopropyl alcohol. Repeat twice more.</li>
</ol>
3. Mouse Oro-tracheal Intubation and Mechanical Ventilation
<ol>
	<li>Have a non-sterile surgical technician place the mouse supine in the pre-warmed surgical area.</li>
	<li>Confirm depth of anesthesia by documenting lack of a response to paw pinch.</li>
	<li>After washing hands and donning surgical attire, mask, and hat, don sterile surgical gloves.</li>
	<li>After draping and using aseptic technique, make a 1 cm vertical incision over the anterior mid-neck to expose the larynx. Lightly retract the strap muscles with curved, serrated 10 cm forceps and expose the larynx and trachea by spreading the strap muscles with the tip of a straight scissors.</li>
	<li>Orally insert a 22 G blunt-tip angiocatheter into the mid-trachea (Figure 1A) and visually confirm placement (Figure 1B).&#160;Maintain anesthesia and ventilate using 1-3% isoflurane through rodent ventilator (225 &#181;l per stroke; 200 stokes per min). Employ a pressure limit of 15 cm H2O.</li>
</ol>
4. Mouse Pneumonectomy
<ol>
	<li>Lay the mouse in the right lateral decubitus position with the mouse&#8217;s back facing the operator (left side up). Use a self-sealing plastic wrap as a sterile drape. Cutting through the drape, use blunt tipped curved scissors to make a 2 cm long cut parallel to the ribs at the 4th and 5th intercostal space. Insert the blunt tip curved scissors and dissect the skin away from the underlying ribs and intercostal muscles.</li>
	<li>Retract the skin with four retractors to expose a square 1.5 x 1.5 cm surgical window (Figure 2A). Secure the retractors to the cork board.</li>
	<li>Dissect down to ribs using curved forceps, and use one tip of the curved forceps to enter the thoracic cavity.</li>
	<li>Using the blunt tip micro-scissors, use the lower blade to enter the thoracic cavity. Make a 0.5 cm incision between the ribs and repeat in the opposite direction.</li>
	<li>Using the two remaining retractors, open the thorax in the anterior-posterior axis and the secure the retractors to the cork board (Figure 2B).</li>
	<li>Using curved blunt-tipped forceps in the left hand, grasp the left lung and displace the upper portion of the left lung laterally and inferiorly through the thoracotomy until the left pulmonary artery and bronchus are exposed (Figure 3A,B).</li>
	<li>Holding the loaded titanium vascular microclip applicator in the right hand with the body of the applicator in the palm and curved tip pointing away from the palm (Figure 3C), slide the applicator tip into the thorax along the curvature of the posterior aspect of the left lung and clip the left bronchus and pulmonary artery (Figure 3D).</li>
	<li>Remove the applicator but keep the left lung retracted. Grasp the blunt tip micro-scissors with the right hand and cut the bronchus and pulmonary artery distal to the clip and remove left lung (Figure 3E).</li>
	<li>Remove the rib retractors.</li>
	<li>Use the curved blunt forceps to pinch up 1 cm of skin inferior to the incision but above the level of the diaphragm and insert a 24 G angiocatheter through the skin and into the left thoracic cavity (Figure&#160;4A,B).</li>
	<li>Use 5-0 prolene suture to place two interrupted sutures around the 4th and 5th ribs to close the thoracic cavity.</li>
	<li>Remove the skin retractors. Use two sets of forceps to approximate the skin along the length of the incision and glue the skin closed.</li>
	<li>Connect a 3 ml luer-lock syringe to the angiocatheter and remove residual air by applying gentle suction and withdrawing the angiocatheter.</li>
	<li>Glue the neck incision closed using two sets of forceps as before.</li>
</ol>
5. Mouse Sham Pneumonectomy
<ol>
	<li>Expose the left lung as noted in &#8220;Mouse Pneumonectomy&#8221; protocol. Lift the rib cage with curved blunt forceps to allow air into the left chest cavity (Figure 5A,B).</li>
	<li>Place a 24 G angiocatheter into the left thoracic cavity as above being careful not to injure the left lung.</li>
	<li>Using 5-0 prolene suture and being careful not to puncture the lung (Figure 5C), place two lengths of suture material into the 3rd/4th and 5th/6th rib interspaces (Figure 5D). Place both lengths of suture material before tying to lessen the risks of left lung herniation. Tie the suture material to make two interrupted stitches (Figure 5E).</li>
	<li>Glue the skin over the thoracic incision, remove residual air with the angiocatheter, and glue the neck incision as above.</li>
</ol>
6. Resuscitation, Analgesia, and Recovery
<ol>
	<li>Turn off the isoflurane, and administer 0.1 mg/kg of buprenorphine and 0.5 ml of normal saline subcutaneously.</li>
	<li>When spontaneous respirations resume, remove the endotracheal tube.</li>
	<li>Observe mouse until it is again ambulatory. Walking typically resumes several minutes after removal of the endotracheal tube.</li>
	<li>Place the mouse in a 27 &#176;C incubator (humidified, 25% oxygen) to recover O/N. 
	NOTE: We place several pellets of chow wetted with water on the cage floor for the first 24 hr after surgery.</li>
	<li>Administer 0.1 mg/kg of buprenorphine by intraperitoneal injection twice daily for three days after surgery. Take care not to open the surgical site when handling animals.</li>
</ol>
7. Mouse Monitoring
<ol>
	<li>Weigh mice at 1, 3, 5, and 7 days after surgery.</li>
</ol>

