This project was supported by KFO 311 Grant from Deutsche Forschungsgemeinschaft.

Experiments were performed on male C57BL/6 mice, aged 12 weeks. This study was conducted in compliance with guidelines of the German Animal Law under Protocol TSA 16/2250.
1. Materials Preparation
NOTE: All steps are performed under clean, non-sterile conditions.&#160;Sterile conditions would be required if animal is to be survived postoperatively.
<ol>
	<li>Introduce 3 fenestrations into a 2-Fr polyurethane tube using a surgical blade under a microscope with 16X magnification. 
	NOTE: All fenestrations must be located in the distal third of the cannula to ensure optimal blood drainage.</li>
	<li>Prepare the priming solution (Materials Table). Include 30 IU/mL heparin and 2.5% v/v of an 8.4% solution of NaHCO3. Refrigerate this solution at 4 &#176;C until it is ready to use. Prime the circuit with 500 uL of priming solution.</li>
	<li>Place the outflow cannula into the priming solution and fill the ECMO machine by switching on the peristaltic pump. Continue to circulate the priming solution through the machine for the next 30 min at a flow rate of 1 mL/min.</li>
	<li>Give 0.5 L/min of 100% oxygen to the oxygenator.</li>
</ol>
2. Anesthesia
<ol>
	<li>Place the animal in an induction chamber filled with a 2.5% v/v isoflurane/oxygen mixture. Provide 0.5 L/min of 100% oxygen to the vaporizer. Before surgery, check that full anesthesia is achieved by testing pedal withdrawal and pain reflexes. Apply eye gel to prevent drying damage.</li>
	<li>Use a warming pad to maintain the body temperature at 37 &#176;C.</li>
	<li>Perform inhalation mask anesthesia using an isoflurane vaporizer and inject 5 mg/kg carprofen subcutaneously.</li>
	<li>Regularly observe spontaneous breathing and adjust the concentration of isoflurane so that it is between 1.3 and 2.5%.</li>
</ol>
3. Surgery
<ol>
	<li>Expose the left jugular vein by using a lateral skin incision of 4 mm with the help of fine scissors on the left side of the neck. Together with sharp and blunt preparation using micro-forceps and cotton swabs, use bipolar coagulation of the small vessels.</li>
	<li>Once the left jugular vein is exposed, ligate the distal part using an 8-0 silk suture with the help of micro-forceps.</li>
	<li>Place a slip knot at the proximal end of the vein. Incise the anterior wall of the vein using micro-scissors.</li>
	<li>To achieve full heparinization, inject 2.5 IU/g heparin into the jugular vein via a 26 G braunula.</li>
	<li>Raise the head side of the animal pad by 30&#176; to avoid excessive blood loss from the vein during insertion of the cannula.</li>
	<li>Insert a 2-Fr polyurethane (PU) cannula into the proximal part of the jugular vein, rotating it slightly while pushing it to a depth of 4 cm; while doing so, the iliac bifurcation of inferior vena cava (IVC) will be reached.</li>
	<li>Secure the cannula with 8-0 silk knots using microforceps.</li>
	<li>Expose the right jugular vein using the steps described in 3.1, 3.2, and 3.3.</li>
	<li>Cannulate the right jugular vein with a 1-Fr PU cannula and gently move it 5 mm towards the direction of right atrium.</li>
	<li>Repeat step 3.7.</li>
	<li>Catheterize the left femoral artery with another 1-Fr PU cannula and use it for invasive pressure monitoring as well as blood sampling for blood gas analysis (BGA).</li>
	<li>Insert electrocardiogram (ECG) needles connected to a data acquisition device subcutaneously into both forelimbs and into the left thoracic wall.</li>
	<li>Insert a rectal thermometer connected to a data acquisition device.</li>
</ol>
4. Veno-Venous Extracorporeal Membrane Oxygenation and Blood Gas Analysis
NOTE: For a schematic of the complete ECMO circuit, see Figure 1.
<ol>
	<li>Initiate ECMO on the animal by turning on the pump with an initial flow rate of 0.1 mL/min. Adjust the flow rate of the pump within the next 2 min to 3-5 mL/min.</li>
	<li>In case of air suction in the outflow cannula via the cannulation site, reduce the flow and add 0.1 mL of priming solution to the circuit via an air-trapping reservoir.</li>
	<li>Under stable flow, continue to monitor in real-time mode all vital parameters via the data acquisition device.</li>
	<li>Constantly observe backflow from the venous drainage and monitor the level of the blood in the air-trapper reservoir.</li>
	<li>Collect any blood leaking from wounds into a 1 cc syringe with the tip of a 24 G branula andreturn it to the ECMO circuit via the air-trapping reservoir.</li>
	<li>For BGA, use a blood sampling cartridge to collect approximately 75 &#181;L of arterial blood at the following time points and from the following locations:
	<ol>
		<li>10 min after the initiation of ECMO, collect blood from the IVC via an extra tube built in before the oxygenator, via similar extra tube&#160;after oxygenator (control), and directly from the femoral artery.</li>
		<li>30 min after the initiation of ECMO, collect blood from the femoral artery.</li>
	</ol>
	</li>
	<li>Give an extra 0.1 mL of priming solution to compensate for intravasal liquid loss every 45 min via the air-trapper or femoral artery catheter or by sucking the air bubbles through the blood draining cannula.</li>
	<li>For BGA, use a blood sampling cartridge to collect approximately 75 &#181;L of arterial blood:
	<ol>
		<li>1 h after the initiation of ECMO from the femoral artery.</li>
		<li>2 h after the initiation of ECMO, collect blood from the IVC via an extra tube built in before the oxygenator, via similar extra tube after oxygenator (control),&#160;and directly from the femoral artery.</li>
	</ol>
	</li>
	<li>After 2 h, reduce the flow rate on the pump gradually (over the course of 5 min), thereby stopping ECMO.</li>
	<li>Continue to record vital parameters for another 10 min.</li>
	<li>Finish the experiment by exsanguinating the animal and harvesting the blood and organs.</li>
</ol>

