Support was provided, in part, by a Merit Review Award (101 BX003482) from the U.S. Department of Veteran Affairs Biomedical Laboratory R&#38;D Service, a NIH grant (5R01 HL123227), a Transformative Project Award (962204) from the American Heart Association, and by institutional funds awarded to Dr. Riess. Dr. Balzer received unrelated funding from the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation), project number BA 6287/1-1. The authors would like to thank Matthew D. Olsen, Chun Zhou, Zhu Li, and Rebecca C. Riess for their valuable contributions to the study.

The in vivo portion of the experiments (from general anesthesia to euthanasia) requires prior approval by the respective Institutional Animal Care and Use Committee (IACUC). All procedures described herein were approved (protocol number M1700168) by the IACUC at Vanderbilt University Medical Center, Nashville, Tennessee, and were performed in compliance with the ARRIVE guidelines14. Prior to experimentation, all the rats were housed in the institute&#39;s animal care facility, with free access to water and food. Including different studies outside the purview of this manuscript, we have used a total of 148 male Sprague Dawley rats, 7-10 weeks old, with a weight between 250 g and 400 g so far.
1. Apparatus construction
NOTE: All parts, including respective manufacturers, are listed in the Table of Materials.
<ol>
	<li>To hold each piece of equipment in place, construct a custom-built lattice to allow for easy configuration and incorporation of the new devices (Figure 1). Attach aluminum poles (1 to 2 ft in length, 1 cm in diameter, available at any hardware store) to each other using cross clamps to create a 3D lattice, and place on a plastic tray (30 in x 21 in) to prevent spillage of fluids.</li>
	<li>Mount pressure transducers at an equal height to the lungs and connect them to the lines for the pulmonary artery (PA), pulmonary vein (PV), and to the ventilation tube. 
	NOTE: These transducers transmit data to their respective bioamplifiers connected to the data acquisition system (DAQ) and its software.</li>
	<li>Rather than using commercially available cannulas, craft custom cannulas (Figure 2) from segments of 2 mm wide, hard plastic tubing that are flared at the end, using an open flame to allow for the secure tying-off of sutures. Bend them into a U-shape to reduce stress on the lungs during hanging and to fit in the lung chamber.</li>
	<li>To make the chamber the lungs are to be ventilated and perfused in, place a 1,000 mL beaker inside a 1,500 mL beaker with a water bath between the two and inside the 1,000 mL beaker, creating a double boiler. Place these beakers on a heating plate set to 48 &#176;C, creating a chamber for the lungs that is both humid and resistant to fluctuations in temperature.</li>
	<li>Keep the buffer for the experiment in a 150 mL volumetric flask placed on a heating plate set to 37 &#176;C. Use a magnetic stirring bar to circulate the buffer inside the beaker. Set the meniscus of the buffer at a height such that it is 4 cm above the lung, to create an innate 4 cmH2O of pressure on the PV. During the surgery, ensure that the animal&#39;s lungs are at the same height as the buffer to reduce hydrostatic edema formation. 
	NOTE: Using a flask minimizes the amount of surface area in contact with room air, which minimizes gas diffusion.</li>
	<li>Use a roller pump to move the buffer through a circuit consisting of a heating coil and an air trap prior to perfusing the lung. Recycle the effluent from the PV back into the volumetric flask. Connect both the heating coil and air trap to a circulating water bath also set to 37 &#176;C. Adjust the temperature of the water bath according to the pump speed, so the perfusate has a constant temperature of 37 &#176;C.</li>
</ol>
2. Procedure
<ol>
	<li>Prior to the start of the experiment, prepare the setup.
	<ol>
		<li>Ensure that the software is running (see below) and properly collecting data.</li>
		<li>Calibrate all pressure transducers daily to ensure they are not drifting.</li>
		<li>Prepare the buffer (see Table 1 for ingredients for a physiological saline solution with 4% bovine serum albumin [BSA]) and ensure that the pH is at 7.4, using HCl to adjust accordingly. Set up the perfusion system with warmed buffer (Table 1) circulating throughout, ensuring that no air bubbles are present. Add BSA at least 30 min prior to the start of the experiment to give it sufficient time to dissolve. In the absence of an oxygenator, bubble gas into the perfusion buffer prior to the addition of BSA with a gas composition of 65% N2, 30% O2, and 5% CO2 to mimic the CO2 levels of in vivo lungs. This prevents the solution from foaming once BSA is added.</li>
		<li>Prepare the operating area with all necessary surgical tools, sutures, and tape. Incline the operating board at a 15&#176; angle, so that the anesthetized rat can be positioned with its head elevated above the rest of the body, and the trachea and heart-lung block can be manipulated easily.</li>
		<li>Wearing proper personal protective equipment, weigh the rat and give an intraperitoneal injection of pentobarbital (65 mg/kg-1). After 10 min, use a toe pinch to ensure that the surgical plane of general anesthesia has been reached. Administer more anesthetic if necessary.</li>
	</ol>
	</li>
	<li>Transfer the rat to the operating area and fixate it to the operating board by taping its front legs separately, followed by the hind legs together, making sure that the front legs are taped loose enough to allow a pain reflex to be visualized should anesthesia not be deep enough (Figure 3A).</li>
	<li>Secure the head of the rat by taping the mouth with a long, thin strip behind the front teeth, without restricting the tongue or airflow for the still spontaneously breathing rat (Figure 3B).</li>
	<li>Perform a tracheostomy by pinching the skin above the trachea using forceps and cutting with surgical scissors. Using curved forceps, bluntly dissect the muscle and tissue to reach the trachea. Ensure that there is no bleeding during this step.