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The authors have nothing to disclose.

We have provided the most detailed description of the mouse PNX and mouse sham PNX procedures reported to date. We have made the reader aware of several of the common pitfalls that investigators learning the procedure commonly encounter, and we have outlined several techniques developed by our laboratory to mitigate against these pitfalls. Other laboratories utilizing this model may have developed other technique modifications or use different instruments. When evaluating differences in techniques, individual investigato...

The principal function of the lung is to provide for oxygen and carbon dioxide exchange between an organism and the atmosphere. In humans, a host of congenital and acquired conditions lead to reduced lung surface area which results in impaired lung function. Although a host of therapies such as inhaled corticosteroids, bronchodilators, supplemental oxygen, and chronic mechanical ventilation are used to mitigate the consequences of impaired lung function1-3, the ideal therapy for these conditions would promote regrowth of functional lung tissue &#8211; i.e., lung regeneration.
Mammalian tissue regeneration has been well documented. The African Spiny Mouse can regenerate large areas of skin without scar formation4. The distal phalanx in humans can regenerate following injury or amputation5-7. Following pneumonectomy (PNX), compensatory lung growth occurs in mice8, rats9, dogs10, and humans11. By definition, compensatory lung growth involves not only expansion of existing airspaces, but re-septation of these enlarged airspaces with expansion of the associated microcirculation12. Gene expression analysis has demonstrated that this model recapitulates many of the signaling events of lung development13. Four weeks after mouse PNX, alveolar surface area is equivalent to that of sham operated animals14. In this manuscript, we describe the mouse PNX and sham PNX procedures.

<table><tbody><tr><td>6 inch Vascular clip applicator</td><td>Teleflex Medical (WECK)</td><td>137062</td><td></td></tr><tr><td>Horizon small titanium red clip</td><td>Teleflex Medical (WECK)</td><td>1201</td><td></td></tr><tr><td>Narrow pattern 12 cm curved forceps</td><td>Fine Science Tools</td><td>11003-12</td><td></td></tr><tr><td>Curved serrated 10 cm Graefe forceps</td><td>Fine Science Tools</td><td>11052-10</td><td></td></tr><tr><td>Castroviejo needle holder</td><td>Fine Science Tools</td><td>12565-14</td><td></td></tr><tr><td>Straight 9 cm Strabismus scissors (blunt tip)</td><td>Fine Science Tools</td><td>14075-09</td><td></td></tr><tr><td>Straight 8.5 cm hardened fine scissors</td><td>Fine Science Tools</td><td>14090-09</td><td></td></tr><tr><td>Straight, blunt tip Cohan-Vannas spring scissors</td><td>Fine Science Tools</td><td>15000-12</td><td></td></tr><tr><td>Skin glue</td><td>Gluture</td><td>32046</td><td></td></tr><tr><td>22 G Angiocatheter</td><td></td><td></td><td></td></tr><tr><td>24 G Angiocatheter</td><td></td><td></td><td></td></tr><tr><td>3 ml Luer lock syringe</td><td></td><td></td><td></td></tr><tr><td>4 Short retractors</td><td></td><td></td><td></td></tr><tr><td>2 Long retractors</td><td></td><td></td><td></td></tr><tr><td>5-0 Prolene on curved cutting needle</td><td>Ethicon</td><td>8698G</td><td></td></tr><tr><td>0.5 ml Syringe on 27 G needle</td><td></td><td></td><td></td></tr><tr><td>Normal saline</td><td></td><td></td><td></td></tr><tr><td>Buprenorphine</td><td></td><td></td><td></td></tr><tr><td>Press-n-Seal wrap</td><td>Glad Products Company</td><td></td><td></td></tr><tr><td>12 x 12 inch Cork board stack</td><td>Office Depot</td><td></td><td></td></tr><tr><td>70% Ethanol</td><td></td><td></td><td></td></tr><tr><td>Betadine</td><td></td><td></td><td></td></tr><tr><td>Mouse ventilator</td><td>Hugo Sachs Elektronnik</td><td>Minivent Type 845</td><td></td></tr><tr><td>Isoflurane vaporizer</td><td>OHMEDA</td><td>Excel 210 SE</td><td></td></tr><tr><td>Artificial tear ointment</td><td>Puralube</td><td>NDC: 17033-211-38</td><td></td></tr></tbody></table>

