Male and female C57BL/6J mice between 8-14 weeks of age were used for these experiments. This protocol has received ethical approval from the Animal Care Committee at the University of Calgary and follows all animal care guidelines.
1. Mouse preparation and surgery
<ol>
	<li>Sterilize surgical tools (via bead sterilizer or autoclave).</li>
	<li>Weigh mouse for presurgical weight and analgesic dose.</li>
	<li>Place the mouse in an induction box with 4% isoflurane and 800 mL/min of oxygen. Confirm the anesthetic plane by pinching the toes and observing the animal for lack of reflex.</li>
	<li>Inject analgesic subcutaneously (0.1 mg/kg of Buprenorphine) (see Table of Materials).</li>
	<li>Place the mouse on a heated surgical pad during surgery and maintain anesthesia with 3% isoflurane delivered via nose cone. Apply ophthalmic ointment to avoid dry eyes to each eye.</li>
	<li>Shave the hair from the chest and neck surgical areas.</li>
	<li>Restrain the paws of the mouse and position them on the surgical table.</li>
	<li>Intubate the mouse by inserting a smooth tipped 23 G catheter into the trachea through the mouth and the pharynx.
	<ol>
		<li>Ventilate the mouse following intubation with 2% isoflurane and 100% oxygen as a carrier gas using a commercial Ventilator (see Table of Materials) set at a rate of 110 breaths/min, a tidal volume of 250 &#181;L, and positive end-expiratory pressure (PEEP) of 4 mmHg.</li>
	</ol>
	</li>
	<li>Roll the mouse 30% on its right side to position the left side of the chest for surgery.</li>
	<li>Clean the surgical area with 3 alternating scrubs of 70% ethanol and betadine in a circular motion(seeTable of Materials). Place sterile surgical drapes around the surgical area.</li>
	<li>Make a 2-3 cm lateral incision in the skin of the chest to visualize the pectoralis muscles on the left side. Cut the pectoralis major and minor using a 1 cm incision from the midline outward to visualize the intercostal muscles between the third and fourth ribs. 
	NOTE: Care should be taken to avoid excess bleeding from the pectoralis muscle through cauterization of bleeding vessels.</li>
	<li>Make a 2 cm incision in the left intercostal muscle to introduce air (by passive air movement) into the chest cavity to allow the heart and lungs to fall away from the surgical incision. Further, expand the opening with the help of a cautery device (see Table of Materials) to incise the intercostal and prevent bleeding 
	NOTE: The ventilator should be temporarily stopped during the period of cauterization to avoid explosive reactions with oxygen. Care is taken not to damage the pericardial sac.</li>
	<li>Using retractors, retract the ribs to expose the heart.</li>
	<li>Observe the pericardium and the underlying heart under a stereomicroscope. 
	NOTE: The mouse pericardium is thin enough to visualize the vasculature of the heart.</li>
	<li>Gently place forceps on the surface of the pericardium to reduce its movement and that of the underlying heart.</li>
	<li>Visually landmark the left anterior descending (LAD) coronary artery by tracing its emergence from under the left appendage.</li>
	<li>Using the micro-needle driver, guide an appropriate suture (see Table of Materials) through the pericardium, under the LAD, with the suture emerging on the other side of the LAD and pericardium. Tie the suture to restrict the blood flow through the coronary artery and trim the excess suture with the help of scissors (Figure 1A). 
	NOTE: Upon restricting blood flow to the coronary artery, blanching of the anterior portion of the left ventricle should be visible. This procedure represents a permanent ligation model. However, a transient ligation approach with different ischemia periods could also be applied at this stage.</li>
	<li>Below the incision site within the sterile area, introduce a 24 G catheter percutaneously into the chest (remove the guide needle after entering the chest cavity). Then, close the ribs followed by muscle layers and skin using an appropriate suture (a taper needle for muscle, conventional cutting needle for skin).</li>
	<li>Once the chest is closed, evacuate the remaining air from the chest cavity via the 24 G catheter using gentle suction with a 3 mL syringe and chest compressions. Once the air is removed, withdraw the 24 G catheter.</li>
	<li>Reduce the isoflurane to 1%.</li>
	<li>Turn off the isoflurane while maintaining ventilation with oxygen to allow the mouse to recover from anesthesia. Once the animal shows signs of breathing independently, remove the 23 G tracheal tube from the mouth and place the mouse in a recovery cage to be monitored for resumption of normal breathing.</li>
	<li>Allow the mouse to recover in the cage with a portion of the cage placed on a warming pad to provide an external heat source.</li>
	<li>Provide maintenance injections of analgesic (Buprenorphine 0.1 mg/kg, subcutaneously) every 12 h for 72 h post-surgery.</li>
	<li>Monitor the health status of mice daily for 7 days, which includes evaluating incisions and animal discomfort. 
	&#8203;NOTE: Due to the invasiveness of this procedure (thoracotomy), the administration of antibiotics may be necessary.</li>
</ol>
2. Functional assessment of cardiac function by echocardiography (ECG)
<ol>
	<li>Induce and maintain the mouse under general anesthesia with 1.5-2% isoflurane and 800 mL/min of oxygen.</li>
	<li>Place mouse in a supine position on a heated stage platform and attach the paws to the ECG leads.</li>
	<li>Shave the chest of the mouse.</li>
