The authors have no acknowledgements.

All animal experiments were performed in compliance with the German Animal Protection Law (TierSchG) and were approved by the local animal welfare committee (Lower Saxony State Office for Consumer Protection and Food Safety, Protocol TSA 14/1556). The minimal weight of mouse suitable for this model is 25 g.
1. Preoperative Preparations
NOTE: All procedures are carried out under clean, non-sterile conditions, with autoclaved instruments.
<ol>
	<li>Place 50-60 8-cm long propylene hollow fibers in parallel in a tube and connect with a T-adapter on both sides. Fill the space between the hollow fibers and the external branch of the T-adapter with glue.
	<ol>
		<li>Allow at least 24 h for the glue to harden. Cut the hollow fibers extending out from the T-adapter using a standard microtome and pull the silicone tube over to the connection sites.</li>
	</ol>
	</li>
	<li>Insert a 27 G metal needle (from a 26G branule)&#160;into a 2 Fr polyurethane cannula. Use a surgical blade and micro-scissors under a microscope (8-12X magnification) to make three to four fenestrations of about 0.15 cm each within the distal third of the cannula to assure optimal venous return flow.
	<ol>
		<li>Remove the wire when completed. Use cotton swabs for blunt dissection and retraction of structures. Use gauze swabs (5 x 5 cm2) to soak up excess fluid to prevent tissue dehydration.</li>
	</ol>
	</li>
	<li>Prepare the priming volume. For completion, add 30 IU of heparin per mL priming solution and 2.5% v/v of an 8.4% solution of NaHCO3 for buffering. Store this solution at 4 &#176;C until used.</li>
	<li>Fill the CPB circuit (i.e., pump, air trapper, tubing and both cannulas) with 850 &#181;L priming solution via the venous cannula. Once filled, keep the CPB circulating until the animal is ready for cannulation.</li>
</ol>
2. Animal Anesthesia
<ol>
	<li>To administer anesthesia, place the animal in an induction chamber under 2.5% v/v isoflurane/oxygen mixture. Confirm proper anesthesia by assessing pedal withdrawal reflex, tail pinch reflex, and eye blink reflex. Apply veterinary eye ointment to avoid eye dryness.</li>
	<li>Place the animal on a warming pad with temperature-regulating function. Measure body temperature with a probe inserted rectally and connected to the data acquisition system.</li>
	<li>After full anesthesia is achieved, intubate the animal orotracheally using a 20G plastic braunule, inserting it orally and pushing it into the trachea under visual control. Start mechanical ventilation using an isoflurane vaporizer connected to a mouse ventilator. Depending on the animal weight, adjust the ventilation so that a tidal volume of 250 - 350 &#181;L is achieved.</li>
	<li>To assure complete analgesia, inject 5 mg/kg animal bodyweight carprofen subcutaneously. It is recommended that isoflurane concentration be kept between 1.3 - 2.5% in oxygen. Isoflurane concentration may be manually adjusted depending on the stage of the procedure.</li>
</ol>
3. Surgical Procedures
<ol>
	<li>After full anesthesia and intubation is achieved, perform a midline skin incision to the neck with sharp fine scissors, separate muscles in a blunt fashion, and expose the right jugular vein and left carotid artery. During preparation, coagulate small vessels using a veterinary coagulator connected to bipolar forceps to ensure minimal blood loss.</li>
	<li>After preparation of the jugular vessels, cranially ligate the distal segment of the left common carotid artery to its bifurcation using 8-0 silk sutures.</li>
	<li>Connect the distal end of a 27 G cannula to the arterial inflow tubing using a silicone connector (0.5 mm ID, 1 mm OD), place 8-0 silk suture slip knots at the proximal segment of the artery, and insert the tip of the cannula into the carotid artery.</li>
	<li>After correct placement of the cannula tip, advance the cannula tip so that it lays 3 - 4 mm proximally to the aortic arch. Fix the cannula tip by securing with 8-0 silk knots.</li>
	<li>Using microsurgical forceps and scissors, perform blunt and sharp dissection, expose the right jugular vein by blunt preparation of tissue laterally to the sternocleidomastoid muscle, close to clavicle, and place an 8-0 silk knot at the distal end and an 8-0 loop at the proximal end.</li>