	<ol>
		<li>Pass curved forceps underneath the trachea and open them to allow room to pass 3-0 sutures underneath that can then be pre-tied into a box knot (Figure 3B). Make a small incision between the cartilage rings of the trachea and insert the tracheal cannula (modified 18 G needle with 1 mm diameter tubing glued around the tip to create a notch). Tie off the suture, ensuring that no air can escape from the tracheal incision and that the cannula is not putting any strain on the trachea.</li>
	</ol>
	</li>
	<li>Set the ventilator to run with a gas mixture of 30% O2, 5% CO2, and 65% N2, with a tidal volume (VT) of 10 mL/kg, a positive end expiratory pressure (PEEP) of 0 cmH2O, and a rate of 60 breaths/min.</li>
	<li>Once the tracheal cannula is secured with the suture, begin ventilating the rat with the above settings. 
	NOTE: The surgical method used was adapted from Nelson et al.9 and is described step-by-step.</li>
	<li>Remove the fur from the abdomen of the rat using large surgical scissors and forceps. 
	NOTE: Hair removal ointments can also be used prior to the start of the experiment.</li>
	<li>While holding the xiphoid process with forceps, make a small horizontal incision below the ribs, ensuring that the diaphragm stays intact. Once the internal organs can safely be visualized, widen the horizontal cut to expose the entire diaphragm.</li>
	<li>Taking extreme care to avoid puncturing the lungs, inject heparin (3,000 U/kg-1) into the inferior vena cava (IVC) with a 22 G syringe.</li>
	<li>Again, hold the xiphoid process with forceps and cut cranially along the sternum while constantly visualizing the lungs to prevent cutting them.</li>
	<li>To properly see the heart-lung block, spread the rib cage apart using two large forceps and be sure to avoid breaking the ribs, which can result in a sharp bone fragment puncturing the lung (Figure 3C). 
	NOTE: Once the rib cage has been opened, use a PEEP of 2-3 cmH2O, set by a water lock attached to the expiratory limb of the ventilator, to avoid atelectasis.</li>
	<li>At this time, cut the IVC to euthanize the rat by exsanguination. Carefully ensure that at least 1 min has passed since the injection of heparin (see step 2.9) to give it enough time to circulate and prevent microclots in the lungs.</li>
	<li>When placing the perfusion cannulas, constantly check the pressures to make sure that no sudden changes occur. 
	NOTE: A sudden spike in pressure while the pump is running at a constant speed indicates blockage formation, while a sudden decrease can indicate a leakage.</li>
	<li>Trim any excess thymus to allow for easier visualization of the pulmonary vasculature. Locate the PA, pass small, curved forceps underneath, and again pass a 3-0 suture underneath and pre-tie into a box knot.</li>
	<li>Make a small incision into the right ventricle of the heart and insert the PA cannula (made from 2 mm diameter plastic tubing flared at the end using an open flame to allow for sutures to be tied around), being gentle to avoid rupturing the adjacent artery. Secure the cannula using the suture and perfuse, starting at 1.5 mL/min-1 (Figure 3D).</li>
	<li>Immediately excise the apex of the heart to avoid a pressure buildup in the lungs. Using small, curved forceps, rupture the mitral valve and visually ensure that the forceps are able to enter the left atrium unrestricted.</li>
	<li>Place a pre-tied 3-0 suture around the heart below the atrium (Figure 3E).</li>
	<li>Insert the PV cannula (similar construction to the PA cannula) into the left atrium and ensure that buffer can flow out of it prior to tying off the suture. 
	NOTE: It is critical that the PV cannula is placed past the mitral valve to ensure that it can be properly tied off, while not being placed too deep as to injure the PV or encumber flow (Figure 3E). Take extra care with tying the suture, as tying it too loose results in lost flow, while tying it too tight can damage the heart, weakening the placement of the cannula and potentially causing leakages.</li>
	<li>Trim excess tissue between the thoracic cavity and the tracheostomy incision using blunt-tipped scissors to avoid damaging the trachea. Ensure that the entire trachea below the tracheal cannula and the entire heart-lung block are visible.</li>
	<li>Remove the heart-lung block and trachea by holding the tracheal cannula and excising connective tissue behind the trachea with a pair of curved, blunt-tipped scissors, leaving the esophagus attached for extra structural support (Figure 3F).</li>
	<li>Gradually increase the flow rate to a maximum flow of 40 mL/kg-1/min-1. Allow the first 50 mL of buffer to flow out of the system to remove any inflammatory cytokines that may be released (Figure 3G).</li>
	<li>Gently move the isolated lungs to the double boiler chamber created with the two beakers. Make sure that the PA, PV, or tracheal cannula do not become twisted at any point while moving the lungs or hanging them. 
	&#8203;NOTE: Prevention of atelectasis during lung retrieval and connection to the setup is important, especially when EVLP is used for already damaged lungs or when interventions prior to EVLP are planned. This can be achieved by, for example, a small clip on the trachea after inspiration before transfer to the ventilator15.</li>
</ol>
3. Data acquisition
<ol>
	<li>For data collection, use any commercially available analog-to-digital converter and DAQ system.</li>
	<li>Pay attention that the sampling frequency is sufficient for the given purpose (200 Hz here) and that all relevant dependent parameters from bioamplifiers and other input devices are collected concurrently (airway pressure, PA, and PV pressure chosen for this protocol)16.</li>
	<li>Record independent parameters (e.g., perfusion speed, ventilation rate, ventilated gas composition, buffer electrolyte composition, and lung weight).</li>
</ol>