	<li>Acquire echocardiography images using a 40 MHz linear transducer probe and contact gel and analyze with the appropriate software (see Table of Materials).</li>
	<li>Turn off the isoflurane and allow the mouse to recover on the heating platform before returning the cage to an active state. 
	&#8203;NOTE: Echocardiography assessment is non-invasive and thus can be performed longitudinally throughout the experiment to determine changes before and after coronary ligation.</li>
</ol>
3. Heart tissue collection for fibrosis staining
<ol>
	<li>Sacrifice the mice with inhalation of 100% CO2 and carefully dissect out the heart. 
	NOTE: Using scissors and forceps, this is achieved by cutting through the large vessels entering (vena cava, pulmonary vein) and exiting (pulmonary artery, aorta) the heart to release it from the circulatory system in the thoracic cavity.</li>
	<li>Fix the heart in 10% formalin for at least 24 h.</li>
	<li>Cut samples using a straight razor blade through the right ventricle, interventricular septum, and left ventricle, ensuring that the incision went through the center of the infarct zone. Samples are then sent to the core facility for paraffin embedding.</li>
	<li>Cut tissue sections of 5 &#181;m thickness with a microtome and place them on glass slides for staining.</li>
	<li>Deparaffinize using commercial xylene and graded alcohol washes (2x 99%, 1x 95%, 1x 70%) with deionized water, then rehydrate.</li>
	<li>Stain with 0.1% Sirius red in picric acid for 2 h at room temperature.</li>
	<li>Wash sections with 0.5% acetic acid for 3 min and rinse with 70% ethanol for 1 min.</li>
	<li>Dehydrate the sections using the reverse order of washes outlined in 3.4, with increasing and graded ethanol concentrations then xylene.</li>
	<li>Mount tissue sections with a mounting solution (see Table of Materials) for microscopic evaluation.</li>
</ol>
4. Flow cytometry of heart and pericardial cavity lavage
<ol>
	<li>Sacrifice the mice with inhalation of 100% CO2 to effect.</li>
	<li>Place the mouse on its back and fix the arms and legs to a surgical board using tape.</li>
	<li>Carefully open the left side (right side from the experimenter view) of the thoracic cavity, starting with cutting the diaphragm to about the midpoint and subsequently cutting through the outside ribs towards the sternum. 
	NOTE: Avoid nicking large blood vessels, particularly those that run parallel to the sternum.</li>
	<li>Retract the ribs using a hemostat to expose the underlying heart and pericardium. 
	NOTE: The pericardium is very fragile, so be sure not to catch it with scissors during the cutting.</li>
	<li>Using a PE-10 tubing (see Table of Materials) catheter inserted into the pericardial space near the junction of the left atrium and left ventricle, inject 100 &#181;L of sterile saline into the pericardial cavity.
	<ol>
		<li>Allow saline to pool and collect from the posterior side of the heart, being careful not to puncture or tear the pericardium in the process. Repeat this step twice and place lavage fluid on ice while processing the heart.</li>
	</ol>
	</li>
	<li>Excise the heart by cutting the major vessels (aorta, pulmonary artery, vein, and vena cava) entering and exiting the heart. Remove the right and left atria and weigh the ventricular heart tissue.</li>
	<li>Mince the tissue in small 1 mm2 pieces using scissors and place in 10 mL of digestion buffer containing 450 U/mL of collagenase I, 125 U/mL of collagenase XI, 60 U/mL of DNase I, and 60 U/mL of hyaluronidase in PBS for 1 h at 37 &#176;C on an orbital shaker.</li>
	<li>Pass heart tissue homogenates through a 70 &#181;m cell strainer (see Table of Materials) and spin down at 60 x g for 5 min at 4 &#176;C to remove cardiac parenchymal cells.</li>
	<li>Collect the supernatant, pass through a 40 &#181;m cell strainer (see Table of Materials) for a single cell suspension, and spin down at 400 x g for 5 min at 4 &#176;C to pellet the cells.</li>
	<li>Perform fragment crystallizable (Fc) receptor blocking and staining of cellular markers on pericardial and cardiac cells as previously described8.</li>
	<li>Run samples on a flow cytometer.</li>
</ol>
5. Labeling pericardial macrophage using the Intercostal Approach to the Pleural Space (ICAPS) method9
<ol>
	<li>Sterilize surgical tools (via bead sterilizer or autoclave) and spray 70% ethanol before commencing.</li>
	<li>Induce and maintain the mouse under general anesthesia with 1.5-2% isoflurane and 800 mL/min of oxygen. Confirm the anesthetic plane by pinching the toes and observing the animal for lack of reflex.</li>
	<li>Inject analgesic subcutaneously (Buprenorphine 0.1 mg/Kg).</li>
	<li>Place the mouse on a heated surgical pad during surgery.</li>
	<li>Shave the right anterolateral thoracic area.</li>
	<li>Clean the surgical area with ethanol and betadine.</li>
	<li>Make a 3 cm long incision in the skin, and with forceps expose the rib cage.</li>
	<li>Load 5 &#181;L of fluorescent beads (commercially available fluorescent microspheres, 1 &#181;m, see Table of Materials) and 45 &#181;L of saline into a PE-10 tubing syringe catheter with a beveled tip.</li>
	<li>Guide the catheter into the intercostal space as previously described9, slowly inject the bead solution and remove the catheter in one motion.</li>
	<li>Close the skin using staples. 
	NOTE: Staples are used in place of sutures to minimize potential re-opening of the incision.</li>
	<li>Turn off the isoflurane, place the mouse in the recovery cage and monitor for complications over the first 24 h.</li>
</ol>