<ol>
	<li>Hill, J. D., 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=Prolonged+Extracorporeal+Oxygenation+for+Acute+Post-Traumatic+Respiratory+Failure+(Shock-Lung+Syndrome).">Prolonged Extracorporeal Oxygenation for Acute Post-Traumatic Respiratory Failure (Shock-Lung Syndrome).</a> New England Journal of Medicine. 286 (12), 629-634 (1972).</li><li>Noah, M. A., 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=Referral+to+an+Extracorporeal+Membrane+Oxygenation+Center+and+Mortality+Among+Patients+With+Severe+2009+Influenza+A(H1N1).">Referral to an Extracorporeal Membrane Oxygenation Center and Mortality Among Patients With Severe 2009 Influenza A(H1N1).</a> Journal of the American Medical Association. 306 (15), 1659 (2011).</li><li>Maslach-Hubbard, A., Bratton, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Extracorporeal+membrane+oxygenation+for+pediatric+respiratory+failure:+History,+development+and+current+status.">Extracorporeal membrane oxygenation for pediatric respiratory failure: History, development and current status.</a> World Journal of Critical. Care Medicine. 2 (4), 29-39 (2013).</li><li>Langer, T., 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="Awake"+extracorporeal+membrane+oxygenation+(ECMO):+pathophysiology,+technical+considerations,+and+clinical+pioneering.">"Awake" extracorporeal membrane oxygenation (ECMO): pathophysiology, technical considerations, and clinical pioneering.</a> Critical Care. 20 (1), 150 (2016).</li><li>Esper, S. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Extracorporeal+Membrane+Oxygenation.">Extracorporeal Membrane Oxygenation.</a> Advances in Anesthesia. 35 (1), 119-143 (2017).</li><li>Millar, J. E., Fanning, J. P., McDonald, C. I., McAuley, D. F., Fraser, J. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+inflammatory+response+to+extracorporeal+membrane+oxygenation+(ECMO):+a+review+of+the+pathophysiology.">The inflammatory response to extracorporeal membrane oxygenation (ECMO): a review of the pathophysiology.</a> Critical Care. 20 (1), 387 (2016).</li><li>Lubnow, M., 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=Technical+complications+during+veno-venous+extracorporeal+membrane+oxygenation+and+their+relevance+predicting+a+system-exchange--retrospective+analysis+of+265+cases.">Technical complications during veno-venous extracorporeal membrane oxygenation and their relevance predicting a system-exchange--retrospective analysis of 265 cases.</a> Public Library of Science One. 9 (12), e112316 (2014).</li><li>Passmore, M. R., 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=Inflammation+and+lung+injury+in+an+ovine+model+of+extracorporeal+membrane+oxygenation+support.">Inflammation and lung injury in an ovine model of extracorporeal membrane oxygenation support.</a> American Journal of Physiology - Lung Cellular and Molecular Physiology. 311 (6), L1202-L1212 (2016).</li><li>Vaquer, S., de Haro, C., Peruga, P., Oliva, J. C., Artigas, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Systematic+review+and+meta-analysis+of+complications+and+mortality+of+veno-venous+extracorporeal+membrane+oxygenation+for+refractory+acute+respiratory+distress+syndrome.">Systematic review and meta-analysis of complications and mortality of veno-venous extracorporeal membrane oxygenation for refractory acute respiratory distress syndrome.</a> Annals of Intensive Care. 7 (1), 51 (2017).</li><li>Houser, S. R., 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=Animal+Models+of+Heart+Failure+A+Scientific+Statement+From+the+American+Heart+Association.">Animal Models of Heart Failure A Scientific Statement From the American Heart Association.</a> Circulation Research. 111 (1), 131-150 (2012).</li><li>Russell, J. C., Proctor, S. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Small+animal+models+of+cardiovascular+disease:+tools+for+the+study+of+the+roles+of+metabolic+syndrome,+dyslipidemia,+and+atherosclerosis.">Small animal models of cardiovascular disease: tools for the study of the roles of metabolic syndrome, dyslipidemia, and atherosclerosis.</a> Cardiovascular Pathology. 15 (6), 318-330 (2006).</li><li>Madrahimov, N., 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=Novel+mouse+model+of+cardiopulmonary+bypass.">Novel mouse model of cardiopulmonary bypass.</a> European Journal of Cardio-thoracic Surgery. 53 (1), 186-193 (2017).</li><li>Madrahimov, N., 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=Cardiopulmonary+Bypass+in+a+Mouse+Model:+A+Novel+Approach.">Cardiopulmonary Bypass in a Mouse Model: A Novel Approach.</a> J. Journal of Visualized Experiments. (127), (2017).</li></ol>