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L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ex+vivo+perfusion+with+adenosine+A2A+receptor+agonist+enhances+rehabilitation+of+murine+donor+lungs+after+circulatory+death.">Ex vivo perfusion with adenosine A2A receptor agonist enhances rehabilitation of murine donor lungs after circulatory death.</a> Transplantation. 99 (12), 2494-2503 (2015).</li><li>Valenza, F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+consumption+of+glucose+during+ex+vivo+lung+perfusion+correlates+with+lung+edema.">The consumption of glucose during ex vivo lung perfusion correlates with lung edema.</a> Transplantation Proceedings. 43 (4), 993-996 (2011).</li><li>Sayner, S. 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The authors declare no conflicts of interest, financial or otherwise.

More than 100 experiments have been successfully performed in our lab using this setup. The modular design of this customized setup gave great flexibility to potential changes in experimental requirements. While other setups utilize a deoxygenator18 to mimic constant oxygen consumption and CO2 production by end organs, this simplified model did not employ this feature, due to the focus on studying the effects of different gas compositions on pulmonary vascular tone. This approach, in wh...

Ex vivo lung perfusion (EVLP) techniques1 have seen a rise in usage in the past decade as a means of studying lung transplantations2, ischemia/reperfusion3, lung metabolism4, and immune responses5. Isolated, but intact, ventilated and perfused lungs offer the critically important ability to directly assess the response of the lungs, including the pulmonary vasculature, to potential interventions and/or therapeutics without potential confounders, such as neuronal and hormonal input or changing hemodynamics in vivo. At the same time, they maintain the physiological interplay of ventilation and perfusion, in contrast to in vitro conditions. A proposal looking at immune responses in lungs5, for example, needs the same quality of data as a study focused on increasing the donor pool size6 for lung transplantations. EVLP can be used across a variety of species, including mice3, rats7,8,9,10,11,12, pigs13, and humans2. Therefore, it is necessary to establish a model that can produce reliable data from a variety of different experimental parameters. Clinical relevance will be generated in subsequent studies using the EVLP model as a tool.
While commercial setups are available for purchase for most species, they can often be cost-prohibitive and confine researchers to a specific brand of equipment and proprietary software. Any deviation from the out-of-the-box setup (e.g., going from one species to another) requires foresight and working around the provided setup, which may prove to be difficult or impossible. In the following, a DIY (do it yourself) setup for rat isolated lungs that is both modular and cost-effective, as well as the surgical procedure for isolating the lungs, are described.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1,000 mL Glass Beaker</td>
			<td>Pyrex, Chicago, IL</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>1,500 mL Glass Beaker</td>
			<td>Pyrex, Chicago, IL</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Air Trap Compliance Chamber</td>
			<td>Radnoti</td>
			<td>130149</td>
			<td></td>
		</tr>
		<tr>
			<td>Bioamplifiers</td>
			<td>CWE Inc</td>
			<td>BPM-832</td>
			<td></td>
		</tr>
		<tr>
			<td>Clamps</td>
			<td>Fisher Scientific</td>
			<td>S02626</td>
			<td></td>
		</tr>
		<tr>
			<td>DAQ (Data Acquisition)</td>
			<td>National Instruments, Austin, TX</td>
			<td>NI USB-6343</td>
			<td></td>
		</tr>
		<tr>
			<td>Gas Mixer</td>
			<td>CWE Inc, Ardmore, PA</td>