<ol><li>Virani, S. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Heart+disease+and+stroke+statistics-2020+update%3A+A+report+from+the+American+Heart+Association%5BTitle%5D)+AND+%22Circulation%22%5BJournal%5D)">Heart disease and stroke statistics-2020 update: A report from the American Heart Association.</a> Circulation. 141, 139(2020).</li>
<li>Iismaa, S. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Comparative+regenerative+mechanisms+across+different+mammalian+tissues%5BTitle%5D)+AND+%22NPJ+Regenerative+Medicine%22%5BJournal%5D)">Comparative regenerative mechanisms across different mammalian tissues.</a> NPJ Regenerative Medicine. 3 (6), eCollection 2018 (2018).</li>
<li>Bayat, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Progressive+heart+failure+after+myocardial+infarction+in+mice%5BTitle%5D)+AND+%22Basic+Research+in+Cardiology%22%5BJournal%5D)">Progressive heart failure after myocardial infarction in mice.</a> Basic Research in Cardiology. 97 (3), 206-213 (2002).</li>
<li>Virag, J. A., Lust, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Coronary+artery+ligation+and+intramyocardial+injection+in+a+murine+model+of+infarction%5BTitle%5D)+AND+%22Journal+of+Visualized+Experiments%22%5BJournal%5D)">Coronary artery ligation and intramyocardial injection in a murine model of infarction.</a> Journal of Visualized Experiments. 52, 2581(2011).</li>
<li>De Villiers, C., Riley, P. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Mouse+models+of+myocardial+infarction%3A+comparing+permanent+ligation+and+ischaemia-reperfusion%5BTitle%5D)+AND+%22Disease+Models+%26+Mechanisms%22%5BJournal%5D)">Mouse models of myocardial infarction: comparing permanent ligation and ischaemia-reperfusion.</a> Disease Models & Mechanisms. 13 (11), (2020).</li>
<li>Borlaug, B. A., Reddy, Y. N. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+role+of+the+pericardium+in+heart+failure%3A+Implications+for+pathophysiology+and+treatment%5BTitle%5D)+AND+%22JACC+Heart+Failure%22%5BJournal%5D)">The role of the pericardium in heart failure: Implications for pathophysiology and treatment.</a> JACC Heart Failure. 7 (7), 574-585 (2019).</li>
<li>Pfaller, M. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+importance+of+the+pericardium+for+cardiac+biomechanics%3A+from+physiology+to+computational+modeling%5BTitle%5D)+AND+%22Biomechanics+and+Modeling+in+Mechanobiology%22%5BJournal%5D)">The importance of the pericardium for cardiac biomechanics: from physiology to computational modeling.</a> Biomechanics and Modeling in Mechanobiology. 18 (2), 503-529 (2019).</li>
<li>Deniset, J. F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Gata6%28%2B%29+Pericardial+Cavity+Macrophages+Relocate+to+the+Injured+Heart+and+Prevent+Cardiac+Fibrosis%5BTitle%5D)+AND+%22Immunity%22%5BJournal%5D)">Gata6(+) Pericardial Cavity Macrophages Relocate to the Injured Heart and Prevent Cardiac Fibrosis.</a> Immunity. 51 (1), 131-140 (2019).</li>
<li>Weber, G. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Immune+targeting+of+the+pleural+space+by+intercostal+approach%5BTitle%5D)+AND+%22BMC+Pulmonary+Medicine%22%5BJournal%5D)">Immune targeting of the pleural space by intercostal approach.</a> BMC Pulmonary Medicine. 15, 14(2015).</li>
<li>Nakatani, T., Shinohara, H., Fukuo, Y., Morisawa, S., Matsuda, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Pericardium+of+rodents%3A+pores+connect+the+pericardial+and+pleural+cavities%5BTitle%5D)+AND+%22The+Anatomical+Record%22%5BJournal%5D)">Pericardium of rodents: pores connect the pericardial and pleural cavities.</a> The Anatomical Record. 220, 132-137 (1988).</li>
<li>Tyberg, J. V., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+relationship+between+pericardial+pressure+and+right+atrial+pressure%3A+an+intraoperative+study%5BTitle%5D)+AND+%22Circulation%22%5BJournal%5D)">The relationship between pericardial pressure and right atrial pressure: an intraoperative study.</a> Circulation. 73, 428-432 (1986).</li>
<li>Hamilton, D. R., Sas, R., Semlacher, R. A., Kieser Prieur, T. M., Tyberg, J. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+relationship+between+left+and+right+pericardial+pressures+in+humans%3A+an+intraoperative+study%5BTitle%5D)+AND+%22The+Canadian+Journal+of+Cardiology%22%5BJournal%5D)">The relationship between left and right pericardial pressures in humans: an intraoperative study.</a> The Canadian Journal of Cardiology. 27, 346-350 (2011).</li>
<li>Park, D. S. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Human+pericardial+proteoglycan+4+%28lubricin%29%3A+Implications+for+postcardiotomy+intrathoracic+adhesion+formation%5BTitle%5D)+AND+%22The+Journal+of+Thoracic+and+Cardiovascular+Surgery%22%5BJournal%5D)">Human pericardial proteoglycan 4 (lubricin): Implications for postcardiotomy intrathoracic adhesion formation.</a> The Journal of Thoracic and Cardiovascular Surgery. 156 (4), 1598-1608 (2018).</li>
</ol>