The authors have nothing to disclose.

Previously, we described a successful model of CPB in a mouse12,13. To implement such a model for acute or end-stage lung disorders we developed an easy-to-use veno-venous ECMO circuit for mice. Different to the CPB model, veno-venous ECMO does not require complicated surgical procedures such as sternotomy and clamping of the aorta, thus reducing the risk of wound bleeding in a fully heparinized animal. To avoid embolization of the oxygenator with blood clots, 2....

Extracorporeal membrane oxygenation (ECMO) is a temporary life support system that takes over functions of the lungs and heart to allow adequate gas exchange and perfusion. Hill et al1 described the first use of ECMO in patients in 1972; however, it only became widely used after its successful application during the H1N1 influenza pandemic in 20092. Today, ECMO is routinely used as a lifesaving procedure in end-stage heart and lung diseases3. Veno-venous ECMO is increasingly employed as an alternative to invasive mechanical ventilation in awake, non-intubated, spontaneously breathing patients with refractory respiratory failure4.
Despite its widespread adoption, diverse complications have been reported for ECMO5,6,7. Complications that can be experienced by patients on ECMO include bleeding, thrombosis, sepsis, thrombocytopenia, device-related malfunctions, and air embolism. Moreover, a systemic inflammatory response syndrome (SIRS) resulting in multi-organ damage is well-described both clinically and in experimental studies8,9. Neurological complications such as brain infarction are also frequently reported in patients undergoing long-term ECMO therapy. To confuse matters, it is often difficult to distinguish whether complications are caused by ECMO itself or arise from the underlying disorders accompanying acute and end-stage diseases.
To specifically study the effects of ECMO on a healthy organism, a reliable experimental animal model must be established. There are very few reports on performance of ECMO on small animals and are all limited to rats. To date, no mouse model of ECMO has been described in the literature. Due to the availability of a large number of genetically modified mouse strains, establishment of a mouse ECMO model would allow further investigation of the molecular mechanisms involved in ECMO-related complications10,11.
Based on our previously described murine model of cardiopulmonary bypass (CPB)12, we have developed a stable method of veno-venous ECMO in non-intubated, spontaneously breathing mice. The ECMO circuit (Figure 1), containing outflow and inflow cannulas, a peristaltic pump, oxygenator, and air-trapping reservoir, is similar to our previously described model of murine CPB12 with the exception of having a smaller priming volume (0.5 mL). This protocol demonstrates the detailed techniques, physiological monitoring, and blood gas analysis involved in a successful ECMO procedure.

