We thank all colleagues and professionals who contributed to and supported the construction of this ex vivo porcine lung model protocol.
This study had no funding sources.

The protocol was approved by the Animal Research Ethics Committee of our Institution (protocol no. 1610/2021).
1. Anesthesia and animal preparation
<ol>
	<li>Initially, place the animal on a scale and check the weight to adjust the medications and sedation necessary for the procedure.</li>
	<li>Administer ketamine 5 mg/kg and midazolam 0.25 mg/kg intramuscularly.</li>
	<li>Puncture the marginal ear vein with a 20 G venous catheter and administer intravenous propofol (5 mg/kg) for anesthesia induction.</li>
	<li>Administer 3 mL of heparin intravenously into the access to the marginal ear vein to assist in the cardiopulmonary extraction and perfusion.</li>
	<li>After anesthesia, perform orotracheal intubation with a 6.5 mm orotracheal cannula (OTC) and fix the OTC with adhesive tape, leaving it firmly fixed to avoid displacement during the procedure. 
	&#8203;NOTE: The depth of sedation is checked by monitoring hemodynamic parameters and using a gas analyzer, such as mean arterial pressure, heart rate, and inspired/expired isoflurane concentration.</li>
</ol>
2. Intraoperative mechanical ventilation
<ol>
	<li>Connect the animal via the OTC to MV, maintaining sedation with 1.5% isoflurane at 50% of the inspired fraction of oxygen (FiO2) and fentanyl 10 mcg/kg bolus + 10 mcg/kg/h continuous infusion.
	<ol>
		<li>Tap on the mechanical ventilator screen and select the volume-controlled ventilation (VCV) mode, select the tidal volume (TV) button, and turn the scroll wheel until the tidal volume value corresponds to 8 mL/kg.</li>
		<li>Tap on the mechanical ventilator screen. Select the FiO2 and turn the scroll wheel until the value of 50% is reached.</li>
		<li>Tap on the mechanical ventilator screen and select the respiratory rate (RR). Turn the wheel until it reaches the ideal value to maintain an end-expired CO2 of 35-45 mmHg measured by capnography coupled to the mechanical ventilator. 
		&#8203;NOTE: The depth of sedation is checked by monitoring hemodynamic parameters and using a gas analyzer such as mean arterial pressure, heart rate, and inspired/expired isoflurane concentration.</li>
	</ol>
	</li>
</ol>
3. Tissue dissection and OTC exchange
<ol>
	<li>Make a medial sternal incision from 2 cm above the manubrium to 2 cm below the xiphoid process of the sternum to access the thoracic cavity. Position the rib retractors, expanding the field of view during the procedure.</li>
	<li>Use a scalpel to make a horizontal tracheal incision at the height of the cricoid cartilage (just at the first tracheal rings) wide enough to introduce a new tracheal cannula.</li>
	<li>Deflate the OTC cuff that is inside the airway and pull slowly to remove it. Meanwhile, insert the new OTC into the incision made in the trachea after removing the old OTC. Leakage may occur due to the deflated cuff, ceasing when repositioning the new OTC.</li>
	<li>Inflate the newly inserted tracheal tube cuff by connecting a 20 mL syringe to the pilot balloon. The syringe delivers air under pressure and inflates the pilot balloon and cuff. Once the cuff inflates, remove the syringe.</li>
	<li>Tie the new tracheal cannula directly to the trachea with 2-0 polyester to prevent leakage and movement while placing the lung in the plexiglass ventilation box.</li>
	<li>With the scalpel, dissect the tissues to remove the cardiopulmonary organs from the thorax.</li>
</ol>
4. Animal euthanasia
<ol>
	<li>Increase the isoflurane concentration to 5% and administer 10 mL of 19.1% potassium chloride. Subsequently, check the absence of vital signs. 
	&#8203;NOTE: This procedure was performed according to the National Institutes of Health Guide for the Care and Use of Laboratory Animals.</li>
</ol>
5. Cardiopulmonary extraction
<ol>
	<li>After euthanasia, dissect the respiratory ligament to remove the lungs.</li>
	<li>After the tissue dissection, clamp the OTC with the appropriate Kelly forceps during the end of inspiration, keeping the lungs inflated.</li>