			<td>GSM-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Heating Coil</td>
			<td>Radnoti, Covina, CA</td>
			<td>158822</td>
			<td></td>
		</tr>
		<tr>
			<td>Heating Plate</td>
			<td>Thermo Fisher Scientific, Waltham, MA</td>
			<td>11-100-49SH</td>
			<td></td>
		</tr>
		<tr>
			<td>Heparin</td>
			<td>Pfizer</td>
			<td>W63422</td>
			<td></td>
		</tr>
		<tr>
			<td>LabVIEW Full Development System 2014</td>
			<td>National Instruments</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Pentobarbital</td>
			<td>Diamondback Drugs</td>
			<td>G2270-0235-50</td>
			<td></td>
		</tr>
		<tr>
			<td>pH700 Probe</td>
			<td>OAKTON, Vernon Hills, IL&#160;</td>
			<td>EW-35419-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Polystat Water Bath</td>
			<td>Cole-Parmer</td>
			<td>EW-12121-02</td>
			<td></td>
		</tr>
		<tr>
			<td>Rodent Ventilator</td>
			<td>Harvard Apparatus, Holliston, MA</td>
			<td>Model 683</td>
			<td></td>
		</tr>
		<tr>
			<td>Roller Pump</td>
			<td>Cole-Parmer, Wertheim, Germany&#160;</td>
			<td>Ismatec REGLO Digital MS 2/8</td>
			<td></td>
		</tr>
		<tr>
			<td>Sprague Dawley Rat</td>
			<td>Charles River, Wilmington, MA</td>
			<td>Strain code 001</td>
			<td></td>
		</tr>
		<tr>
			<td>VetScan i-STAT</td>
			<td>Abraxis, Chicago, IL</td>
			<td>i-STAT 1</td>
			<td></td>
		</tr>
	</tbody>
</table>

design and implementation of a rat ex vivo lung perfusion model

Ex vivo lung preparations are a useful model that can be translated to many different fields of research, complementing corresponding in vivo and in vitro models. Laboratories wishing to use isolated lungs need to be aware of important steps and inherent challenges to establish a setup that is affordable, reliable, and that can be easily adapted to fit the topic of interest. This paper describes a DIY (do it yourself) model for ex vivo rat lung ventilation and perfusion to study drug and gas effects on pulmonary vascular tone, independent of changes in cardiac output. Creating this model includes a) the design and construction of the apparatus, and b) the lung isolation procedure. This model results in a setup that is more cost-effective than commercial alternatives and yet modular enough to adapt to changes in specific research questions. Various obstacles had to be resolved to ensure a consistent model that is capable of being used for a variety of different research topics. Once established, this model has proven to be highly adaptable to different questions and can easily be altered for different fields of study.

Following 10 min of stabilization and baseline readings, we randomized a first set of 10 male Sprague Dawley rats into five small groups: global no-flow ischemia for 5, 7.5, 8, 9, or 10 min (n = 2 per group) followed by reperfusion; these limited preliminary dose-finding experiments were conducted to identify the longest possible ischemia time to still allow sufficient ventilation and reperfusion before the eventual development of a precipitous and irreversible increase in airway pressure and edema formation. Importantly...

Watch this Scientific Journal Video about Design and Implementation of a Rat Ex Vivo Lung Perfusion Model at JoVE.com

Design and Implementation of a Rat Ex Vivo Lung Perfusion Model

Ex vivo lungs are useful for a variety of experiments to collect physiological data while excluding the confounding variables of in vivo experiments. Commercial setups are often expensive and limited in the types of data they can collect. We describe a method for building a fully modular setup, adaptable for various study designs.

Design and Implementation of a Rat Ex Vivo Lung Perfusion Model

Ex vivo lung preparations are a useful model that can be translated to many different fields of research, complementing corresponding in vivo and in vitro ...