<table><tbody><tr><td>Sterofundin</td><td>B.Braun Petzold GmbH</td><td>PZN:8609189</td><td>in 1:1 with Tetraspan</td></tr><tr><td>Tetraspan 6% Solution</td><td>B. Braun Melsungen AG</td><td>PZN: 05565416</td><td>in 1:1 with Sterofundin</td></tr><tr><td>Heparin Natrium 25.000</td><td>Ratiopharm GmbH</td><td>PZN: 3029843</td><td>2,5 IU per ml of priming</td></tr><tr><td>NaHCO3 8,4% Solution</td><td>B. Braun Melsungen AG</td><td>PZN: 1579775</td><td>3% in priming solution</td></tr><tr><td>Carprofen</td><td>Zoetis Inc., USA</td><td>PZN:00289615</td><td>5mg/kg/BW</td></tr><tr><td>1 Fr PU Catheter</td><td>Instechlabs INC., USA</td><td>C10PU-MCA1301</td><td>carotide artery</td></tr><tr><td>2 Fr PU Catheter</td><td>Instechlabs INC., USA</td><td>C20PU-MJV1302</td><td>jugular vein</td></tr><tr><td>8-0 Silk suture braided</td><td>Ashaway Line &amp;amp; Twine Co., USA</td><td>75290</td><td>ligature</td></tr><tr><td>Isoflurane</td><td>Piramal Critical Care GmbH</td><td>PZN:9714675</td><td>narcosis</td></tr><tr><td>Spring Scissors - 6mm Blades</td><td>Fine Science Tools GmbH</td><td>15020-15</td><td>instruments</td></tr><tr><td>Spring Scissors - 2mm Blades</td><td>Fine Science Tools GmbH</td><td>15000-03</td><td>instruments</td></tr><tr><td>Halsted-Mosquito Hemostat</td><td>Fine Science Tools GmbH</td><td>13009-12</td><td>instruments</td></tr><tr><td>Dumont #55 Forceps</td><td>Fine Science Tools GmbH</td><td>11295-51</td><td>instruments</td></tr><tr><td>Castroviejo Micro Needle Holder - 9cm</td><td>Fine Science Tools GmbH</td><td>12060-02</td><td>instruments</td></tr><tr><td>Micro Serrefines</td><td>Fine Science Tools GmbH</td><td>18555-01</td><td>instruments</td></tr><tr><td>Bulldog Serrefine</td><td>Fine Science Tools GmbH</td><td>18050-28</td><td>instruments</td></tr><tr><td>Isoflurane Vaporizer Drager 19.1</td><td>Dr&amp;auml;gerwerk AG &amp;amp; Co. KGaA</td><td></td><td>anesthesia 1,3 -2,5%</td></tr><tr><td>Multichannel Data Aquisition Device with ISOHEART Software</td><td>Hugo Sachs Elektronik GmbH, Germany</td><td></td><td>invasive pressure, ECG, t &amp;deg;C</td></tr><tr><td>i-STAT portable device</td><td>Abbott Laboratories, Lake Bluff, Illinois, USA</td><td></td><td>blood gas analysis</td></tr><tr><td>i-STAT CG4+ and CG8+ cartridges</td><td>Abbott Laboratories, Lake Bluff, Illinois, USA</td><td></td><td>blood gas analysis</td></tr><tr><td>C57Bl/6 mice, male, 30 g, 14 weeks old</td><td>Charles River Laboratories</td><td></td><td>housed 1 week before</td></tr></tbody></table>

veno-venous extracorporeal membrane oxygenation in a mouse

The use of extracorporeal membrane oxygenation (ECMO) has increased substantially in recent years. ECMO has become a reliable and effective therapy for acute as well as end-stage lung diseases. With the increase in clinical demand and prolonged use of ECMO, procedural optimization and prevention of multi-organ damage are of critical importance. The aim of this protocol is to present a detailed technique of veno-venous ECMO in a non-intubated, spontaneously breathing mouse. This protocol demonstrates the technical design of the ECMO and surgical steps. This murine ECMO model will facilitate the study of pathophysiology related to ECMO (e.g., inflammation,bleeding and thromboembolic events). Due to the abundance of genetically modified mice, the molecular mechanisms involved in ECMO-related complications can also be dissected.

This protocol describes the method of veno-venous ECMO in a mouse. This model is reliable and reproducible, and compared to our previously described model of CPB with respiratory and circulatory arrest12,13, it is less technically demanding to establish.
ECMO flow in the venous system was maintained between 1.5 and 5 mL/min. The mean arterial pressure was kept between 70...

Watch this Scientific Journal Video about Veno-Venous Extracorporeal Membrane Oxygenation in a Mouse at JoVE.com

Veno-Venous Extracorporeal Membrane Oxygenation in a Mouse

Here we present a protocol describing the technique of veno-venous extracorporeal membrane oxygenation (ECMO) in a non-intubated, spontaneously breathing mouse. This murine model of ECMO can be effectively implemented in experimental studies of acute and end-stage lung diseases.