	<li>After ligation of the distal end of the jugular vein, place the tip of the venous cannula in the right jugular vein and progress towards the right atrium and secure using the silk knot. For optimal venous return, it may be necessary to advance the venous tip into the right ventricle.</li>
	<li>Once the correct cannula position is achieved, carry out systemic heparinization by adding 2.5 IU of heparin per gram of the animal bodyweight via direct intravenous injection into the jugular vein.</li>
	<li>For real-time invasive pressure monitoring, cannulate the left femoral artery. Expose the groin area and gently separate the common femoral artery from the femoral vein and femoral nerve under 25X magnification.
	<ol>
		<li>Ligate the distal part of the femoral artery. Temporarily occlude the proximal part with a slip-knot and make a small incision on the anterior wall using micro-forceps.</li>
		<li>Afterwards, insert a 1 Fr polyurethane cannula into the femoral artery and secure it with a silk 10-0 suture. This cannula is used to extract blood samples for blood gas analysis.</li>
	</ol>
	</li>
	<li>After successful placement of the arterial and venous cannulas, initiate cardiopulmonary bypass by turning on the pump. The time from intubation to starting of CPB is approximately 45 min. Start using a flow rate of 0.5 mL/min, depending on systemic pressure measurements, and increase blood flow within 2 min to full flow (4 - 6 mL/min) by increasing the pump speed.</li>
	<li>Under full monitoring, perform an upper sternotomy using sharp scissors starting from the manubrium and going 5 mm in the caudal direction. Coagulate the sternal edges immediately to avoid bleeding. Expose the aortic arch by pulling the right carotid artery in the cranial direction.</li>
	<li>For optimal control, circumferentially free the aortic arch from the thymus and surrounding tissue to facilitate clamping. Place a 8-0 silk loop around the ascending aorta. To place the cross clamp for cardioplegia, pull the silk loop in the cranial direction to better expose the ascending aorta.</li>
</ol>
4. Cardiopulmonary Bypass and Blood Gas Analysis
<ol>
	<li>For blood gas analysis (BGA), use glass capillary tubes to collect 60 - 90 &#181;L arterial blood via the femoral artery catheter.
	<ol>
		<li>Use a small clamp on the silicone tubing and detach the tubing from the catheter. Release pressure slowly on the clamp to allow controlled filling of the capillary tube.</li>
		<li>Reattach the silicone tube to the catheter. Insert capillary tubes into the blood gas analyzer for measurements at the following time points: 
		10 min after initiation of CPB with ventilation (BGA1) 
		after 25 min of CPB and 15 min of respiratory arrest (BGA2) 
		after 40 min of CPB and 30 min of respiratory arrest (BGA3) 
		after 55 min of CPB, 45 min of respiratory, and 20 min of cardiac arrest (BGA4)</li>
	</ol>
	</li>
	<li>Under stable CPB flow, initiate respiratory arrest by stopping ventilation.</li>
	<li>After termination of ventilation, set the warming pad to 28 &#186;C and&#160;start topically cooling the animal to 28 &#186;C body temperature within the first 20 min of respiratory arrest using gauze soaked in cold saline.</li>
	<li>Once a body temperature of 28 &#186;C is achieved, and after a total of 30 min of respiratory arrest, administer 0.3 mL of 7.45% KCl solution into the CPB circuit to initiate cardioplegia.</li>
	<li>For cross clamping of the aorta, pull the previously placed silk loop (step 3.10) in the cranial direction for better exposure and place a micro-serrefine clamp on the ascending part of the aorta.</li>
	<li>Perform a total of 60 min respiratory arrest and 30 min of cardiac arrest. Remove the micro-serrefine clamp from the ascending aorta to initiate reperfusion of the heart. Simultaneously, re-ventilate and warm the animal to 37 &#176;C.</li>
	<li>After reperfusion is completed and the animal has reached normothermia, terminate the CPB by turning off the pump and continue to monitor physiological measurements (e.g. body temperature, ECG, and invasive blood pressure) for 20 min.</li>
	<li>At the end of the experiment, exsanguinate the animal under full anesthesia (5% isoflurane) and collect blood and organs for further analysis14.</li>
</ol>