	<li>Disconnect the OTC from the mechanical ventilator, but keep it clamped.</li>
	<li>Section the aortic artery, position the aspirator inside the thoracic cavity to remove the extravasated blood, maintain visualization of the cavity while finishing the dissections, and free the organs to be removed from the thoracic cavity. 
	NOTE: The inferior pulmonary ligament should be carefully released to avoid pulmonary laceration.</li>
	<li>Remove the heart and lung from the ribcage with the OTC clamped, without separating them, and place them on a tray.</li>
</ol>
6. Cardiopulmonary preparation
<ol>
	<li>With the lung on a tray, cannulate the pulmonary artery with a large-bore single-lumen catheter and connect it to the infusion set to continuously administer 2,000 mL of cold 0.9% saline solution (SS) or until clear liquid flows from the aorta. 
	NOTE: SS should be administered at a normal rate, avoid squeezing the intravenous (IV) bag.</li>
	<li>After clearing the flow, suture the aortic artery with 2-0 polyester and administer another 100 mL of 0.9% SS. Close the single-lumen catheter outlet as the liquid will remain inside until the end of the experiment.</li>
	<li>Unclamp the OTC, note that the lungs will deflate and remain closed, ready to receive MV and the ARM.</li>
</ol>
7. MV inside an acrylic box
<ol>
	<li>After preparation, open the acrylic box and position the lungs vertically inside the box. Pass the OTC through the hole in the lid and connect the tracheal cannula to the mechanical ventilator. 
	NOTE: Make sure the tracheal cannula is firmly secured in the trachea.</li>
	<li>Select the Start Ventilation button.
	<ol>
		<li>Tap on the mechanical ventilator screen and select the mechanical ventilator for VCV.</li>
		<li>Tap on the VCV mode settings screen, and select the TV button, turn the wheel until it reaches the value of 6 mL/kg. Do the same to adjust PEEP to 5 cm H2O, FiO2 to 21%, RR to 15 breaths per minute, and inspiratory pause time to 10%.</li>
	</ol>
	</li>
</ol>
8. ARM
<ol>
	<li>To start the recruitment, increase PEEP from 5 cm H2O to 6 cm H2O and then increase it in step-by-step increments of 2 cm H2O until reaching 14 cm H2O.PEEP is increased using the on-screen button below the PEEP value displayed on the screen. Turn the wheel to increase the value.
	<ol>
		<li>For each PEEP, write down the peak pressure, plateau pressure, dynamic compliance, and airway resistance values displayed on the mechanical ventilator screen. Write down the driving pressure, which is the plateau pressure value minus the PEEP value adjusted at that time.</li>
	</ol>
	</li>
	<li>After reaching 14 cm H2O, reduce PEEP in step-by-step decrements of 2 cm H2O until reaching 6 cm H2O, then reduce it to 5 cm H2O. PEEP is reduced using the on-screen button below the PEEP value displayed on the screen. Turn the wheel to decrease the value.
	<ol>
		<li>For each PEEP, write down the peak pressure, plateau pressure, dynamic compliance, and airway resistance values displayed on the mechanical ventilator screen. Write down the driving pressure, which is the plateau pressure value minus the PEEP value adjusted at that time. 
		&#8203;NOTE: Maintain each PEEP value for 10 min during the increment and for 5 min at each step during the decrement.</li>
	</ol>
	</li>
</ol>
9. Cardiopulmonary maintenance
<ol>
	<li>At the end of the recruitment stage, gently clamp the tracheal cannula with the clamp during inspiration, keeping the lungs inflated. Open the acrylic box.</li>
	<li>Remove the lungs from the acrylic box and carefully place them in a glass container. 
	&#8203;NOTE: Make sure the tracheal cannula is firmly secured in the trachea.</li>
	<li>Pour 500 mL of 0.9% SS.</li>
	<li>Store it in the refrigerator in a plastic-wrapped glass container at a temperature of 2 to 8 &#176;C for 24 h.</li>
	<li>Repeat the steps 7, 8, and 9 for five consecutive days.</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64850/64850fig01.jpg" /> 
Figure 1: Study flowchart. <a href="https://www.jove.com/files/ftp_upload/64850/64850fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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The authors declare no conflicts of interest.