The use of extracorporeal membrane oxygenation (ECMO) has increased substantially in recent years. ECMO has become a reliable and effective therapy for ...

veno-venous-extracorporeal-membrane-oxygenation-in-a-mouse

Research

JoVE Journal

Medicine

12.4K Views.  Hannover Medical School. Here we present a protocol describing the technique of veno-venous extracorporeal membrane oxygenation (ECMO) in a non-intubated, spontaneously breathing mouse. This murine model of ECMO can be effectively implemented in experimental studies of acute and end-stage lung diseases.

Video: Veno-Venous Extracorporeal Membrane Oxygenation in a Mouse

In vivo Measurement of the Mouse Pulmonary Endothelial Surface Layer

The endothelial glycocalyx is a layer of proteoglycans and associated glycosaminoglycans lining the vascular lumen. In vivo, the glycocalyx is highly hydrated, forming a substantial endothelial surface layer (ESL) that contributes to the maintenance of endothelial function. As the endothelial glycocalyx is often aberrant in vitro and is lost during standard tissue fixation techniques, study of the ESL requires use of intravital microscopy. To best approximate the complex physiology of the alveolar microvasculature, pulmonary intravital imaging is ideally performed on a freely-moving lung. These preparations, however, typically suffer from extensive motion artifact. We demonstrate how closed-chest intravital microscopy of a freely-moving mouse lung can be used to measure glycocalyx integrity via ESL exclusion of fluorescently-labeled high molecular weight dextrans from the endothelial surface. This non-recovery surgical technique, which requires simultaneous brightfield and fluorescent imaging of the mouse lung, allows for longitudinal observation of the subpleural microvasculature without evidence of inducing confounding lung injury.

In vivo Measurement of the Mouse Pulmonary Endothelial Surface Layer

The endothelial glycocalyx/endothelial surface layer is ideally studied using intravital microscopy. Intravital microscopy is technically challenging in a moving organ such as the lung. We demonstrate how simultaneous brightfield and fluorescent microscopy may be used to estimate endothelial surface layer thickness in a freely-moving in vivo mouse lung.

The endothelial glycocalyx is a layer of proteoglycans and associated glycosaminoglycans lining the vascular lumen. In vivo, the glycocalyx is highly ...

Ex Vivo Hepatic Perfusion Through the Portal Vein in Mouse

Metabolic diseases such as diabetes, pre-diabetes, non-alcoholic fatty liver disease (NAFLD), and nonalcoholic steatohepatitis (NASH) are becoming increasingly common. Ex vivo liver perfusions allow for a comprehensive analysis of liver metabolism using nuclear magnetic resonance (NMR), in nutritional conditions that can be rigorously controlled. As in silico simulations remain a primarily theoretical means of assessing hormone actions and the effects of pharmaceutical intervention, the perfused liver remains one of the most valuable test beds for understanding hepatic metabolism. As these studies guide basic insights into hepatic physiology, results must be accurate and reproducible. The greatest factor in the reproducibility of ex vivo hepatic perfusion is the quality of surgery. Therefore, we have introduced an organized and streamlined method to perform ex vivo mouse liver perfusions in the context of in situ NMR experiments. We also describe a unique application and discuss common issues encountered in these studies. The overall purpose is to provide an uncomplicated guide to a technique we have refined over several years that we deem the golden standard for obtaining reproducible results in hepatic resections and perfusions in the context of in situ NMR experiments. The distance to the center of the field for the magnet as well as the inaccessibility of the tissue to intervention during the NMR experiment makes our methods novel.

Ex Vivo Hepatic Perfusion Through the Portal Vein in Mouse

The protocol describes a straightforward method of resectioning an intact mouse liver for metabolic studies through portal vein perfusion.

Metabolic diseases such as diabetes, pre-diabetes, non-alcoholic fatty liver disease (NAFLD), and nonalcoholic steatohepatitis (NASH) are becoming ...