mouse pneumonectomy model of compensatory lung growth

In humans, disrupted repair and remodeling of injured lung contributes to a host of acute and chronic lung disorders which may ultimately lead to disability or death. Injury-based animal models of lung repair and regeneration are limited by injury-specific responses making it difficult to differentiate changes related to the injury response and injury resolution from changes related to lung repair and lung regeneration. However, use of animal models to identify these repair and regeneration signaling pathways is critical to the development of new therapies aimed at improving pulmonary function following lung injury. The mouse pneumonectomy model utilizes compensatory lung growth to isolate those repair and regeneration signals in order to more clearly define mechanisms of alveolar re-septation. Here, we describe our technique for performing mouse pneumonectomy and sham pneumonectomy. This technique may be utilized in conjunction with lineage tracing or other transgenic mouse models to define molecular and cellular mechanism of lung repair and regeneration.

A plot of PNX and sham operated mouse weights is provided in Figure 6. In our hands, survival is consistently 95 - 100% for both PNX and sham pneumonectomy. For descriptions of how the right lung re-grows in this model and the expected time course, we refer the reader to manuscripts of Gibney et al.15 and Wang et al.14
Several common pitfalls must be avoided to successfully perform the mouse PNX and mouse sham pneumonectomy procedures.</...

Watch this Scientific Journal Video about Mouse Pneumonectomy Model of Compensatory Lung Growth at JoVE.com

Mouse Pneumonectomy Model of Compensatory Lung Growth

Mouse pneumonectomy is a commonly employed model of compensatory lung growth. This procedure can be used in conjunction with lineage tracing or transgenic mouse models to elucidate underlying mechanisms.

In humans, disrupted repair and remodeling of injured lung contributes to a host of acute and chronic lung disorders which may ultimately lead to ...

mouse-pneumonectomy-model-of-compensatory-lung-growth

Research

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12.8K Views.  Cincinnati Children's Hospital Medical Center. Mouse pneumonectomy is a commonly employed model of compensatory lung growth. This procedure can be used in conjunction with lineage tracing or transgenic mouse models to elucidate underlying mechanisms.

Video: Mouse Pneumonectomy Model of Compensatory Lung Growth

Development of Obliterative Bronchiolitis in a Murine Model of Orthotopic Lung Transplantation

Orthotopic lung transplantation in rats was first reported by Asimacopoulos and colleagues in 1971 1. Currently, this method is well accepted and standardized not only for the study of allo-rejection but also between syngeneic strains for examining mechanisms of ischemia-reperfusion injury after lung transplantation. Although the application of the rat and other large animal model 2 contributed significantly to the elucidation of these studies, the scope of those investigations is limited by the scarcity of knockout and transgenic rats. Due to no effective therapies for obliterative bronchiolitis, the leading cause of death in lung transplant patients, there has been an intensive search for pre-clinical models that replicate obliterative bronchiolitis. The tracheal allograft model is the most widely used and may reproduce some of the histopathologic features of obliterative bronchiolitis 3. However, the lack of an intact vasculature with no connection to the recipient's conducting airways, and incomplete pathologic features of obliterative bronchiolitis limit the utility of this model 4. Unlike transplantation of other solid organs, vascularized mouse lung transplants have only recently been reported by Okazaki and colleagues for the first time in 2007 5. Applying the basic principles of the rat lung transplant, our lab initiated the obliterative bronchiolitis model using minor histoincompatible antigen murine orthotopic single-left lung transplants which allows the further study of obliterative bronchiolitis immunopathogenesis6.