The described protocol is useful for producing an ex vivo porcine lung model under positive-pressure MV. It can be used for studying and teaching lung mechanics through visual feedback from the lungs during recruitment and analysis of the curves and values projected on the device screen. To achieve this result, pilot studies are needed to understand the behavior of the lungs outside the rib cage and to identify the need for adaptations.
We identified that the critical point was the fo...

Mechanical ventilation (MV) is widely used in intensive care units (ICUs) and surgical centers. Its monitoring is essential to help recognize asynchronies and prevent injuries for all patients, especially when the patient has serious lung injuries1,2,3,4,5,6. Monitoring respiratory mechanics can also contribute to the clinical understanding of the disease progression and therapeutic applications, such as the use of positive end-expiratory pressure (PEEP) or the alveolar recruitment maneuver (ARM). However, the use of these techniques requires a proficient understanding of curves and basic lung mechanics3,4.
Students, residents, and medical professionals feel insecure about MV management, from turning on the ventilator and initial adjustments to monitoring plateau and driving pressures, and this insecurity is associated with a lack of knowledge and adequate prior training7,8,9,10. We observed that professionals who participated in simulations and used a lung model reported greater confidence, understanding of the parameters, and understanding of the components of lung mechanics8,11,12.
Models for studying and training MV with test lungs, bellows, and pistons can simulate different pressures and volumes, as well as different lung mechanics conditions13,14,15. Computational and software models also contribute to the study of cardiopulmonary interaction by generating simulations that can be used to teach the principles of MV11 to health professionals16,17.
While computational models may present difficulties in representing pulmonary hysteresis16, models with test lung and bellows13,14,15can produce pressure-volume curves similar to the physiological curve and demonstrate pulmonary dynamics. As an advantage, the ex vivo porcine lung presents similar anatomy to humans18, also producing MV curves, pulmonary hysteresis, and providing visual feedback of the lungs inside the acrylic box during the lung mechanics analysis. Visual models are important and can help understand difficult-to-imagine components and concepts. Thus, ex vivo lung models represent a practical way of teaching.
Studies with ex vivo porcine lungs, such as those on MV with positive and negative pressure19,20,21, analysis of aerosol distribution22,23, pediatric simulations24, and lung perfusion25 can improve the knowledge on MV. Recent studies analyzing models in positive and negative pressure have shown that positive-pressure ventilation can lead to abrupt recruitment with greater local deformation, greater distension, hysteresis curve differences, and possible tissue lesions compared to negative pressure pressure19,20,21. Nevertheless, positive-pressure models are necessary because patients are under positive pressure during MV pressure19,20,21. The development of a lung model for preclinical studies opens possibilities for new research and applications, including MV teaching and training.
Here, we present an ex vivo porcine lung model for studying and training purposes. Our primary objective is to describe the steps for the generation of this ex vivo porcine lung model under positive-pressure MV. It can be used in the future to study and teach lung mechanics.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.9% Saline solution</td>
			<td></td>
			<td></td>
			<td>2500ml</td>
		</tr>
		<tr>
			<td>Anesthesia machine - Primus</td>
			<td>Drager</td>
			<td>REF 8603800-18</td>
			<td>Anesthesia work station used in the procedure</td>
		</tr>
		<tr>
			<td>Aspirator</td>
			<td></td>
			<td></td>
			<td>For blood aspiration from thorax</td>
		</tr>
		<tr>
			<td>Bedside Monitor - Life Scope</td>
			<td>Nihon Kohden</td>
			<td>BSM-7363</td>
			<td>Multiparameter monitor used during the procedure</td>
		</tr>
		<tr>
			<td>Bonney Tissue Forceps</td>
			<td></td>
			<td></td>
			<td>Any tissue forceps is suitable</td>
		</tr>
		<tr>
			<td>Disposable scalper, #23</td>
			<td></td>
			<td></td>
			<td>Any scalper is suitable</td>
		</tr>
		<tr>
			<td>Disposable syringe needles, 18G x 1 1/2&#34;, 23G x 1&#34;</td>
			<td>BD</td>
			<td>302814</td>
			<td>Widely available</td>
		</tr>
		<tr>
			<td>Disposable syringes, 10ml</td>
			<td></td>
			<td></td>
			<td>Widely available</td>
		</tr>
		<tr>
			<td>Electrosurgical unit - SS-501</td>
			<td>WEM</td>
			<td></td>
			<td>For cutting and coagulation during thorax incision</td>
		</tr>
		<tr>
			<td>Fentanyl</td>
			<td></td>
			<td></td>
			<td>10 mcg/kg bolus + 10 mcg/kg/hour continuous infusion</td>
		</tr>
		<tr>
			<td>Finochietto retractor</td>
			<td></td>
			<td></td>
			<td>Any finochietto retractor is suitable</td>
		</tr>
		<tr>
			<td>heparin</td>
			<td></td>
			<td></td>
			<td>3ml</td>
		</tr>
		<tr>
			<td>Infusion set</td>
			<td></td>
			<td></td>
			<td>Any infusion set is suitable</td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td></td>
			<td></td>
			<td>1.5%</td>
		</tr>
		<tr>
			<td>Kelly Forceps Curved</td>
			<td></td>
			<td></td>
			<td>Any kelly forceps is suitable</td>
		</tr>
		<tr>
			<td>Ketamine</td>
			<td></td>
			<td></td>
			<td>5mg/kg</td>
		</tr>
		<tr>
			<td>Lactated Ringer solution</td>
			<td></td>
			<td></td>
			<td>500ml</td>
		</tr>
		<tr>
			<td>Mechanical ventilator - Servo I</td>
			<td>Maquet</td>
			<td>REF 6449701</td>
			<td>Mechanical ventilator used in the procedure</td>
		</tr>
		<tr>
			<td>Metzenbaum Scissor (Straight and curved)</td>
			<td></td>
			<td></td>
			<td>Any metzenbaum scissor is suitable</td>
		</tr>
		<tr>
			<td>Midazolam</td>
			<td></td>
			<td></td>
			<td>0.25mg/kg</td>
		</tr>
		<tr>
			<td>Orotracheal intubation cannula, #6.5</td>
			<td>Rusch</td>
			<td>112282</td>
			<td>Widely available</td>
		</tr>
		<tr>
			<td>Plexiglass</td>
			<td></td>
			<td></td>
			<td>Custom made plexiglass box: 30x45x60cm</td>
		</tr>
		<tr>
			<td>Polyester suture, 2-0</td>
			<td></td>
			<td></td>
			<td>Widely available</td>
		</tr>
		<tr>
			<td>Potassium choride</td>
			<td></td>
			<td></td>
			<td>10 ml, 19.1% potassium chloride.</td>
		</tr>
		<tr>
			<td>propofol</td>
			<td></td>
			<td></td>
			<td>5mg/kg</td>
		</tr>
		<tr>
			<td>Three way stopcock</td>
			<td></td>
			<td></td>
			<td>Widely available</td>
		</tr>
		<tr>
			<td>Venous catheter, G20 x 1&#34;</td>
			<td>BD</td>
			<td>38183314</td>
			<td>Widely available</td>
		</tr>
	</tbody>
</table>