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699 Views.  Vanderbilt University Medical Center. Ex vivo lungs are useful for a variety of experiments to collect physiological data while excluding the confounding variables of in vivo experiments. Commercial setups are often expensive and limited in the types of data they can collect. We describe a method for building a fully modular setup, adaptable for various study designs. 

Video: Design and Implementation of a Rat Ex Vivo Lung Perfusion Model

Method for Culture of Early Chick Embryos ex vivo (New Culture)

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying  the formation and patterning of brain, neural tube, somite and heart primordia. Applications of chick embryo culture include electroporation of DNA or RNA constructs in order to analyze gene function, grafts of growth factor coated beads such as FGFs and BMPs , as well as whole mount in situ hybridization and immunohistochemistry. This video demonstrates the different steps in chick embryo culture; First, the embryo is explanted in saline. Then, the embryo is centered on a glass ring. The membranes surrounding the embryo are lifted along the walls of the ring. The ring is then placed in a culture dish containing a pool of albumine. The culture dish is sealed and placed in a humid chamber, where the embryo is cultured for up to 24 hrs. Finally, the embryo is removed from the ring, fixed and processed for further applications. A troubleshooting guide is also presented.

Method for Culture of Early Chick Embryos ex vivo New Culture

This video demonstrates New culture, a method by which chick embryos are cultured outside the egg for up to 24 hr. This method enables one to study early development (primitive streak to 14 som.), a period corresponding to E7-9 in mouse. Applications of this technique include electroporation, in situ hybridization and immunohistochemistry.

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ...

Isolation of Rat Portal Fibroblasts by In situ Liver Perfusion

Liver fibrosis is defined by the excessive deposition of extracellular matrix by activated myofibroblasts. There are multiple precursors of hepatic myofibroblasts, including hepatic stellate cells, portal fibroblasts and bone marrow derived fibroblasts 1. Hepatic stellate cells have been the best studied, but portal fibroblasts are increasingly recognized as important contributors to the myofibroblast pool, particularly in biliary fibrosis 2. Portal fibroblasts undergo proliferation in response to biliary epithelial injury, potentially playing a key role in the early stages of biliary scarring 3-5. A method of isolating portal fibroblasts would allow in vitro study of this cell population and lead to greater understanding of the role portal fibroblasts play in biliary fibrosis.
Portal fibroblasts have been isolated using various techniques including outgrowth 6, 7 and liver perfusion with enzymatic digestion followed by size selection 8. The advantage of the digestion and size selection technique compared to the outgrowth technique is that cells can be studied without the necessity of passage in culture. Here, we describe a modified version of the original technique described by Kruglov and Dranoff 8 for isolation of portal fibroblasts from rat liver that results in a relatively pure population of primary cells.

Isolation of Rat Portal Fibroblasts by In situ Liver Perfusion

A technique for isolating portal fibroblasts from rat liver is described. Livers are perfused and digested in situ with collagenase, followed by ex vivo digestion of the liver slurry and size selection of cells. This method provides a pure population of portal fibroblasts without the need for passage in culture.

Liver fibrosis is defined  by the excessive deposition of extracellular matrix by activated  myofibroblasts. There are multiple precursors of hepatic ...

A System for ex vivo Culturing of Embryonic Pancreas

The pancreas controls vital functions of our body, including the production of digestive enzymes and regulation of blood sugar levels1. Although in the past decade many studies have contributed to a solid foundation for understanding pancreatic organogenesis, important gaps persist in our knowledge of early pancreas formation2. A complete understanding of these early events will provide insight into the development of this organ, but also into incurable diseases that target the pancreas, such as diabetes or pancreatic cancer. Finally, this information will generate a blueprint for developing cell-replacement therapies in the context of diabetes.
 During embryogenesis, the pancreas originates from distinct embryonic outgrowths of the dorsal and ventral foregut endoderm at embryonic day (E) 9.5 in the mouse embryo3,4. Both outgrowths evaginate into the surrounding mesenchyme as solid epithelial buds, which undergo proliferation, branching and differentiation to generate a fully mature organ2,5,6. Recent evidences have suggested that growth and differentiation of pancreatic cell lineages, including the insulin-producing &beta;-cells, depends on proper tissue-architecture, epithelial remodeling and cell positioning within the branching pancreatic epithelium7,8. However, how branching morphogenesis occurs and is coordinated with proliferation and differentiation in the pancreas is largely unknown. This is in part due to the fact that current knowledge about these developmental processes has relied almost exclusively on analysis of fixed specimens, while morphogenetic events are highly dynamic.
 Here, we report a method for dissecting and culturing mouse embryonic pancreatic buds ex vivo on glass bottom dishes, which allow direct visualization of the developing pancreas (Figure 1). This culture system is ideally devised for confocal laser scanning microscopy and, in particular, live-cell imaging. Pancreatic explants can be prepared not only from wild-type mouse embryos, but also from genetically engineered mouse strains (e.g. transgenic or knockout), allowing real-time studies of mutant phenotypes. Moreover, this ex vivo culture system is valuable to study the effects of chemical compounds on pancreatic development, enabling to obtain quantitative data about proliferation and growth, elongation, branching, tubulogenesis and differentiation. In conclusion, the development of an ex vivo pancreatic explant culture method combined with high-resolution imaging provides a strong platform for observing morphogenetic and differentiation events as they occur within the developing mouse embryo.