Obliterative bronchiolitis is the key impediment to the long-term survival of lung transplant recipients and the lack of a robust preclinical model precludes examining obliterative bronchiolitis immunopathogenesis. Unlike other solid organ transplants, vascularized mouse lung transplantation has only recently been developed. Here we show our independently developed obliterative bronchiolitis model after murine orthotopic single-lung transplantation.

Orthotopic lung transplantation in rats was first reported by Asimacopoulos and colleagues in 1971 1. Currently, this method is well accepted and ...

A Standardized Method for Measuring Internal Lung Surface Area via Mouse Pneumonectomy and Prosthesis Implantation

Pulmonary morphology, physiology, and respiratory functions change in both physiological and pathological conditions. Internal lung surface area (ISA), representing the gas-exchange capacity of the lung, is a critical criterion to assess respiratory function. However, observer bias can significantly influence measured values for lung morphological parameters. The protocol that we describe here minimizes variations during measurements of two morphological parameters used for ISA calculation: internal lung volume (ILV) and mean linear intercept (MLI). Using ISA as a morphometric and functional parameter to determine the outcome of alveolar regeneration in both pneumonectomy (PNX) and prosthesis implantation mouse models, we found that the increased ISA following PNX treatment was significantly blocked by implantation of a prosthesis into the thoracic cavity1. The ability to accurately quantify ISA is not only expected to improve the reliability and reproducibility of lung function studies in injured-induced alveolar regeneration models, but also to promote mechanistic discoveries of multiple pulmonary diseases.

Internal lung surface area (ISA) is a critical criterion for assessing lung morphology and physiology in lung diseases and injury-induced alveolar regeneration. We describe here a standardized method that can minimize the measurement bias for ISA in both lung pneumonectomy and prosthesis implantation mouse models.

Pulmonary morphology, physiology, and respiratory functions change in both physiological and pathological conditions. Internal lung surface area (ISA), ...

Developmental Biology

Measurement of the Pressure-volume Curve in Mouse Lungs

In recent decades the mouse has become the primary animal model of a variety of lung diseases. In models of emphysema or fibrosis, the essential phenotypic changes are best assessed by measurement of the changes in lung elasticity. To best understand specific mechanisms underlying such pathologies in mice, it is essential to make functional measurements that can reflect the developing pathology. Although there are many ways to measure elasticity, the classical method is that of the total lung pressure-volume (PV) curve done over the whole range of lung volumes. This measurement has been made on adult lungs from nearly all mammalian species dating back almost 100 years, and such PV curves also played a major role in the discovery and understanding of the function of pulmonary surfactant in fetal lung development. Unfortunately, such total PV curves have not been widely reported in the mouse, despite the fact that they can provide useful information on the macroscopic effects of structural changes in the lung. Although partial PV curves measuring just the changes in lung volume are sometimes reported, without a measure of absolute volume, the nonlinear nature of the total PV curve makes these partial ones very difficult to interpret. In the present study, we describe a standardized way to measure the total PV curve. We have then tested the ability of these curves to detect changes in mouse lung structure in two common lung pathologies, emphysema and fibrosis. Results showed significant changes in several variables consistent with expected structural changes with these pathologies. This measurement of the lung PV curve in mice thus provides a straightforward means to monitor the progression of the pathophysiologic changes over time and the potential effect of therapeutic procedures.

Here we present a protocol to simply and reliably measure the lung pressure-volume curve in mice, showing that it is sufficiently sensitive to detect phenotypic parenchymal changes in two common lung pathologies, pulmonary fibrosis and emphysema. This metric provides a means to quantify the lung&#8217;s structural changes with developing pathology.

In recent decades the mouse has become the primary animal model of a variety of lung diseases. In models of emphysema or fibrosis, the essential ...