<ol><li>Edmunds, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Cardiopulmonary+Bypass+after+50+Years%5BTitle%5D)+AND+%22N.+Engl.+J.+Med%22%5BJournal%5D)">Cardiopulmonary Bypass after 50 Years.</a> N. Engl. J. Med. 351 (16), 1601-1603 (2004).</li>
<li>Goto, T., Maekawa, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Cerebral+dysfunction+after+coronary+artery+bypass+surgery%5BTitle%5D)+AND+%22J.+Anesth%22%5BJournal%5D)">Cerebral dysfunction after coronary artery bypass surgery.</a> J. Anesth. 28 (2), 242-248 (2014).</li>
<li>Uysal, S., Reich, D. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Neurocognitive+outcomes+of+cardiac+surgery%5BTitle%5D)+AND+%22J.+Cardiothorac.+Vasc.+Anesth%22%5BJournal%5D)">Neurocognitive outcomes of cardiac surgery.</a> J. Cardiothorac. Vasc. Anesth. 27 (5), 958-971 (2013).</li>
<li>Ballaux, P. K., Gourlay, T., Ratnatunga, C. P., Taylor, K. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+literature+review+of+cardiopulmonary+bypass+models+for+rats%5BTitle%5D)+AND+%22Perfusion%22%5BJournal%5D)">A literature review of cardiopulmonary bypass models for rats.</a> Perfusion. 14 (6), 411-417 (1999).</li>
<li>Jungwirth, B., de Lange, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Animal+models+of+cardiopulmonary+bypass%3A+development%2C+applications%2C+and+impact%5BTitle%5D)+AND+%22Semin.+Cardiothorac.+Vasc.+Anesth%22%5BJournal%5D)">Animal models of cardiopulmonary bypass: development, applications, and impact.</a> Semin. Cardiothorac. Vasc. Anesth. 14 (2), 136-140 (2010).</li>
<li>Günzinger, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+rat+model+of+cardiopulmonary+bypass+with+cardioplegic+arrest+and+hemodynamic+assessment+by+conductance+catheter+technique%5BTitle%5D)+AND+%22Basic+Res+Cardiol%22%5BJournal%5D)">A rat model of cardiopulmonary bypass with cardioplegic arrest and hemodynamic assessment by conductance catheter technique.</a> Basic Res Cardiol. 102 (6), 508-517 (2007).</li>
<li>Waterbury, T., Clark, T. J., Niles, S., Farivar, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Rat+model+of+cardiopulmonary+bypass+for+deep+hypothermic+circulatory+arrest%5BTitle%5D)+AND+%22J.+Thorac.+Cardiovasc.+Surg%22%5BJournal%5D)">Rat model of cardiopulmonary bypass for deep hypothermic circulatory arrest.</a> J. Thorac. Cardiovasc. Surg. 141 (6), 1549-1551 (2011).</li>
<li>Schnoering, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+newly+developed+miniaturized+heart-lung+machine-expression+of+inflammation+in+a+small+animal+model%5BTitle%5D)+AND+%22Artif.+Organs%22%5BJournal%5D)">A newly developed miniaturized heart-lung machine-expression of inflammation in a small animal model.</a> Artif. Organs. 34 (11), 911-917 (2010).</li>
<li>Kim, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+responses+of+tissues+from+the+brain%2C+heart%2C+kidney%2C+and+liver+to+resuscitation+following+prolonged+cardiac+arrest+by+examining+mitochondrial+respiration+in+rats%5BTitle%5D)+AND+%22Oxid.+Med.+Cell.+Longev%22%5BJournal%5D)">The responses of tissues from the brain, heart, kidney, and liver to resuscitation following prolonged cardiac arrest by examining mitochondrial respiration in rats.</a> Oxid. Med. Cell. Longev. 2016, (2016).</li>
<li>Shappell, S. B., Gurpinar, T., Lechago, J., Suki, W. N., Truong, L. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Chronic+obstructive+uropathy+in+severe+combined+immunodeficient+%28SCID%29+mice%3A+lymphocyte+infiltration+is+not+required+for+progressive+tubulointerstitial+injury%5BTitle%5D)+AND+%22J.+Am.+Soc.+Nephrol%22%5BJournal%5D)">Chronic obstructive uropathy in severe combined immunodeficient (SCID) mice: lymphocyte infiltration is not required for progressive tubulointerstitial injury.</a> J. Am. Soc. Nephrol. 9 (6), 1008-1017 (1998).</li>
<li>Majzoub, J. A., Muglia, L. J. <a target="_blank" href="http://web.mit.edu/hst.160/www/Knockout.pdf">Knockout mice.</a> N. Engl. J. Med. , 904-907 (1996).</li>
<li>Houser, S. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Animal+Models+of+Heart+Failure+A+Scientific+Statement+From+the+American+Heart+Association%5BTitle%5D)+AND+%22Circ.+Res%22%5BJournal%5D)">Animal Models of Heart Failure A Scientific Statement From the American Heart Association.</a> Circ. Res. 111 (1), 131-150 (2012).</li>
<li>Russell, J. C., Proctor, S. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Small+animal+models+of+cardiovascular+disease%3A+tools+for+the+study+of+the+roles+of+metabolic+syndrome%2C+dyslipidemia%2C+and+atherosclerosis%5BTitle%5D)+AND+%22Cardiovasc.+Pathol%22%5BJournal%5D)">Small animal models of cardiovascular disease: tools for the study of the roles of metabolic syndrome, dyslipidemia, and atherosclerosis.</a> Cardiovasc. Pathol. 15 (6), 318-330 (2006).</li>
<li>Iurascu-Gagea, M., Craig, S. Euthanasia and necropsy. <a target="_blank" href="https://books.google.com/books?id=HhEs-xsYp6IC&lpg=PP1&ots=kQsV14dfx9&dq=%22The%20laboratory%20rabbit%2C%20guinea%20pig%2C%20hamster%2C%20and%20other%20rodents%22&lr&pg=PP1#v=onepage&q=%22The%20laboratory%20rabbit,%20guinea%20pig,%20hamster,%20and%20other%20rodents%22&f=false">The laboratory rabbit, guinea pig, hamster, and other rodents</a>. Suckow, M. A., Stevens, K. A., Wilson, R. P. , Academic Press (Elsevier). 117-141 (2012).</li>
</ol>