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.

We used five female pigs weighing between 23.4-26.9 kg and followed the described protocol for cardiopulmonary extraction and lung mechanics analysis. Our intention is that the model is useful for the study of lung mechanics by analyzing peak pressure, plateau pressure, resistance, driving pressure, and dynamic compliance variables collected directly from the mechanical ventilator screen. The model flowchart is shown in Figure 1.
The lungs were analyzed for five c...

Watch this Scientific Journal Video about Ex Vivo Porcine Experimental Model for Studying and Teaching Lung Mechanics at JoVE.com

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.

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 ...

ex-vivo-porcine-experimental-model-for-studying-teaching-lung

Research

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JoVE Journal

Medicine

1.3K Views.  Universidade de São Paulo. 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.

Video: Ex Vivo Porcine Experimental Model for Studying and Teaching Lung Mechanics

Steps for the Autologous Ex vivo Perfused Porcine Liver-kidney Experiment

The use of ex vivo perfused models can mimic the physiological conditions of the liver for short periods, but to maintain normal homeostasis for an extended perfusion period is challenging. We have added the kidney to our previous ex vivo perfused liver experiment model to reproduce a more accurate physiological state for prolonged experiments without using live animals. Five intact livers and kidneys were retrieved post-mortem from sacrificed pigs on different days and perfused for a minimum of 6 hr. Hourly arterial blood gases were obtained to analyze pH, lactate, glucose and renal parameters. The primary endpoint was to investigate the effect of adding one kidney to the model on the acid base balance, glucose, and electrolyte levels. The result of this liver-kidney experiment was compared to the results of five previous liver only perfusion models. In summary, with the addition of one kidney to the ex vivo liver circuit, hyperglycemia and metabolic acidosis were improved. In addition this model reproduces the physiological and metabolic responses of the liver sufficiently accurately to obviate the need for the use of live animals. The ex vivo liver-kidney perfusion model can be used as an alternative method in organ specific studies. It provides a disconnection from numerous systemic influences and allows specific and accurate adjustments of arterial and venous pressures and flow.