Generation of a Chronic Obstructive Pulmonary Disease Model in Mice by Repeated Ozone Exposure

Chronic obstructive pulmonary disease (COPD) is characterized by persistent airflow limitation and lung parenchymal destruction. It has a very high incidence in aging populations. The current conventional therapies for COPD focus mainly on symptom-modifying drugs; thus, the development of new therapies is urgently needed. Qualified animal models of COPD could help to characterize the underlying mechanisms and can be used for new drug screening. Current COPD models, such as lipopolysaccharide (LPS) or the porcine pancreatic elastase (PPE)-induced emphysema model, generate COPD-like lesions in the lungs and airways but do not otherwise resemble the pathogenesis of human COPD. A cigarette smoke (CS)-induced model remains one of the most popular because it not only simulates COPD-like lesions in the respiratory system, but it is also based on one of the main hazardous materials that causes COPD in humans. However, the time-consuming and labor-intensive aspects of the CS-induced model dramatically limit its application in new drug screening. In this study, we successfully generated a new COPD model by exposing mice to high levels of ozone. This model demonstrated the following: 1) decreased forced expiratory volume 25, 50, and 75/forced vital capacity (FEV25/FVC, FEV50/FVC, and FEV75/FVC), indicating the deterioration of lung function; 2) enlarged lung alveoli, with lung parenchymal destruction; 3) reduced fatigue time and distance; and 4) increased inflammation. Taken together, these data demonstrate that the ozone exposure (OE) model is a reliable animal model that is similar to humans because ozone overexposure is one of the etiological factors of COPD. Additionally, it only took 6 - 8 weeks, based on our previous work, to create an OE model, whereas it requires 3 - 12 months to induce the cigarette smoke model, indicating that the OE model might be a good choice for COPD research.

This study describes the successful generation of a new chronic obstructive pulmonary disease (COPD) animal model by repeatedly exposing mice to high concentrations of ozone.

Chronic obstructive pulmonary disease (COPD) is characterized by persistent airflow limitation and lung parenchymal destruction. It has a very high ...

The Left Pneumonectomy Combined with Monocrotaline or Sugen as a Model of Pulmonary Hypertension in Rats

In this protocol, we detail the correct procedural steps and necessary precautions to successfully perform a left pneumonectomy and induce PAH in rats with the additional administration of monocrotaline (MCT) or SU5416 (Sugen). We also compare these two models to other PAH models commonly used in research. In the last few years, the focus of animal PAH models has moved towards studying the mechanism of angioproliferation of plexiform lesions, in which the role of increased pulmonary blood flow is considered as an important trigger in the development of severe pulmonary vascular remodeling. One of the most promising rodent models of increased pulmonary flow is the unilateral left pneumonectomy combined with a "second hit" of MCT or Sugen. The removal of the left lung leads to increased and turbulent pulmonary blood flow and vascular remodeling. Currently, there is no detailed procedure of the pneumonectomy surgery in rats. This article details a step-by-step protocol of the pneumonectomy surgical procedure and post-operative care in male Sprague-Dawley rats. Briefly, the animal is anesthetized and the chest is opened. Once the left pulmonary artery, pulmonary vein, and bronchus are visualized, they are ligated and the left lung is removed. The chest then closed and the animal recovered. Blood is forced to circulate only on the right lung. This increased vascular pressure leads to a progressive remodeling and occlusion of small pulmonary arteries. The second hit of MCT or Sugen is used one week post-surgery to induce endothelial dysfunction. The combination of increased blood flow in the lung and endothelial dysfunction produces severe PAH. The primary limitation of this procedure is that it requires general surgical skills.

The rodent left pneumonectomy is a valuable technique in pulmonary hypertension research. Here, we present a protocol to describe the rat pneumonectomy procedure and postoperative care to ensure minimal morbidity and mortality.

In this protocol, we detail the correct procedural steps and necessary precautions to successfully perform a left pneumonectomy and induce PAH in rats ...