The authors have no conflicts to disclose.

Inducing an MI in a closed pericardium in rodents is unique and can have potentially significant applications. The procedure relies heavily on the surgeon&#39;s familiarity with the rodent model and rodent cardiac anatomy. Success is also dependent on the care given during three critical steps: intercostal muscle incision and rib retraction (Steps 1.11-1.13), creating the infarct (Step 1.17 ), and animal recovery (Steps 1.22-1.24).
The thoracotomy must be done diligently to avoid puncturing or lacerating the pericardium. The most crucial step of this protocol is the suturing of the LAD to induce an infarction. As is the case with all LAD ligation models, appropriate placement of the ligation suture on the LAD is critical: proximal ligation may result in a fatal MI, whereas distal ligation may not cause a functionally relevant MI. Landmarking the LAD in the approximate center of the heart avoids these issues. As LAD ligation is performed with the heart beating, gently stabilizing the heart with forceps can help minimize movement, allowing the LAD to be sutured without damaging it. Small vessel laceration on the epicardium can occur during this procedure. Minor bleeds will resolve over 2-3 days and will not contaminate the pericardial fluid. The rodent pericardium, especially in mice, is very thin and can be easily torn if the surgeon does not exercise caution. Lastly, the operator must pay close attention to the animal in the post-procedure (i.e., recovery) phase. The timing of stopping isoflurane and removing the endotracheal tube must be done methodically to ensure the rodent can self-ventilate. The mouse should also be monitored after recovery to ensure no post-operative complications necessitate immediate intervention before being put into animal housing facilities. Examples of these complications include hemothorax, pneumothorax, and the inability to regain consciousness after anesthesia.
Most current mouse models of MI require opening the pericardium to ligate the LAD, resulting in a non-intact pericardium. The present model is unique as it preserves the homeostatic aspect of the pericardial space during infarction, hence providing a more clinically relevant representation of an MI. Maintaining the pericardial space results in improved functional characteristics of the mouse heart compared to procedures that divide the pericardium. Preservation of the native pericardial fluid also provides significant benefits to research possibilities as well as infarct healing. Intra-pericardial pressure is significant11,12, while pericardial fluid contains proteins that promote non-fibrotic healing pathways13. Recent research has revealed that macrophages that reside in the pericardial fluid also play an essential role in cardiac tissue repair and healing8. The current protocol provides a specific labeling method to track the fate of these macrophages following MI. Other cells within the pericardial space can be similarly labeled to assess their role in cardiac remodeling. Animal models that maintain the pericardium may better preserve these crucial pathways, making them a more accurate representation of patients&#39; pathophysiology and the healing processes.
This model allows the user to study and manipulate the entire pericardial space, potentiating research that explores the complex healing and inflammatory pathways mediated by pericardial cells. This model also provides an improved rodent infarct model for research not focused on the pericardial space. The preserved pericardial injury pathways allow for infarcts to have more human relevancy. The significant limitations of this model lay in user skill due to its technical nature. If the surgeon is not proficient in tissue handling and surgical techniques, errors can result in a torn pericardium or mortality. Finally, to leverage the advantages of this protocol, users should be able to use established and advanced imaging modalities, such as echocardiography and microscopy.