The authors have nothing to disclose.

We have developed a fully-functioning clinically-relevant model of CPB in a mouse. With more than thirty strains of mice having cardiovascular diseases, our model could be a starting point for development of new prospective protocols related to CPB. Moreover, due to the plethora of mouse-specific reagents and knockout-out mice, this model can not only replace the current rat model of CPB but will facilitate dissection of the molecular mechanisms involved in CPB-related organ damage. To date, CPB has not been applied in m...

Since the introduction of cardiopulmonary bypass (CPB) into the clinic, it has played an essential role in cardiac surgery1. In modern cardiac surgery, prolonged CPB time is essential to perform extensive aortic reconstructions and combined procedures. Although technological advances have been tremendous, the use of extracorporal circulation is associated with intra- and postoperative systemic and local organ damage2,3.
Large animal models have been developed to investigate the role of CPB on physiological processes4,5. Although these models have provided insight into some of the CPB associated complications, they are extremely costly and molecular tools (e.g., antibodies) are very limited. A more cost-efficient alternative has been developed in small animals. Since their development, multiple studies have been conducted to optimize a CPB model in rats and rabbits5,6,7,8,9. These models provide a good basis for measurements of pathophysiological disease processes; however, they are still insufficient to investigate cellular and humoral immunology due to the lack of relevant antibodies and reagents. This impairs their role in this field of research.
We have recently developed a mouse model of CPB. Due to a wide variety of mouse-specific reagents and genetically-modified mice, mouse models are in general the model of choice for physiological, molecular, and immunological research10,11. Therefore, our model will facilitate the study of CPB in relation to various comorbidities as there are many mice strains available with clinically-relevant diseases12,13. Accordingly, this paper describes, in detail, how to perform CPB in mice. Oxygen and hemodynamic parameters are closely monitored after deep respiratory and circulatory arrest.