Steps for the Autologous Ex vivo Perfused Porcine Liver-kidney Experiment

Study of the animal organ physiology can be achieved by performing an experimental ex vivo perfusion system. The addition of a porcine kidney, as a homeostatic organ to our previously developed ex vivo liver perfusion model can be a principal step to achieve a better physiological environment.

The use of ex vivo perfused models can mimic the physiological conditions of the liver for short periods, but to maintain normal homeostasis for an ...

Procedure for Human Saphenous Veins Ex Vivo Perfusion and External Reinforcement

The mainstay of contemporary therapies for extensive occlusive arterial disease is venous bypass graft. However, its durability is threatened by intimal hyperplasia (IH) that eventually leads to vessel occlusion and graft failure. Mechanical forces, particularly low shear stress and high wall tension, are thought to initiate and to sustain these cellular and molecular changes, but their exact contribution remains to be unraveled. To selectively evaluate the role of pressure and shear stress on the biology of IH, an ex&#160;vivo perfusion system (EVPS) was created to perfuse segments of human saphenous veins under arterial regimen (high shear stress and high pressure). Further technical innovations allowed the simultaneous perfusion of two segments from the same vein, one reinforced with an external mesh. Veins were harvested using a no-touch technique and immediately transferred to the laboratory for assembly in the EVPS. One segment of the freshly isolated vein was not perfused (control, day 0). The two others segments were perfused for up to 7 days, one being completely sheltered with a 4 mm (diameter) external mesh. The pressure, flow velocity, and pulse rate were continuously monitored and adjusted to mimic the hemodynamic conditions prevailing in the femoral artery. Upon completion of the perfusion, veins were dismounted and used for histological and molecular analysis. Under ex&#160;vivo conditions, high pressure perfusion (arterial, mean = 100 mm Hg) is sufficient to generate IH and remodeling of human veins. These alterations are reduced in the presence of an external polyester mesh.

Procedure for Human Saphenous Veins Ex Vivo Perfusion and External Reinforcement

The mechanisms leading to the development of intimal hyperplasia (IH) and vein graft failure are still poorly understood. This study describes an ex&#160;vivo system to perfuse human veins under controlled flow and pressure. Furthermore the efficiency of external mesh reinforcement to limit the development of IH was evaluated.

The mainstay of contemporary therapies for extensive occlusive arterial disease is venous bypass graft. However, its durability is threatened by intimal ...

Method of Isolated Ex Vivo Lung Perfusion in a Rat Model: Lessons Learned from Developing a Rat EVLP Program

The number of acceptable donor lungs available for lung transplantation is severely limited due to poor quality. Ex-Vivo Lung Perfusion (EVLP) has allowed lung transplantation in humans to become more readily available by enabling the ability to assess organs and expand the donor pool. As this technology expands and improves, the ability to potentially evaluate and improve the quality of substandard lungs prior to transplant is a critical need. In order to more rigorously evaluate these approaches, a reproducible animal model needs to be established that would allow for testing of improved techniques and management of the donated lungs as well as to the lung-transplant recipient. In addition, an EVLP animal model of associated pathologies, e.g., ventilation induced lung injury (VILI), would provide a novel method to evaluate treatments for these pathologies. Here, we describe the development of a rat EVLP lung program and refinements to this method that allow for a reproducible model for future expansion. We also describe the application of this EVLP system to model VILI in rat lungs. The goal is to provide the research community with key information and &#8220;pearls of wisdom&#8221;/techniques that arose from trial and error and are critical to establishing an EVLP system that is robust and reproducible.

Method of Isolated Ex Vivo Lung Perfusion in a Rat Model: Lessons Learned from Developing a Rat EVLP Program

Ex-Vivo Lung Perfusion (EVLP) has allowed lung transplantation in humans to become more readily available by enabling the ability to assess organs and expand the donor pool. Here, we describe the development of a rat EVLP program and refinements that allow for a reproducible model for future expansion.

The number of acceptable donor lungs available for lung transplantation is severely limited due to poor quality. Ex-Vivo Lung Perfusion (EVLP) has allowed ...