To this day, cardiovascular disease (CVD) is recognized as the leading cause of death globally, resulting in a significant financial burden and reduction in patient quality of life1. Coronary artery disease (CAD) is a sub-type of CVD and plays an essential role in the development of myocardial infarction (MI), which is a chief contributor to mortality. By definition, MI results from irreversible injury to the myocardial tissue due to prolonged conditions of ischemia and hypoxia. Myocardial tissue lacks regeneration capacity, so injuries are permanent and result in the replacement of heart muscle with a fibrotic scar that can be initially protective but ultimately contributes to adverse cardiac remodeling and eventual heart failure2.
Although the management of patients with CAD has dramatically improved over the past few decades, chronic heart failure (CHF) secondary to ischemia affects many patients worldwide. For preventing and managing this epidemic, it is necessary to understand the underlying mechanisms more extensively and develop new therapeutic approaches. Moreover, past findings highlight the limitations of systemic therapy and the necessity of developing precise alternatives. Given investigating the molecular sequelae of MI in humans is affected by the ability to access infarcted tissue, animal models that recapitulate the characteristics and development of human MI and CHF related to CVD are indispensable.
As ideal animal models closely resemble a human disorder for structural and functional characteristics, disease etiology should guide their conception. In CAD, it is the chronic atherosclerotic stenosis of coronary arteries or acute thrombotic occlusion. Different methods have been developed and applied in various species of laboratory animals to induce coronary artery narrowing or occlusion. Such strategies can be broadly classified into two groups: (1) mechanical manipulation of a coronary artery to induce an MI and (2) expediting atherosclerosis to facilitate coronary narrowing leading to an MI. The first strategy usually involves either the ligation of a coronary artery or the placement of a stent within the artery. The second approach tends to rely on modifying the animal&#39;s diet to include high fat/cholesterol food. Some of the limitations of this latter approach include the lack of control on the timing and site of coronary occlusions.
In contrast, the surgical induction of MI or ischemia in an animal model has several advantages, such as location, precise timing, and extent of the coronary event, leading to more reproducible results. The most widely used method is surgical ligation of the left anterior descending coronary artery (LAD). Such models recapitulate human responses to acute ischemic injury, as well as the progression to CHF3. Initially developed in larger animals, LAD surgery on small animals such as rodents has become more feasible with advancements in technology4. In establishing such models, mice have been favored for various reasons, including their relative availability, low expense in housing, and their capacity for genetic manipulation.
Contemporary surgical models of ischemic heart disease using LAD occlusion require the researcher to open the pericardium to temporarily or permanently ligate the artery5. Such strategies result in the disruption of the pericardial space, which plays an essentially mechanical and lubricating function to ensure proper cardiac function. Another disadvantage of opening the pericardium is losing the animal&#39;s native pericardial fluid with its various cellular and protein components6,7. In response, a method to induce MI while keeping the pericardium intact was developed by us. In addition to minimizing the perturbation of this homeostatic environment, this approach allows for tagging and tracing specific cells after causing an MI. In addition, this approach better represents myocardial ischemic injury in the human setting.