<table><tbody><tr><td>Sterofundin</td><td>B.Braun Petzold GmbH</td><td>PZN:8609189</td><td>priming volume, 1:1 with Tetraspan</td></tr><tr><td>Tetraspan 6% HES Solution</td><td>B. Braun Melsungen AG</td><td>PZN: 05565416</td><td>priming volume, 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 solution</td></tr><tr><td>NaHCO&lt;sub&gt;3&lt;/sub&gt; 8,4% Solution</td><td>B. Braun Melsungen AG</td><td>PZN: 1579775</td><td>3% in priming solution</td></tr><tr><td>KCL 7.45 % Solution</td><td>B. Braun Melsungen AG</td><td>PZN: 2418577</td><td>0.1 mL for cardioplegia</td></tr><tr><td>Carprofen</td><td>Zoetis Inc., USA</td><td>PZN:00289615 08859153</td><td>5 mg/kg/BW</td></tr><tr><td>1 Fr PU Catheter</td><td>Instechlabs INC., USA</td><td>C10PU-MCA1301</td><td>carotid 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>Vasofix Safety catheter 20G</td><td>B.Braun Medical</td><td>4268113S-01</td><td>orotracheal intubation</td></tr><tr><td>8-0 Silk suture braided</td><td>Ashaway Line &amp;amp; Twine Mfg. Co., USA</td><td>75290</td><td>ligature</td></tr><tr><td>Isoflurane</td><td>Piramal Critical Care Deutschland GmbH</td><td>PZN:9714675</td><td>narcosis</td></tr><tr><td>CLINITUBES blood capillaries</td><td>Radiomed GmbH</td><td>51750132</td><td>blood sampling 60 - 95 microliter</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>MiniVent Ventilator for Mice (Model 845)</td><td>Harvard Apparatus</td><td>73-0044</td><td>mechanical ventilation</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>PowerLab data acquisition device 4/35</td><td>ADInstruments Ltd, New Zealand</td><td>PL3504</td><td>invasive pressure, ECG, temperature</td></tr><tr><td>ABL 800 Flex</td><td>Radiometer GmbH</td><td></td><td>blood gas analysis</td></tr><tr><td>NMRI mice</td><td>Charles River Laboratories</td><td>Crl:NMRI(Han)</td><td>male, 30 - 35 g, 12 weeks old, housed at least 1 week before the experiment</td></tr></tbody></table>

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 protocol describes the perfusion circuit, surgical procedures, and monitoring of physiological parameters during CPB of a mouse. When performed by an adequately skilled microsurgeon, the results are consistently and reproducibly obtained.
To maintain adequate tissue perfusion, the mean arterial pressure is always kept between 40 and 60 mmHg by adjusting the CPB blood flow and adding of extra volume. Depending on the weight ...

Watch this Scientific Journal Video about Cardiopulmonary Bypass in a Mouse Model: A Novel Approach at JoVE.com

Cardiopulmonary Bypass in a Mouse Model: A Novel Approach

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 ...

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10.9K Views.  Hannover Medical School. 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.