An Ex vivo Model to Study Hormone Action in the Human Breast

The study of hormone action in the human breast has been hampered by lack of adequate model systems. Upon in vitro culture, primary mammary epithelial cells tend to lose hormone receptor expression. Widely used hormone receptor positive breast cancer cell lines are of limited relevance to the in vivo situation. Here, we describe an ex vivo model to study hormone action in the human breast. Fresh human breast tissue specimens from surgical discard material such as reduction mammoplasties or mammectomies are mechanically and enzymatically digested to obtain tissue fragments containing ducts and lobules and multiple stromal cell types. These tissue microstructures kept in basal medium without growth factors preserve their intercellular contacts, the tissue architecture, and remain hormone responsive for several days. They are readily processed for RNA and protein extraction, histological analysis or stored in freezing medium. Fluorescence activated cell sorting (FACS) can be used to enrich for specific cell populations. This protocol provides a straightforward, standard approach for translational studies with highly complex, varied human specimens.

An Ex vivo Model to Study Hormone Action in the Human Breast

We have developed a novel ex vivo model to study hormone action in the human breast. It is based on tissue microstructures isolated from surgical breast tissue specimens which preserve tissue architecture, intercellular interactions, and paracrine signaling.

The study of hormone action in the human breast has been hampered by lack of adequate model systems. Upon in vitro culture, primary mammary epithelial ...

Ex vivo Live Imaging of Lung Metastasis and Their Microenvironment

Metastasis is a major cause for cancer-related morbidity and mortality. Metastasis is a multistep process and due to its complexity, the exact cellular and molecular processes that govern metastatic dissemination and growth are still elusive. Live imaging allows visualization of the dynamic and spatial interactions of cells and their microenvironment. Solid tumors commonly metastasize to the lungs. However, the anatomical location of the lungs poses a challenge to intravital imaging. This protocol provides a relatively simple and quick method for ex vivo live imaging of the dynamic interactions between tumor cells and their surrounding stroma within lung metastasis. Using this method, the motility of cancer cells as well as interactions between cancer cells and stromal cells in their microenvironment can be visualized in real time for several hours. By using transgenic fluorescent reporter mice, a fluorescent cell line, injectable fluorescently labeled molecules and/or antibodies, multiple components of the lung microenvironment can be visualized, such as blood vessels and immune cells. To image the different cell types, a spinning disk confocal microscope that allows long-term continuous imaging with rapid, four-color image acquisition has been used. Time-lapse movies compiled from images collected over multiple positions and focal planes show interactions between live metastatic and immune cells for at least 4 hr. This technique can be further used to test chemotherapy or targeted therapy. Moreover, this method could be adapted for the study of other lung-related pathologies that may affect the lung microenvironment.

Ex vivo Live Imaging of Lung Metastasis and Their Microenvironment

We describe a relatively simple method for ex vivo live imaging of the tumor cell-stroma interactions within lung metastasis, utilizing fluorescent reporters in mice. Using spinning-disk confocal microscopy, this technique enables visualization of live cells for at least 4 hr and could be adapted to study other inflammatory lung conditions.

Metastasis is a major cause for cancer-related morbidity and mortality. Metastasis is a multistep process and due to its complexity, the exact cellular ...

A Small Animal Model of Ex Vivo Normothermic Liver Perfusion

There is a significant shortage of liver allografts available for transplantation, and in response the donor criteria have been expanded. As a result, normothermic ex vivo liver perfusion (NEVLP) has been introduced as a method to evaluate and modify organ function. NEVLP has many advantages in comparison to hypothermic and subnormothermic perfusion including reduced preservation injury, restoration of normal organ function under physiologic conditions, assessment of organ performance, and as a platform for organ repair, remodeling, and modification. Both murine and porcine NEVLP models have been described. We demonstrate a rat model of NEVLP and use this model to show one of its important applications &#8212; the use of a therapeutic molecule added to liver perfusate. Catalase is an endogenous reactive oxygen species (ROS) scavenger and has been demonstrated to decrease ischemia-reperfusion in the eye, brain, and lung. Pegylation has been shown to target catalase to the endothelium. Here, we added pegylated-catalase (PEG-CAT) to the base perfusate and demonstrated its ability to mitigate liver preservation injury. An advantage of our rodent NEVLP model is that it is inexpensive in comparison to larger animal models. A limitation of this study is that it does not currently include post-perfusion liver transplantation. Therefore, prediction of the function of the organ post-transplantation cannot be made with certainty. However, the rat liver transplant model is well established and certainly could be used in conjunction with this model. In conclusion, we have demonstrated an inexpensive, simple, easily replicable NEVLP model using rats. Applications of this model can include testing novel perfusates and perfusate additives, testing software designed for organ evaluation, and experiments designed to repair organs.