A System for ex vivo Culturing of Embryonic Pancreas

Here, we describe a method for isolation, culture and manipulation of mouse embryonic pancreas. This represents an excellent ex vivo system for studying various aspects of pancreatic development, including morphogenesis, differentiation and growth. Pancreatic bud explants can be cultured for several days and used in a range of different applications, including whole-mount immunofluorescence and live imaging.

The pancreas controls vital functions of our body,  including the production of digestive enzymes and regulation of blood sugar  levels1. Although in the ...

A Novel Ex vivo Culture Method for the Embryonic Mouse Heart

Developmental studies in the mouse are hampered by the inaccessibility of the embryo during gestation. Thus, protocols to isolate and culture individual organs of interest are essential to provide a method of both visualizing changes in development and allowing novel treatment strategies. To promote the long-term culture of the embryonic heart at late stages of gestation, we developed a protocol in which the excised heart is cultured in a semi-solid, dilute Matrigel. This substrate provides enough support to maintain the three-dimensional structure but is flexible enough to allow continued contraction. In brief, hearts are excised from the embryo and placed in a mixture of cold Matrigel diluted 1:1 with growth medium. After the diluted Matrigel solidifies, growth medium is added to the culture dish. Hearts excised as late as embryonic day 16.5 were viable for four days post-dissection. Analysis of the coronary plexus shows that this method does not disrupt coronary vascular development. Thus, we present a novel method for long-term culture of embryonic hearts.

A Novel Ex vivo Culture Method for the Embryonic Mouse Heart

Developmental studies in the mouse are hampered by the inaccessibility of the embryo during gestation. To promote the long-term culture of the embryonic heart at late stages of gestation, we developed a protocol in which the excised heart is cultured in a semi-solid, dilute Matrigel.

Developmental studies in the mouse are hampered  by the inaccessibility of the embryo during gestation. Thus, protocols to  isolate and culture individual ...

The c-FOS Protein Immunohistological Detection: A Useful Tool As a Marker of Central Pathways Involved in Specific Physiological Responses In Vivo and Ex Vivo

Many studies seek to identify and map the brain regions involved in specific physiological regulations. The proto-oncogene c-fos, an immediate early gene, is expressed in neurons in response to various stimuli. The protein product can be readily detected with immunohistochemical techniques leading to the use of c-FOS detection to map groups of neurons that display changes in their activity. In this article, we focused on the identification of brainstem neuronal populations involved in the ventilatory adaptation to hypoxia or hypercapnia. Two approaches were described to identify involved neuronal populations in vivo in animals and ex vivo in deafferented brainstem preparations. In vivo, animals were exposed to hypercapnic or hypoxic gas mixtures. Ex vivo, deafferented preparations were superfused with hypoxic or hypercapnic artificial cerebrospinal fluid. In both cases, either control in vivo animals or ex vivo preparations were maintained under normoxic and normocapnic conditions. The comparison of these two approaches allows the determination of the origin of the neuronal activation i.e., peripheral and/or central. In vivo and ex vivo, brainstems were collected, fixed, and sliced into sections. Once sections were prepared, immunohistochemical detection of the c-FOS protein was made in order to identify the brainstem groups of cells activated by hypoxic or hypercapnic stimulations. Labeled cells were counted in brainstem respiratory structures. In comparison to the control condition, hypoxia or hypercapnia increased the number of c-FOS labeled cells in several specific brainstem sites that are thus constitutive of the neuronal pathways involved in the adaptation of the central respiratory drive.

The c-FOS Protein Immunohistological Detection: A Useful Tool As a Marker of Central Pathways Involved in Specific Physiological Responses In Vivo and Ex Vivo

Here, we present a protocol based on c-FOS protein immunohistological detection, a classical technique used for the identification of neuronal populations involved in specific physiological responses in vivo and ex vivo.

Many studies seek to identify and map the brain regions involved in specific physiological regulations. The proto-oncogene c-fos, an immediate early gene, ...