Biochemistry

Normothermic Ex Vivo Liver Machine Perfusion in Mouse

This protocol presents an optimized erythrocytes-free NEVLP system using mouse livers. Ex vivo preservation of mouse livers was achieved by employing modified cannulas and techniques adapted from conventional commercial ex vivo perfusion equipment. The system was utilized to evaluate the preservation outcomes following 12 h of perfusion. C57BL/6J mice served as liver donors, and the livers were explanted by cannulating the portal vein (PV) and bile duct (BD), and subsequently flushing the organ with warm (37 &#176;C) heparinized saline. Then, the explanted livers were transferred to the perfusion chamber and subjected to normothermic oxygenated machine perfusion (NEVLP). Inlet and outlet perfusate samples were collected at 3 h intervals for perfusate analysis. Upon completion of the perfusion, liver samples were obtained for histological analysis, with morphological integrity assessed using modified Suzuki-Score through Hematoxylin-Eosin (HE) staining. The optimization experiments yielded the following findings: (1) mice weighing over 30 g were deemed more suitable for the experiment due to the larger size of their bile duct (BD). (2) a 2 Fr (outer diameter = 0.66 mm) polyurethane cannula was better suited for cannulating the portal vein (PV) when compared to a polypropylene cannula. This was attributed to the polyurethane material&#39;s enhanced grip, resulting in reduced catheter slippage during the transfer from the body to the organ chamber. (3) for cannulation of the bile duct (BD), a 1 Fr (outer diameter = 0.33 mm) polyurethane cannula was found to be more effective compared to the polypropylene UT - 03 (outer diameter = 0.30 mm) cannula. With this optimized protocol, mouse livers were successfully preserved for a duration of 12 h without significant impact on the histological structure. Hematoxylin-Eosin (HE) staining revealed a well-preserved morphological architecture of the liver, characterized by predominantly viable hepatocytes with clearly visible nuclei and mild dilation of hepatic sinusoids.

Normothermic Ex Vivo Liver Machine Perfusion in Mouse

A normothermic ex vivo liver perfusion (NEVLP) system was created for mouse livers. This system requires experience in microsurgery but allows for reproducible perfusion results. The ability to utilize mouse livers facilitates the investigation of molecular pathways to identify novel perfusate additives and enables the execution of experiments focused on organ repair.

This protocol presents an optimized erythrocytes-free NEVLP system using mouse livers. Ex vivo preservation of mouse livers was achieved by employing ...

Engineering

Use of a Central Venous Line for Fluids, Drugs and Nutrient Administration in a Mouse Model of Critical Illness

This protocol describes a centrally catheterized mouse model of prolonged critical illness. We combine the cecal ligation and puncture method to induce sepsis with the use of a central venous line for fluids, drugs and nutrient administration to mimic the human clinical setting. Critically ill patients require intensive medical support in order to survive. While the majority of patients will recover within a few days, about a quarter of the patients need prolonged intensive care and are at high risk of dying from non-resolving multiple organ failure. Furthermore, the prolonged phase of critical illness is hallmarked by profound muscle weakness, and endocrine and metabolic changes, of which the pathogenesis is currently incompletely understood. The most widely used animal model in critical care research is the cecal ligation and puncture model to induce sepsis. This is a very reproducible model, with acute inflammatory and hemodynamic changes similar to human sepsis, which is designed to study the acute phase of critical illness. However, this model is hallmarked by a high lethality, which is different from the clinical human situation, and is not developed to study the prolonged phase of critical illness. Therefore, we adapted the technique by placing a central venous catheter in the jugular vein allowing us to administer clinically relevant supportive care, to better mimic the human clinical situation of critical illness. This mouse model requires an extensive surgical procedure and daily intensive care of the animals, but it results in a relevant model of the acute and prolonged phase of critical illness.

This protocol describes a centrally catheterized mouse model of prolonged critical illness. We combine the cecal ligation and puncture method to induce sepsis with the use of a central venous line for fluids, drugs and nutrient administration to mimic the human clinical setting.

This protocol describes a centrally catheterized mouse model of prolonged critical illness. We combine the cecal ligation and puncture method to induce ...

Cardiopulmonary Bypass in a Mouse Model: A Novel Approach

As prolonged cardiopulmonary bypass becomes more essential during cardiac interventions, an increasing clinical demand arises for procedure optimization and for minimizing organ damage resulting from prolonged extracorporal circulation. The goal of this paper was to demonstrate a fully functional and clinically relevant model of cardiopulmonary bypass in a mouse. We report on the device design, perfusion circuit optimization, and microsurgical techniques. This model is an acute model, which is not compatible with survival due to the need for multiple blood drawings. Because of the range of tools available for mice (e.g., markers, knockouts, etc.), this model will facilitate investigation into the molecular mechanisms of organ damage and the effect of cardiopulmonary bypass in relation to other comorbidities.

This paper describes how to perform cardiopulmonary bypass in mice. This novel model will facilitate the investigation of the molecular mechanisms involved in organ damage.

As prolonged cardiopulmonary bypass becomes more essential during cardiac interventions, an increasing clinical demand arises for procedure optimization ...