<table><tbody><tr><td>Steri-350 Bead Sterilizer</td><td>Inotech</td><td>NC9449759</td><td></td></tr><tr><td>10% Formalin</td><td>Millipore Sigma</td><td>HT501128-4L</td><td></td></tr><tr><td>40 &amp;micro;m Cell strainer</td><td>VWR</td><td>CA21008-949</td><td>Falcon, 352340</td></tr><tr><td>70 &amp;micro;m Cell strainer</td><td>VWR</td><td>CA21008-952</td><td>Falcon, 352350</td></tr><tr><td>ACK Lysis Buffer</td><td>Thermo Fisher</td><td>A1049201</td><td></td></tr><tr><td>BD Insyte-W Catheter Needle 24 G X 3/4&quot;</td><td>CDMV Inc</td><td>108778</td><td></td></tr><tr><td>Betadine (10% povidone-iodine topical solution)</td><td>CDMV Inc</td><td>104826</td><td></td></tr><tr><td>Blunt Forceps</td><td>Fine Science Tools</td><td>FST 11000-12</td><td></td></tr><tr><td>BNP Ophthalmic Ointment</td><td>CDMV Inc</td><td>17909</td><td></td></tr><tr><td>Castroviejo Needle Driver</td><td>Fine Science Tools</td><td>FST 12061-01</td><td></td></tr><tr><td>Centrifuge 5810R</td><td>Eppendorf</td><td>22625101</td><td></td></tr><tr><td>Collagenase I</td><td>Millipore Sigma</td><td>SCR103</td><td></td></tr><tr><td>Collagenase XI</td><td>Millipore Sigma</td><td>C7657</td><td></td></tr><tr><td>Covidien 5-0 Polysorb Suture - CV-11 taper needle</td><td>Medtronic Canada</td><td>GL-890</td><td></td></tr><tr><td>Covidien 5-0 Polysorb Suture - PC-13 cutting needle</td><td>Medtronic Canada</td><td>SL-1659</td><td></td></tr><tr><td>Curved Blunt Forceps</td><td>Fine Science Tools</td><td>FST 11009-13</td><td></td></tr><tr><td>Dako Mounting Medium</td><td>Agilen</td><td>CS70330-2</td><td></td></tr><tr><td>DNase I</td><td>Millipore Sigma</td><td>11284932001</td><td></td></tr><tr><td>Ethanol, 100%</td><td>Millipore Sigma</td><td>MFCD00003568</td><td></td></tr><tr><td>Ethicon 8-0 Ethilon Suture - BV-130-4 taper needle</td><td>Johnson &amp;amp; Johnson Inc.</td><td>2815G</td><td></td></tr><tr><td>Fiber-Optic Light</td><td>Nikon</td><td>2208502</td><td></td></tr><tr><td>Fine Forceps</td><td>Fine Science Tools</td><td>FST 11150-10</td><td></td></tr><tr><td>Fluoresbrite&amp;reg; YG Carboxylate Microspheres 1.00 &amp;micro;m</td><td>Polysciences, Inc.</td><td>15702</td><td></td></tr><tr><td>Geiger Thermal Cautery Unit</td><td>World Precision Instruments</td><td>501293</td><td>Model 150-ST</td></tr><tr><td>Hyaluronidase</td><td>Millipore Sigma</td><td>H4272</td><td></td></tr><tr><td>Isofluorane Vaporizer</td><td>Harvard Apparatus</td><td>75-0951</td><td></td></tr><tr><td>Isoflurane USP, 250 mL</td><td>CDMV Inc</td><td>108737</td><td></td></tr><tr><td>Magnetic Fixator Retraction System</td><td>Fine Science Tools</td><td>18200-20</td><td></td></tr><tr><td>MX550D- 40 MHz probe</td><td>Fujifilm- Visual Sonics</td><td></td><td></td></tr><tr><td>Needle Driver</td><td>Fine Science Tools</td><td>FST 12002-12</td><td></td></tr><tr><td>PE-10 Tubing</td><td>Braintree Scienctific, Inc.</td><td>PE10 50 FT</td><td></td></tr><tr><td>Scissors</td><td>Fine Science Tools</td><td>FST 14184-09</td><td></td></tr><tr><td>SMZ-1B Stereo Microscope</td><td>Nikon</td><td>SMZ1-PS</td><td></td></tr><tr><td>VentElite Small Animal Ventilator</td><td>Harvard Apparatus</td><td>55-7040</td><td></td></tr><tr><td>Vetergesic (10 mL, 0.3mg/mL buprenorphine))</td><td>CDMV Inc</td><td>124918</td><td>controlled drug</td></tr><tr><td>Vevo 2100 Software</td><td>Fujifilm-Visual Sonics</td><td></td><td></td></tr></tbody></table>

an intact pericardium ischemic rodent model

This protocol has shown that the pericardium and its contents play an essential anti-fibrotic role in the ischemic rodent model (coronary ligation to induce myocardial injury). The majority of pre-clinical myocardial infarction models require the disruption of pericardial integrity with loss of the homeostatic cellular milieu. However, recently a methodology has been developed by us to induce myocardial infarction, which minimizes pericardial damage and retains the heart&#39;s resident immune cell population. An improved cardiac functional recovery in mice with an intact pericardial space following coronary ligation has been observed. This method provides an opportunity to study inflammatory responses in the pericardial space following myocardial infarction. Further development of the labeling techniques can be combined with this model to understand the fate and function of pericardial immune cells in regulating the inflammatory mechanisms that drive remodeling in the heart, including fibrosis.