Video: Cardiopulmonary Bypass in a Mouse Model: A Novel Approach

Murine Cervical Heart Transplantation Model Using a Modified Cuff Technique

Mouse models are of special interest in research since a wide variety of monoclonal antibodies and commercially defined inbred and knockout strains are available to perform mechanistic in vivo studies. While heart transplantation models using a suture technique were first successfully developed in rats, the translation into an equally widespread used murine equivalent was never achieved due the technical complexity of the microsurgical procedure. In contrast, non-suture cuff techniques, also developed initially in rats, were successfully adapted for use in mice1-3. This technique for revascularization involves two major steps I) everting the recipient vessel over a polyethylene cuff; II) pulling the donor vessel over the formerly everted recipient vessel and holding it in place with a circumferential tie. This ensures a continuity of the endothelial layer, short operating time and very high patency rates4.
Using this technique for vascular anastomosis we performed more than 1,000 cervical heart transplants with an overall success rate of 95%. For arterial inflow the common carotid artery and the proximal aortic arch were anastomosed resulting in a retrograde perfusion of the transplanted heart. For venous drainage the pulmonary artery of the graft was anastomosed with the external jugular vein of the recipient5.
Herein, we provide additional details of this technique to supplement the video.

The murine cervical heart transplantation model is well suited for immunological as well as ischemia reperfusion injury studies. We modified the procedure using a non-suture cuff technique and performed more than 1,000 successful transplants with this approach.
Herein, we provide additional details of this technique to supplement the video.

Mouse models are of special interest in research since a wide variety of monoclonal antibodies and commercially defined inbred and knockout strains are ...

A Murine Model of Myocardial Ischemia-reperfusion Injury through Ligation of the Left Anterior Descending Artery

Acute or chronic myocardial infarction (MI) are cardiovascular events resulting in high morbidity and mortality. Establishing the pathological mechanisms at work during MI and developing effective therapeutic approaches requires methodology to reproducibly simulate the clinical incidence and reflect the pathophysiological changes associated with MI. Here, we describe a surgical method to induce MI in mouse models that can be used for short-term ischemia-reperfusion (I/R) injury as well as permanent ligation. The major advantage of this method is to facilitate location of the left anterior descending artery (LAD) to allow for accurate ligation of this artery to induce ischemia in the left ventricle of the mouse heart. Accurate positioning of the ligature on the LAD increases reproducibility of infarct size and thus produces more reliable results. Greater precision in placement of the ligature will improve the standard surgical approaches to simulate MI in mice, thus reducing the number of experimental animals necessary for statistically relevant studies and improving our understanding of the mechanisms producing cardiac dysfunction following MI. This mouse model of MI is also useful for the preclinical testing of treatments targeting myocardial damage following MI.

We introduce a surgical method to induce experimental ischemia/reperfusion (I/R) injury to simulate myocardial infarction (MI) in mouse models that allows for more clarity in positioning of the ligation on the left anterior descending artery (LAD) to increase the reproducibility of MI experiments in mice.

Acute or chronic myocardial infarction (MI) are cardiovascular events resulting in high morbidity and mortality. Establishing the pathological mechanisms ...

A Modified Method for Heterotopic Mouse Heart Transplantion

Mice are often used as heart transplant donors and recipients in studies of transplant immunology due to the wide range of transgenic mice and reagents available. A difficulty is presented due to the small size of the animal and the considerable technical challenges of the microsurgery involved in heart transplantation. In particular, a high rate of technical failure early after transplantation may result from recipient death and post-operative complications such as hind limb paralysis or a non-beating heart. Here, the complete technique for heterotopic mouse heart transplantation is demonstrated, involving harvesting the donor heart and its subsequent implantation into a recipient mouse. The donor heart is harvested immediately following in situ perfusion with cold heparinized saline and transection of the ascending aorta and pulmonary artery. The recipient operation involves preparation of the abdominal aorta and inferior vena cava (IVC), followed by end-to-side anastomosis of the donor aorta with the recipient aorta using a single running 10-0 microsuture and a similar anastomosis of the donor pulmonary artery with the recipient IVC. Following the operation the animal is injected with 0.6 ml normal saline subcutaneously and allowed to recover on a 37&#160;&#176;C heating pad. The results from 227 mouse heart transplants are summarized with a success rate at 48 hr of 86.8%. Of the 13.2% failures within 48 hr, 5 (2.2%) experienced hind limb paralysis, 10 (4.4%) had a non-beating heart due to graft ischemic injury and/or thrombosis, while 15 (6.6%) died within 48 hr.