A Small Animal Model of Ex Vivo Normothermic Liver Perfusion

There is a significant liver donor shortage, and criteria for liver donors have been expanded. Normothermic ex vivo liver perfusion (NEVLP) has been developed to evaluate and modify organ function. This study demonstrates a rat model of NEVLP and tests the ability of pegylated-catalase, to mitigate liver preservation injury.

There is a significant shortage of liver allografts available for transplantation, and in response the donor criteria have been expanded. As a result, ...

Ex Vivo Corneal Organ Culture Model for Wound Healing Studies

The cornea has been used extensively as a model system to study wound healing. The ability to generate and utilize primary mammalian cells in two dimensional (2D) and three dimensional (3D) culture has generated a wealth of information not only about corneal biology but also about wound healing, myofibroblast biology, and scarring in general. The goal of the protocol is an assay system for quantifying myofibroblast development, which characterizes scarring. We demonstrate a corneal organ culture ex vivo model using pig eyes. In this anterior keratectomy wound, corneas still in the globe are wounded with a circular blade called a trephine. A plug of approximately 1/3 of the anterior cornea is removed including the epithelium, the basement membrane, and the anterior part of the stroma. After wounding, corneas are cut from the globe, mounted on a collagen/agar base, and cultured for two weeks in supplemented-serum free medium with stabilized vitamin C to augment cell proliferation and extracellular matrix secretion by resident fibroblasts. Activation of myofibroblasts in the anterior stroma is evident in the healed cornea. This model can be used to assay wound closure, the development of myofibroblasts and fibrotic markers, and for toxicology studies. In addition, the effects of small molecule inhibitors as well as lipid-mediated siRNA transfection for gene knockdown can be tested in this system.

A protocol for an ex vivo corneal organ culture model useful for wound healing studies is described. This model system can be used to assess the effects of agents to promote regenerative healing or drug toxicity in an organized 3D multicellular environment.

The cornea has been used extensively as a model system to study wound healing. The ability to generate and utilize primary mammalian cells in two ...

An Ex Vivo Tissue Culture Model for Fibrovascular Complications in Proliferative Diabetic Retinopathy

Diabetic retinopathy (DR) is the most common microvascular complication of diabetes and one of the leading causes of blindness in working-age adults. No current animal models of diabetes and oxygen-induced retinopathy develop the full-range progressive changes manifested in human proliferative diabetic retinopathy (PDR). Therefore, understanding of the disease pathogenesis and pathophysiology has relied largely on the use of histological sections and vitreous samples in approaches that only provide steady-state information on the involved pathogenic factors. Increasing evidence indicates that dynamic cell-cell and cell-extracellular matrix (ECM) interactions in the context of three-dimensional (3D) microenvironments are essential for the mechanistic and functional studies towards the development of new treatment strategies. Therefore, we hypothesized that the pathological fibrovascular tissue surgically excised from eyes with PDR could be utilized to reliably unravel the cellular and molecular mechanisms of this devastating disease and to test the potential for novel clinical interventions. Towards this end, we developed a novel method for 3D ex vivo culture of surgically-excised patient-derived fibrovascular tissue (FT), which will serve as a relevant model of human PDR pathophysiology. The FTs are dissected into explants and embedded in fibrin matrix for ex vivo culture and 3D characterization. Whole-mount immunofluorescence of the native FTs and end-point cultures allows thorough investigation of tissue composition and multicellular processes, highlighting the importance of 3D tissue-level characterization for uncovering relevant features of PDR pathophysiology. This model will allow the simultaneous assessment of molecular mechanisms, cellular/tissue processes and treatment responses in the complex context of dynamic biochemical and physical interactions within the PDR tissue architecture and microenvironment. Since this model recapitulates PDR pathophysiology, it will also be amenable for testing or developing new treatments.

Here, we present a protocol to study the pathophysiology of proliferative diabetic retinopathy by using patient-derived, surgically-excised, fibrovascular tissues for three-dimensional native tissue characterization and ex vivo culture. This ex vivo culture model is also amenable for testing or developing new treatments.

Diabetic retinopathy (DR) is the most common microvascular complication of diabetes and one of the leading causes of blindness in working-age adults. No ...