Isolation of CD146+ Resident Lung Mesenchymal Stromal Cells from Rat Lungs

Mesenchymal stromal cells (MSCs) are increasingly recognized for their therapeutic potential in a wide range of diseases, including lung diseases. Besides the use of bone marrow and umbilical cord MSCs for exogenous cell therapy, there is also increasing interest in the repair and regenerative potential of resident tissue MSCs. Moreover, they likely have a role in normal organ development, and have been attributed roles in disease, particularly those with a fibrotic nature. The main hurdle for the study of these resident tissue MSCs is the lack of a clear marker for the isolation and identification of these cells. The isolation technique described here applies multiple characteristics of lung resident MSCs (L-MSCs). Upon sacrifice of the rats, lungs are removed and rinsed multiple times to remove blood. Following mechanical dissociation by scalpel, the lungs are digested for 2-3 hr using a mix of collagenase type I, neutral protease and DNase type I. The obtained single cell suspension is subsequently washed and layered over density gradient medium (density 1.073 g/ml). After centrifugation, cells from the interphase are washed and plated in culture-treated flasks. Cells are cultured for 4-7 days in physiological 5% O2, 5% CO2 conditions. To deplete fibroblasts (CD146-) and to ensure a population of only L-MSCs (CD146+), positive selection for CD146+ cells is performed through magnetic bead selection. In summary, this procedure reliably produces a population of primary L-MSCs for further in vitro study and manipulation. Because of the nature of the protocol, it can easily be translated to other experimental animal models.

Isolation of CD146+ Resident Lung Mesenchymal Stromal Cells from Rat Lungs

This protocol describes an isolation technique for obtaining primary lung resident mesenchymal stromal cells from rats, through the use of enzymatic digestion, density gradient separation, plastic adherence and CD146+ magnetic bead selection.

Mesenchymal stromal cells (MSCs) are increasingly recognized for their therapeutic potential in a wide range of diseases, including lung diseases. Besides ...

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a versatile analytical tool for characterizing viscoelastic mechanical properties for a variety of materials ranging from hard (plastic, glass, metal, etc.) surfaces to cells on any substrate. It has been widely used to measure the stiffness of cells, but less frequently used to measure the stiffness of aortas. In this paper, we will describe the procedures for using AFM in contact mode to measure the ex vivo elastic modulus of unloaded mouse arteries. We describe our procedure for isolation of mouse aortas, and then provide detailed information for the AFM analysis. This includes step-by-step instructions for alignment of the laser beam, calibration of the spring constant and deflection sensitivity of the AFM probe, and acquisition of force curves. We also provide a detailed protocol for data analysis of the force curves.

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

We present detailed protocols for isolation of aortas from mouse and measurement of their elastic modulus using atomic force microscopy.

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a ...

An Ex Vivo Tissue Culture Model of Cartilage Remodeling in Bovine Knee Explants

Ex vivo culture systems cover a broad range of experiments dedicated to studying tissue and cellular function in a native setting. Cartilage is a unique tissue important for proper function of the synovial joint and is constituted by a dense extracellular matrix (ECM), rich in proteoglycan and type II collagen. Chondrocytes are the only cell type present within cartilage and are widespread and relatively low in number. Altered external stimuli and cellular signalling can lead to changes in ECM composition and deterioration, which are important pathological hallmarks in diseases such as osteoarthritis (OA) and rheumatoid arthritis.
Ex vivo cartilage models allow 1) profiling of chondrocyte mediated alterations of cartilage tissue turnover, 2) visualizing the cartilage ECM composition, and 3) chondrocyte rearrangement directly in the tissue. Profiling these alterations in response to stimuli or treatments are of high importance in various aspects of cartilage biology, and complement in vitro experiments in isolated chondrocytes, or more complex models in live animals where experimental conditions are more difficult to control.
Cartilage explants present a translational and easily accessible method for assessing tissue remodeling in the cartilage ECM in controllable settings. Here, we describe a protocol for isolating and culturing live bovine cartilage explants. The method uses tissue from the bovine knee, which is easily accessible from the local butchery. Both explants and conditioned culture medium can be analyzed to investigate tissue turnover, ECM composition, and chondrocyte function, thus profiling ECM modulation.

Here, we present a protocol describing isolation and culturing of cartilage explants from bovine knees. This method provides an easy and accessible tool to describe tissue changes in response to biological stimuli or novel therapeutics targeting the joint.

Ex vivo culture systems cover a broad range of experiments dedicated to studying tissue and cellular function in a native setting. Cartilage is a unique ...