Effects of Extracorporeal Membrane Oxygenation on the Coagulation System

This study aimed to investigate the effects of long-term awake extracorporeal membrane oxygenation (ECMO) on the coagulation system in a sheep model. A total of ten healthy sheep were included in the study, with 5 sheep in each group. In the veno-arterial ECMO (V-A ECMO) group, cannulation was performed in the right carotid artery and the right external jugular vein. In the veno-venous ECMO (V-V ECMO) group, a dual-lumen catheter was utilized to insert into the right external jugular vein. After initiating ECMO, the sheep were recovered from anesthesia and remained awake for 7 days. The target activated clotting time (ACT) goal was set at 220-250 s. In both groups, the actual ACT fluctuated around 250 s with the dose of heparin gradually increasing, reaching almost 60 IU/kg/min at the end of the experiments. Moreover, the activated partial thromboplastin time (APTT) and thrombin time (TT) values were significantly higher in the V-A ECMO group compared to the V-V ECMO group, despite receiving the same doses of heparin. Although laboratory test results fluctuated within a normal and reasonable range, infarct foci in the kidneys were observed in both groups at the end of the study.

We present a protocol for establishing a long-term awake extracorporeal membrane oxygenation (ECMO) model in sheep. Special attention is given to the management and evaluation of the coagulation system during the ECMO model.

This study aimed to investigate the effects of long-term awake extracorporeal membrane oxygenation (ECMO) on the coagulation system in a sheep model. A ...

Bioengineering

Murine Model of Central Venous Stenosis using Aortocaval Fistula with an Outflow Stenosis

Central venous stenosis is an important entity contributing to arteriovenous fistula (AVF) failure. A murine AVF model was modified to create a partial ligation of the inferior vena cava (IVC) in the outflow of the fistula, mimicking central venous stenosis. Technical aspects of this model are introduced. The aorta and IVC are exposed, following an abdominal incision. The infra-renal aorta and IVC are dissected for proximal clamping, and the distal aorta is exposed for puncture. The IVC at the midpoint between the left renal vein and the aortic bifurcation is carefully dissected to place an 8-0 suture beneath the IVC. After clamping the aorta and IVC, an AVF is created by puncturing the infra-renal aorta through both walls into the IVC with a 25 G needle, followed by ligating a 22 G intra-venous (IV) catheter and IVC together. The catheter is then removed, creating a reproducible venous stenosis without occlusion. The aorta and IVC are unclamped after confirming primary hemostasis. This novel model of central vein stenosis is easy to perform, reproducible, and will facilitate studies on AVF failure.

An aortocaval fistula was created by puncturing the murine infra-renal aorta through both walls into the inferior vena cava and was followed by creation of a stenosis in its outflow via partial ligation of the inferior vena cava. This reproducible model can be used to study central venous stenosis.

Central venous stenosis is an important entity contributing to arteriovenous fistula (AVF) failure. A murine AVF model was modified to create a partial ...

A Mouse Model of Orotracheal Intubation and Ventilated Lung Ischemia Reperfusion Surgery

Ischemia reperfusion (IR) injury frequently results from processes that involve a transient period of interrupted blood flow. In the lung, isolated IR permits the experimental study of this specific process with continued alveolar ventilation, thereby avoiding the compounding injurious processes of hypoxia and atelectasis. In the clinical context, lung ischemia reperfusion injury (also known as lung IRI or LIRI) is caused by numerous processes, including but not limited to pulmonary embolism, resuscitated hemorrhagic trauma, and lung transplantation. There are currently limited effective treatment options for LIRI. Here, we present a reversible surgical model of lung IR involving first orotracheal intubation followed by unilateral left lung ischemia and reperfusion with preserved alveolar ventilation or gas exchange. Mice undergo a left thoracotomy, through which the left pulmonary artery is exposed, visualized, isolated, and compressed using a reversible slipknot. The surgical incision is then closed during the ischemic period, and the animal is awakened and extubated. With the mouse spontaneously breathing, reperfusion is established by releasing the slipknot around the pulmonary artery. This clinically relevant survival model permits the evaluation of lung IR injury, the resolution phase, downstream effects on lung function, as well as two-hit models involving experimental pneumonia. While technically challenging, this model can be mastered over the course of a few weeks to months with an eventual survival or success rate of 80%-90%.

A mouse surgical model to create left lung ischemia reperfusion (IR) injury while maintaining ventilation and avoiding hypoxia.

Ischemia reperfusion (IR) injury frequently results from processes that involve a transient period of interrupted blood flow. In the lung, isolated IR ...