Experimental Model to Evaluate Resolution of Pneumonia

Acute respiratory distress syndrome (ARDS) causes acute lung injury, characterized by rapid alveolar damage and severe hypoxemia. This, in turn, leads to high morbidity and mortality. Currently, there are no pre-clinical models that recapitulate the complexity of human ARDS. However, infectious models of pneumonia (PNA) can replicate the main pathophysiological features of ARDS. Here, we describe a model of PNA induced by the intratracheal instillation of live Streptococcus pneumoniae and Klebsiella pneumoniae in C57BL6 mice. In order to evaluate and characterize the model, after inducing injury, we carried out serial measurements of body weight and bronchoalveolar lavage (BAL) for measuring markers of lung injury. Additionally, we harvested lungs for cell count and differentials, BAL protein quantification, cytospin, bacterial colony-forming unit counts, and histology. Lastly, high dimensional flow cytometry was performed. We propose this model as a tool to understand the immune landscape during the early and late resolution phases of lung injury.

This manuscript describes the establishment of an infectious model of pneumonia in mice and the respective characterization of injury resolution along with methods for growing bacteria and intratracheal instillation. A novel approach using high-dimensional flow cytometry to evaluate the immune landscape is also described.

Acute respiratory distress syndrome (ARDS) causes acute lung injury, characterized by rapid alveolar damage and severe hypoxemia. This, in turn, leads to ...

Immunology and Infection

A Rat Lung Transplantation Model of Warm Ischemia/Reperfusion Injury: Optimizations to Improve Outcomes

From our experience with rat lung transplantation, we have found several areas for improvement. Information in the existing literature regarding methods for choosing appropriate cuff sizes for the pulmonary vein (PV), pulmonary artery (PA), or bronchus (Br) are varied, thus making the determination of proper cuff size during rat lung transplantation an exercise of trial and error. By standardizing the cuffing technique to use the smallest effective cuff appropriate for the size of the vessel or bronchus, one can make the transplantation procedure safer, faster, and more successful. Since diameters of the PV, PA, and Br are related to the body weight of the rat, we present a strategy to choosing an appropriate size using a weight-based guide. Since lung volume is also related to body weight, we recommend that this relationship should also be considered when choosing the proper volume of air for donor lung inflation during warm ischemia as well as for the proper volume of PBS to be instilled during bronchoalveolar lavage (BAL) fluid collection. We also describe methods for 4th intercostal space dissection, wound closure, and sample collection from both the native and transplanted lobes.

Here, we present optimizations to a rat lung transplantation model that serve to improve outcomes. We provide a size guide for cuffs based on body weight, a measurement strategy to ascertain the 4th intercostal space, and methods of wound closure and BAL (bronchoalveolar lavage) fluid and tissue collection.

From our experience with rat lung transplantation, we have found several areas for improvement. Information in the existing literature regarding methods ...

Normothermic Negative Pressure Ventilation Ex Situ Lung Perfusion: Evaluation of Lung Function and Metabolism

Lung transplantation (LTx) remains the standard of care for end-stage lung disease. A shortage of suitable donor organs and concerns over donor organ quality exacerbated by excessive geographic transportation distance and stringent donor organ acceptance criteria pose limitations to current LTx efforts. Ex situ lung perfusion (ESLP) is an innovative technology that has shown promise in attenuating these limitations. The physiologic ventilation and perfusion of the lungs outside of the inflammatory milieu of the donor body affords ESLP several advantages over traditional cold static preservation (CSP). There is evidence that negative pressure ventilation (NPV) ESLP is superior to positive pressure ventilation (PPV) ESLP, with PPV inducing more significant ventilator-induced lung injury, pro-inflammatory cytokine production, pulmonary edema, and bullae formation. The NPV advantage is perhaps due to the homogenous distribution of intrathoracic pressure across the entire lung surface. The clinical safety and feasibility of a custom NPV-ESLP device have been demonstrated in a recent clinical trial involving extender criteria donor (ECD) human lungs. Herein, the use of this custom device is described in a juvenile porcine model of normothermic NPV-ESLP over a 12 h duration, paying particular attention to management techniques. Pre-surgical preparation, including ESLP software initialization, priming, and de-airing of the ESLP circuit, and the addition of anti-thrombotic, anti-microbial, and anti-inflammatory agents, is specified. The intraoperative techniques of central line insertion, lung biopsy, exsanguination, blood collection, cardiectomy, and pneumonectomy are described. Furthermore, particular focus is paid to anesthetic considerations, with anesthesia induction, maintenance, and dynamic modifications outlined. The protocol also specifies the custom device's initialization, maintenance, and termination of perfusion and ventilation. Dynamic organ management techniques, including alterations in ventilation and metabolic parameters to optimize organ function, are thoroughly described. Finally, the physiological and metabolic assessment of lung function is characterized and depicted in the representative results.