A technique is demonstrated for the microsurgical procedure for heterotopic transplantation of hearts in mice, including simplified methods for donor harvesting and recipient vessel anastomosis.

Mice are often used as heart transplant donors and recipients in studies of transplant immunology due to the wide range of transgenic mice and reagents ...

Heterotopic Cervical Heart Transplantation in Mice

The heterotopic cervical heart transplantation in mice is a valuable tool in transplant and cardiovascular research. The cuff technique greatly simplifies this model by avoiding challenging suture anastomoses of small vessels thereby reducing warm ischemia time. In comparison to abdominal graft implantation the cervical model is less invasive and the implanted graft is easily accessible for further follow-up examinations. Anastomoses are performed by pulling the ascending aorta of the graft over the cuff with the recipient&#8217;s common carotid artery and by pulling the main pulmonary artery over the cuff with the external jugular vein. Selection of appropriate cuff size and complete mobilization of the vessels are important for successful revascularization. Ischemia-reperfusion (I/R) injury can be minimized by perfusing the graft with a cardioplegic solution and by hypothermia. In this article, we provide technical details for a simplified and improved cuff technique, which should allow surgeons with basic microsurgical skills to perform the procedure with a high success rate.

The cuff technique shortens and facilitates the model of heterotopic cervical heart transplantation in mice by avoiding technically challenging suture anastomoses of small vessels. The technical details shown in this video paper should allow researches to establish this model in their laboratories.

The heterotopic cervical heart transplantation in mice is a valuable tool in transplant and cardiovascular research. The cuff technique greatly simplifies ...

A Novel Microsurgical Model for Heterotopic, En Bloc Chest Wall, Thymus, and Heart Transplantation in Mice

Exploration of novel strategies in organ transplantation to prolong allograft survival and minimizing the need for long-term maintenance immunosuppression must be pursued. Employing vascularized bone marrow transplantation and co-transplantation of the thymus have shown promise in this regard in various animal models.1-11 Vascularized bone marrow transplantation allows for the uninterrupted transfer of donor bone marrow cells within the preserved donor microenvironment, and the incorporation of thymus tissue with vascularized bone marrow transplantation has shown to increase T-cell chimerism ultimately playing a supportive role in the induction of immune regulation. The combination of solid organ and vascularized composite allotransplantation can uniquely combine these strategies in the form of a novel transplant model. Murine models serve as an excellent paradigm to explore the mechanisms of acute and chronic rejection, chimerism, and tolerance induction, thus providing the foundation to propagate superior allograft survival strategies for larger animal models and future clinical application. Herein, we developed a novel heterotopic en bloc chest wall, thymus, and heart transplant model in mice using a cervical non-suture cuff technique. The experience in syngeneic and allogeneic transplant settings is described for future broader immunological investigations via an instructional manuscript and video supplement.

To study combined solid organ and vascularized composite allotransplantation, we describe a novel heterotopic en bloc chest wall, thymus, and heart transplant model in mice using a cervical non-suture cuff technique.

Exploration of novel strategies in organ transplantation to prolong allograft survival and minimizing the need for long-term maintenance immunosuppression ...

A Recovery Cardiopulmonary Bypass Model Without Transfusion or Inotropic Agents in Rats

Cardiopulmonary bypass (CPB) is indispensable in cardiovascular surgery. Despite the dramatic refinement of CPB technique and devices, multi-organ complications related to prolonged CPB still compromise the outcome of cardiovascular surgeries, and may worsen postoperative morbidity and mortality. Animal models recapitulating the clinical usage of CPB enable the clarification of the pathophysiological processes that occur during CPB, and facilitate pre-clinical studies to develop strategies protecting against these complications. Rat CPB models are advantageous because of their greater cost-effectiveness, convenient experimental processes, abundant testing methods at the genetic or protein levels, and genetic consistency. They can be used for investigating the immune system activation and synthesis of proinflammatory cytokines, compliment activation, and production of oxygen free radicals. The rat models have been refined and have gradually taken the place of large-animal models. Here, we describe a simple CPB model without transfusion and/or inotropic agents in a rat. This recovery model allows the study of the long-term multiple organ sequelae of CPB.