Mass Spectrometry Analysis to Identify Ubiquitylation of EYFP-tagged CENP-A (EYFP-CENP-A)

Studying the structure and the dynamics of kinetochores and centromeres is important in understanding chromosomal instability (CIN) and cancer progression. How the chromosomal location and function of a centromere (i.e., centromere identity) are determined and participate in accurate chromosome segregation is a fundamental question. CENP-A is proposed to be the non-DNA indicator (epigenetic mark) of centromere identity, and CENP-A ubiquitylation is required for CENP-A deposition at the centromere, inherited through dimerization between cell division, and indispensable to cell viability.
Here we describe mass spectrometry analysis to identify ubiquitylation of EYFP-CENP-A K124R mutant suggesting that ubiquitylation at a different lysine is induced because of the EYFP tagging in the CENP-A K124R mutant protein. Lysine 306 (K306) ubiquitylation in EYFP-CENP-A K124R was successfully identified, which corresponds to lysine 56 (K56) in CENP-A through mass spectrometry analysis. A caveat is discussed in the use of GFP/EYFP or the tagging of high molecular weight protein as a tool to analyze the function of a protein. Current technical limit is also discussed for the detection of ubiquitylated bands, identification of site-specific ubiquitylation(s), and visualization of ubiquitylation in living cells or a specific single cell during the whole cell cycle.
The method of mass spectrometry analysis presented here can be applied to human CENP-A protein with different tags and other centromere-kinetochore proteins. These combinatory methods consisting of several assays/analyses could be recommended for researchers who are interested in identifying functional roles of ubiquitylation.

Mass Spectrometry Analysis to Identify Ubiquitylation of EYFP-tagged CENP-A EYFP-CENP-A

CENP-A ubiquitylation is an important requirement for CENP-A deposition at the centromere, inherited through dimerization between cell division, and indispensable to cell viability. Here we describe mass spectrometry analysis to identify ubiquitylation of EYFP-tagged CENP-A (EYFP-CENP-A) protein.

Studying the structure and the dynamics of kinetochores and centromeres is important in understanding chromosomal instability (CIN) and cancer ...

Isolation of Primary Rat Hepatocytes with Multiparameter Perfusion Control

Primary hepatocytes are widely used in basic research on liver diseases and for toxicity testing in vitro. The two-step collagenase perfusion procedure for primary hepatocyte isolation is technically challenging, especially in portal vein cannulation. The procedure is also prone to occasional contamination and variations in perfusion conditions due to difficulties in the assembly, optimization, or maintenance of the perfusion setup. Here, a detailed protocol for an improved two-step collagenase perfusion procedure with multiparameter perfusion control is presented. Primary rat hepatocytes were successfully and reliably isolated by taking the necessary technical precautions at critical steps of the procedure, and by reducing the operational difficulty and mitigating the variability of perfusion parameters through the adoption of a special intravenous catheter, standardized sterile disposable tubing, temperature control, and real-time monitoring and alarm system. The isolated primary rat hepatocytes consistently exhibit high cell viability (85%-95%), yield (2-5 x 108 cells per 200-300 g rat) and functionality (albumin, urea and CYP activity). The procedure was complemented by an integrated perfusion system, which is compact enough to be set up in the laminar flow hood to ensure aseptic operation.

This protocol details the use of a special intravenous catheter, standardized sterile disposable tubing, temperature control complemented by real-time monitoring, and an alarm system for two-step collagenase perfusion procedure to improve the consistency in the viability, yield, and functionality of isolated primary rat hepatocytes.

Primary hepatocytes are widely used in basic research on liver diseases and for toxicity testing in vitro. The two-step collagenase perfusion procedure ...

Design and Implementation of a Rat Ex Vivo Lung Perfusion Model

Summary

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Chapters in this video

Method for Culture of Early Chick Embryos ex vivo (New Culture)

Isolation of Rat Portal Fibroblasts by In situ Liver Perfusion

A System for ex vivo Culturing of Embryonic Pancreas

A Novel Ex vivo Culture Method for the Embryonic Mouse Heart

The c-FOS Protein Immunohistological Detection: A Useful Tool As a Marker of Central Pathways Involved in Specific Physiological Responses In Vivo and Ex Vivo

Isolation of CD146⁺ Resident Lung Mesenchymal Stromal Cells from Rat Lungs

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

An Ex Vivo Tissue Culture Model of Cartilage Remodeling in Bovine Knee Explants

Mass Spectrometry Analysis to Identify Ubiquitylation of EYFP-tagged CENP-A (EYFP-CENP-A)

Isolation of Primary Rat Hepatocytes with Multiparameter Perfusion Control