This modified coronary ligation model has been optimized to achieve reproducibility and animal survival. However, due to the significant injury induced in the heart, some expected intra-operative and post-operative mortality are associated with the procedure. The standard mortality is typically higher in males (~25-35%) than in females (~ 10-15%).
Successful induction of an MI with the modified coronary ligation should be evident by changes in the heart&#39;s functional parameters and structural features. For function, decreases in parameters such as left ventricle (LV) ejection fraction as assessed by echocardiography will be noticeable within 3-4 weeks post-MI (Figure 1A). These functional changes should be accompanied by significant fibrosis of the free wall of the LV as assessed by histological staining such as picrosirius red (PR) (Figure 1B). For this analysis, the use of longitudinal cross-sections through the infarcted should allow representation of the infarcted area, peri-infarct and remote zones of the heart.
Maintaining an intact pericardium throughout the procedure provides an opportunity to study the concurrent inflammatory response in the pericardial cavity. It also allows for determining how immune cells within this compartment can contribute to ongoing remodeling processes. Combining the fluorescent bead labeling method with flow cytometry analysis provides one approach to track with high selectivity resident Gata6+ pericardial macrophages (GPCMs). This procedure involves injecting beads directly into the pleural space. These are equally taken up by resident Gata6 macrophages in both the pleural and pericardial cavities (Figure 2A) due to communication between this two cavities10. Importantly, little to no labeling should be detected in the heart or blood (Figure 2A). Once labeled, the relocation of the cells following inflammatory challenges such as MI can be tracked by flow cytometry (Figure 2B) and/or imaging. For avoiding any potential inflammatory effects from the ICAPS procedure, this labeling should be performed one week before subsequent interventions.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62720/62720fig01.jpg" /> 
Figure 1: Intact pericardial coronary ligation model induces functional and structural alterations in the heart. (A) Schematic timeline and LV ejection fraction quantification at baseline or 4 weeks post-coronary ligation for animals with disrupted or intact pericardium. Data are represented as mean &#177; SD. ***= p &#60; 0.001, *= p &#60; 0.05 vs baseline, one-way ANOVA. Adapted from Deniset JF, et al., with permission from Elsevier8. (B) Representative images and quantification of picrosirius red fibrosis staining in mouse heart cross-sections at 4 weeks post-infarct with the disrupted or intact pericardial coronary ligation models. <a href="https://www.jove.com/files/ftp_upload/62720/62720fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62720/62720fig02.jpg" /> 
Figure 2: Labeling and tracking of pericardial cavity macrophages following MI. (A) Representative flow cytometry plots of fluorescent bead containing myeloid cells from the pleural cavity, pericardial cavity, cardiac tissue, and blood at baseline or 7 days following local injection of fluorescent beads using the ICAPS method. Bottom panels- Characterization of bead and cells in the pericardial cavity as predominantly Gata6+ pericardial macrophages (GPCMs). (B) Flow cytometry analysis and quantification of fluorescent bead labeled pericardial myeloid cells in pericardial cavity and heart with or without MI. *= p &#60; 0.05, ** = p &#60; 0.01. Adapted from Deniset JF, et al., with permission from Elsevier8. <a href="https://www.jove.com/files/ftp_upload/62720/62720fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about An Intact Pericardium Ischemic Rodent Model at JoVE.com

An Intact Pericardium Ischemic Rodent Model

This protocol outlines the steps for inducing myocardial infarction in mice while preserving the pericardium and its contents.

This protocol has shown that the pericardium and its contents play an essential anti-fibrotic role in the ischemic rodent model (coronary ligation to ...

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2.4K Views.  Cumming School of Medicine. This protocol outlines the steps for inducing myocardial infarction in mice while preserving the pericardium and its contents.