Normothermic Negative Pressure Ventilation Ex Situ Lung Perfusion: Evaluation of Lung Function and Metabolism

This paper describes a porcine model of negative pressure ventilation ex situ lung perfusion, including procurement, attachment, and management on the custom-made platform. Focus is made on anesthetic and surgical techniques, as well as troubleshooting.

Lung transplantation (LTx) remains the standard of care for end-stage lung disease. A shortage of suitable donor organs and concerns over donor organ ...

A Simple Cuff Technique for Murine Left Lung Transplantation

Over the past decade, our laboratory has made significant progress in developing and refining vascularized mouse lung transplantation models using an efficient and highly reliable "cuff technique" of transplantation. This article describes a sophisticated and comprehensive method for orthotopic lung transplantation in a vascularized orthotopic lung model, representing the most physiologic and clinically relevant model of mouse lung transplantation to date. The transplantation process consists of two distinct stages: donor harvest and subsequent implantation into the recipient. The method has been successfully mastered, and with several months of sufficient training, a skilled practitioner can perform the procedure in approximately 90 min from skin-to-skin. Surprisingly, once individuals overcome the initial learning curve, the survival rate during the perioperative period approaches nearly 100%. The mouse model allows for the use of multiple commercially available transgenic and mutant strains of mice, enabling the study of tolerance and rejection. Additionally, the unique features of this model make it a valuable tool for investigating tumor biology and immunology.

The present protocol describes a cuff technique for a mouse left lung transplantation model. This technique has been developed over several years and has performed well, serving effectively in immunological research.

Over the past decade, our laboratory has made significant progress in developing and refining vascularized mouse lung transplantation models using an ...

Ex Vivo Porcine Experimental Model for Studying and Teaching Lung Mechanics

Mechanical ventilation is widely used and requires specific knowledge for understanding and management. Health professionals in this field may feel insecure and lack knowledge because of inadequate training and teaching methods. Therefore, the objective of this article is to outline the steps involved in generating an ex vivo porcine lung model to be used in the future, to study and teach lung mechanics. To generate the model, five porcine lungs were carefully removed from the thorax following the guidelines of the Animal Research Ethics Committee with adequate care and were connected to the mechanical ventilator through a tracheal cannula. These lungs were then subjected to the alveolar recruitment maneuver. Respiratory mechanics parameters were recorded, and video cameras were used to obtain videos of the lungs during this process. This process was repeated for five consecutive days. When not used, the lungs were kept refrigerated. The model showed different lung mechanics after the alveolar recruitment maneuver every day; not being influenced by the days, only by the maneuver. Therefore, we conclude that the ex vivo lung model can provide a better understanding of lung mechanics and its effects, and even of the alveolar recruitment maneuver through visual feedback during all stages of the process.

Ex Vivo Porcine Experimental Model for Studying and Teaching Lung Mechanics

We present an ex vivo pig lung model for the demonstration of pulmonary mechanics and alveolar recruitment maneuvers for teaching purposes. The lungs can be used for more than one day (up to five days) with minimal changes in pulmonary mechanics variables.

Mechanical ventilation is widely used and requires specific knowledge for understanding and management. Health professionals in this field may feel ...

Mouse Pneumonectomy Model of Compensatory Lung Growth

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Development of Obliterative Bronchiolitis in a Murine Model of Orthotopic Lung Transplantation

A Standardized Method for Measuring Internal Lung Surface Area via Mouse Pneumonectomy and Prosthesis Implantation

Measurement of the Pressure-volume Curve in Mouse Lungs

Generation of a Chronic Obstructive Pulmonary Disease Model in Mice by Repeated Ozone Exposure

The Left Pneumonectomy Combined with Monocrotaline or Sugen as a Model of Pulmonary Hypertension in Rats

Experimental Model to Evaluate Resolution of Pneumonia

A Rat Lung Transplantation Model of Warm Ischemia/Reperfusion Injury: Optimizations to Improve Outcomes

Normothermic Negative Pressure Ventilation Ex Situ Lung Perfusion: Evaluation of Lung Function and Metabolism

A Simple Cuff Technique for Murine Left Lung Transplantation

Ex Vivo Porcine Experimental Model for Studying and Teaching Lung Mechanics