Immunology and Infection

Multiple Intravenous Bolus Dosing and Invasive Hemodynamic Assessment in a Hypoxia-Induced Mouse Pulmonary Artery Hypertension Model

Pulmonary arterial hypertension (PAH) is a progressive life-threatening disease, primarily affecting small pulmonary arterioles of the lung. Currently, there is no cure for PAH. It is important to discover new compounds that can be used to treat PAH. The mouse hypoxia-induced PAH model is a widely used model for PAH research. This model recapitulates human clinical manifestations of PAH Group 3 disease and is an important research tool to evaluate the effectiveness of new experimental therapies for PAH. Research using this model often requires the administration of compounds in mice. For a compound that needs to be given directly into the bloodstream, optimizing intravenous (IV) administration is a key part of the experimental procedures. Ideally, the IV injection system should permit multiple injections over a set time course. Although the mouse hypoxia-induced PAH model is very popular in many laboratories, it is technically challenging to perform multiple IV bolus dosing and invasive hemodynamic assessment in this model. In this protocol, we present step-by-step instructions on how to carry out multiple IV bolus dosing via mouse jugular vein and perform arterial and right ventricle catheterization for hemodynamic assessment in mouse hypoxia-induced PAH model.

This protocol provides a step-by-step procedure for executing multiple intravenous bolus dose administration and invasive hemodynamic monitoring in mice. Investigators can use this protocol for future therapeutic compound screening for pulmonary artery hypertension.

Pulmonary arterial hypertension (PAH) is a progressive life-threatening disease, primarily affecting small pulmonary arterioles of the lung. Currently, ...

Veno-Arterial Extracorporeal Membrane Oxygenation for Cardiogenic Shock

Cardiogenic shock (CS) is a clinical condition characterized by inadequate tissue perfusion in the setting of low cardiac output. CS is the leading cause of death following acute myocardial infarction (AMI). Several temporary mechanical support devices are available for hemodynamic support in CS until clinical recovery ensues or until more definitive surgical procedures have been performed. Veno-arterial (VA) extracorporeal membrane oxygenation (ECMO) has evolved as a powerful treatment option for short-term circulatory support in refractory CS. In the absence of randomized clinical trials, the utilization of ECMO has been guided by clinical experience and based on data from registries and observational studies. Survival to hospital discharge with the use of VA-ECMO ranges from 28-67%. The initiation of ECMO requires venous and arterial cannulation, which can be performed either percutaneously or by surgical cutdown. Components of an ECMO circuit include an inflow cannula that draws blood from the venous system, a pump, an oxygenator, and an outflow cannula that returns blood to the arterial system. Management considerations post ECMO initiation include systemic anticoagulation to prevent thrombosis, left ventricle unloading strategies to augment myocardial recovery, prevention of limb ischemia with a distal perfusion catheter in cases of femoral arterial cannulation, and prevention of other complications such as hemolysis, air embolism, and Harlequin syndrome. ECMO is contraindicated in patients with uncontrolled bleeding, unrepaired aortic dissection, severe aortic insufficiency, and in futile cases such as severe neurological injury or metastatic malignancies. A multi-disciplinary shock team approach is recommended while considering patients for ECMO. Ongoing studies will evaluate whether the addition of routine ECMO improves survival in AMI patients with CS who undergo revascularization.

The following article highlights various steps involved in initiating and maintaining veno-arterial extracorporeal membrane oxygenation in patients with cardiogenic shock.

Cardiogenic shock (CS) is a clinical condition characterized by inadequate tissue perfusion in the setting of low cardiac output. CS is the leading cause ...

Veno-Venous Extracorporeal Membrane Oxygenation in a Mouse

Summary

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Chapters in this video

In vivo Measurement of the Mouse Pulmonary Endothelial Surface Layer

Ex Vivo Hepatic Perfusion Through the Portal Vein in Mouse

Normothermic Ex Vivo Liver Machine Perfusion in Mouse

Use of a Central Venous Line for Fluids, Drugs and Nutrient Administration in a Mouse Model of Critical Illness

Cardiopulmonary Bypass in a Mouse Model: A Novel Approach

Effects of Extracorporeal Membrane Oxygenation on the Coagulation System

Murine Model of Central Venous Stenosis using Aortocaval Fistula with an Outflow Stenosis

A Mouse Model of Orotracheal Intubation and Ventilated Lung Ischemia Reperfusion Surgery

Multiple Intravenous Bolus Dosing and Invasive Hemodynamic Assessment in a Hypoxia-Induced Mouse Pulmonary Artery Hypertension Model

Veno-Arterial Extracorporeal Membrane Oxygenation for Cardiogenic Shock