Here, we present a protocol to describe a simple recovery cardiopulmonary bypass model without transfusion or inotropic agents in a rat. This model allows the study of the long-term multiple organ sequelae of cardiopulmonary bypass.

Cardiopulmonary bypass (CPB) is indispensable in cardiovascular surgery. Despite the dramatic refinement of CPB technique and devices, multi-organ ...

Immunology and Infection

Optimization of the Cuff Technique for Murine Heart Transplantation

Murine cardiac transplantation has been performed for more than 40 years. With advancements in microsurgery, certain new techniques have been used to improve surgical efficiency. In our lab, we have optimized the cuff technique with two major steps. First, we used the inner tube technique to insert a temporary inner tube into the external jugular vein and carotid artery blood vessel to facilitate eversion of the vessel over the cuff. Second, we performed complete heterotopic cardiac transplantation through the collaboration of two experienced surgeons. These modifications effectively reduced the operation time to 25 minutes, with a success rate of 95%. In this report, we describe these procedures in detail and provide a supplemental video. We believe that this report on the improved cuff technique will offer practical guidance for murine heterotopic heart transplantation and will enhance the utility of this mouse model for basic research.

We introduce an inner tube approach to the cuff technique for mouse cervical heterotopic heart transplantation to help evert the vessel over the cuff. We found that cooperation between two experienced surgeons markedly shortens the operation time.

Murine cardiac transplantation has been performed for more than 40 years. With advancements in microsurgery, certain new techniques have been used to ...

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.

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 ...

Mouse Heterotopic Cervical Cardiac Transplantation Utilizing Vascular Cuffs

Murine models of cardiac transplantation are frequently utilized to study ischemia-reperfusion injury, innate and adaptive immune responses after transplantation, and the impact of immunomodulatory therapies on graft rejection. Heterotopic cervical heart transplantation in mice was first described in 1991 using sutured anastomoses and subsequently modified to include cuff techniques. This modification allowed for improved success rates, and since then, there have been multiple reports that have proposed further technical improvements. However, translation into more widespread utilization remains limited due to the technical difficulty associated with graft anastomoses, which requires precision to achieve adequate length and caliber of the cuffs to avoid vascular anastomotic twisting or excessive tension, which can result in damage to the graft. The present protocol describes a modified technique for performing heterotopic cervical cardiac transplantation in mice which involves cuff placement on the recipient's common carotid artery and the donor's pulmonary artery in alignment with the direction of the blood flow.

Mouse cardiac transplantation models represent valuable research tools for studying transplantation immunology. The present protocol details mouse heterotopic cervical cardiac transplantation that involves the placement of cuffs on the recipient's common carotid artery and the donor's pulmonary artery trunk to allow for laminar blood flow.

Murine models of cardiac transplantation are frequently utilized to study ischemia-reperfusion injury, innate and adaptive immune responses after ...

Blood Circuit Reconstruction in an Abdominal Mouse Heart Transplantation Model

The surgical technique of heterotopic abdominal heart transplantation in mice is a standard model for research in transplantation immunology. Here, the established technique for a modified blood circuit reconstruction in a heterotopic abdominal heart transplantation model is presented. This method uses the intrathoracic inferior vena cava (IIVC) instead of the pulmonary artery of the donor heart for the anastomosis to the inferior vena cava of the recipient. It is facilitating and improving success rates for abdominal heart transplantation in mice.

A novel technique for blood circuit reconstruction in a heterotopic abdominal mouse heart transplantation model is demonstrated.

The surgical technique of heterotopic abdominal heart transplantation in mice is a standard model for research in transplantation immunology. Here, the ...