Manufacture of a Multi-Purpose Low-Cost Animal Bench-Model for Teaching Tracheostomy

Tracheostomy is one of the most frequent procedures, performed through various techniques in the intensive care unit and emergency situations. Despite this, there is a lack of training on this procedure that affects its outcome, which is also dependent on operator's dexterity. Here, we take the specific training and simulation into consideration. This article aims to describe every step of the manufacture of a new multi-purpose low-cost animal bench-model, with the support of video and images, and to obtain an opinion about the quality of this model by administering a questionnaire to professionals with experience in the procedures. 
Ten experts in the technique were enrolled. The model scored an average of 3.45/5 for its anatomical realism; 4.75/5 for its usefulness as a training tool for simulation courses and assessments. The time necessary to build the model was 15 minutes, and the cost amounted to 10&#8364;. The animal bench-model was considered a very useful simulator for tracheostomy training and assessments. Therefore, it could be used as a tool for medical courses and residencies.

This article illustrates every step of the manufacture of a new multi-purpose low-cost animal bench-model for subglottic airway access management. All the procedures are shown in the video. The model's realism and its suitability for training the given clinical maneuvers were assessed by independent senior otolaryngologists and anesthesiologists.

Tracheostomy is one of the most frequent procedures, performed through various techniques in the intensive care unit and emergency situations. Despite ...

Precision Cut Lung Slices as an Efficient Tool for Ex vivo Pulmonary Vessel Structure and Contractility Studies

The visualization of murine lung tissue provides valuable structural and cellular information regarding the underlying airway and vasculature. However, the preservation of pulmonary vessels that truly represents physiological conditions still presents challenges. In addition, the delicate configuration of murine lungs result in technical challenges preparing samples for high-quality images that preserve both cellular composition and architecture. Similarly, cellular contractility assays can be performed to study the potential of cells to respond to vasoconstrictors in vitro, but these assays do not reproduce the complex environment of the intact lung. In contrast to these technical issues, the precision-cut lung slice (PCLS) method can be applied as an efficient alternative to visualize lung tissue in three dimensions without regional bias and serve as a live surrogate contractility model for up to 10 days. Tissue prepared using PCLS has preserved structure and spatial orientation, making it ideal to study disease processes ex vivo. The location of endogenous tdTomato-labeled cells in PCLS harvested from an inducible tdTomato reporter murine model can be successfully visualized by confocal microscopy. After exposure to vasoconstrictors, PCLS demonstrates the preservation of both vessel contractility and lung structure, which can be captured by a time-lapse module. In combination with the other procedures, such as western blot and RNA analysis, PCLS can contribute to the comprehensive understanding of signaling cascades that underlie a wide variety of disorders and lead to a better understanding of the pathophysiology in pulmonary vascular diseases.

Precision Cut Lung Slices as an Efficient Tool for Ex vivo Pulmonary Vessel Structure and Contractility Studies

Presented here is a protocol for preserving the vascular contractility of PCLS murine lung tissue, resulting in a sophisticated three-dimensional image of the pulmonary vasculature and airway, which can be preserved for up to 10 days that is susceptible to numerous procedures.

The visualization of murine lung tissue provides valuable structural and cellular information regarding the underlying airway and vasculature. However, ...

Ex Vivo Porcine Experimental Model for Studying and Teaching Lung Mechanics

Summary

Explore More Videos

Chapters in this video

Steps for the Autologous Ex vivo Perfused Porcine Liver-kidney Experiment

Procedure for Human Saphenous Veins Ex Vivo Perfusion and External Reinforcement

Method of Isolated Ex Vivo Lung Perfusion in a Rat Model: Lessons Learned from Developing a Rat EVLP Program

An Ex vivo Model to Study Hormone Action in the Human Breast

Ex vivo Live Imaging of Lung Metastasis and Their Microenvironment

A Small Animal Model of Ex Vivo Normothermic Liver Perfusion

Ex Vivo Corneal Organ Culture Model for Wound Healing Studies

An Ex Vivo Tissue Culture Model for Fibrovascular Complications in Proliferative Diabetic Retinopathy

Manufacture of a Multi-Purpose Low-Cost Animal Bench-Model for Teaching Tracheostomy

Precision Cut Lung Slices as an Efficient Tool for Ex vivo Pulmonary Vessel Structure and Contractility Studies