Video: An Intact Pericardium Ischemic Rodent Model

Acute Myocardial Infarction in Rats

With heart failure leading the cause of death in the USA (Hunt), biomedical research is fundamental to advance medical treatments for cardiovascular diseases. Animal models that mimic human cardiac disease, such as myocardial infarction (MI) and ischemia-reperfusion (IR) that induces heart failure as well as pressure-overload (transverse aortic constriction) that induces cardiac hypertrophy and heart failure (Goldman and Tarnavski), are useful models to study cardiovascular disease. In particular, myocardial ischemia (MI) is a leading cause for cardiovascular morbidity and mortality despite controlling certain risk factors such as arteriosclerosis and treatments via surgical intervention (Thygesen). Furthermore, an acute loss of the myocardium following myocardial ischemia (MI) results in increased loading conditions that induces ventricular remodeling of the infarcted border zone and the remote non-infarcted myocardium. Myocyte apoptosis, necrosis and the resultant increased hemodynamic load activate multiple biochemical intracellular signaling that initiates LV dilatation, hypertrophy, ventricular shape distortion, and collagen scar formation. This pathological remodeling and failure to normalize the increased wall stresses results in progressive dilatation, recruitment of the border zone myocardium into the scar, and eventually deterioration in myocardial contractile function (i.e. heart failure). The progression of LV dysfunction and heart failure in rats is similar to that observed in patients who sustain a large myocardial infarction, survive and subsequently develops heart failure (Goldman). The acute myocardial infarction (AMI) model in rats has been used to mimic human cardiovascular disease; specifically used to study cardiac signaling mechanisms associated with heart failure as well as to assess the contribution of therapeutic strategies for the treatment of heart failure. The method described in this report is the rat model of acute myocardial infarction (AMI). This model is also referred to as an acute ischemic cardiomyopathy or ischemia followed by reperfusion (IR); which is induced by an acute 30-minute period of ischemia by ligation of the left anterior descending artery (LAD) followed by reperfusion of the tissue by releasing the LAD ligation (Vasilyev and McConnell). This protocol will focus on assessment of the infarct size and the area-at-risk (AAR) by Evan's blue dye and triphenyl tetrazolium chloride (TTC) following 4-hours of reperfusion; additional comments toward the evaluation of cardiac function and remodeling by modifying the duration of reperfusion, is also presented. Overall, this AMI rat animal model is useful for studying the consequence of a myocardial infarction on cardiac pathophysiological and physiological function.

The rat model of acute myocardial infarction (AMI) is useful to study the consequence of a MI on cardiac pathophysiological and physiological function.

With heart failure leading the cause of death in the USA (Hunt), biomedical research is fundamental to advance medical treatments for cardiovascular ...

Coronary Artery Ligation and Intramyocardial Injection in a Murine Model of Infarction

Mouse models are a valuable tool for studying acute injury and chronic remodeling of the myocardium in vivo. With the advent of genetic modifications to the whole organism or the myocardium and an array of biological and/or synthetic materials, there is great potential for any combination of these to assuage the extent of acute ischemic injury and impede the onset of heart failure pursuant to myocardial remodeling. 
Here we present the methods and materials used to reliably perform this microsurgery and the modifications involved for temporary (with reperfusion) or permanent coronary artery occlusion studies as well as intramyocardial injections. The effects on the heart that can be seen during the procedure and at the termination of the experiment in addition to histological evaluation will verify efficacy. 
Briefly, surgical preparation involves anesthetizing the mice, removing the fur on the chest, and then disinfecting the surgical area. Intratracheal intubation is achieved by transesophageal illumination using a fiber optic light. The tubing is then connected to a ventilator. An incision made on the chest exposes the pectoral muscles which will be cut to view the ribs. For ischemia/reperfusion studies, a 1 cm piece of PE tubing placed over the heart is used to tie the ligature to so that occlusion/reperfusion can be customized. For intramyocardial injections, a Hamilton syringe with sterile 30gauge beveled needle is used. When the myocardial manipulations are complete, the rib cage, the pectoral muscles, and the skin are closed sequentially. Line block analgesia is effected by 0.25% marcaine in sterile saline which is applied to muscle layer prior to closure of the skin. The mice are given a subcutaneous injection of saline and placed in a warming chamber until they are sternally recumbent. They are then returned to the vivarium and housed under standard conditions until the time of tissue collection. At the time of sacrifice, the mice are anesthetized, the heart is arrested in diastole with KCl or BDM, rinsed with saline, and immersed in fixative. Subsequently, routine procedures for processing, embedding, sectioning, and histological staining are performed. 
Nonsurgical intubation of a mouse and the microsurgical manipulations described make this a technically challenging model to learn and achieve reproducibility. These procedures, combined with the difficulty in performing consistent manipulations of the ligature for timed occlusion(s) and reperfusion or intramyocardial injections, can also affect the survival rate so optimization and consistency are critical.

Numerous genetic manipulations and/or intramyocardial injections of genes, proteins, cells, and/or biomaterials are superimposed upon the dimension of time in studies of acute ischemia/ reperfusion injury and chronic remodeling in mice. This video illustrates the microsurgical procedures for ischemia/reperfusion, permanent coronary artery ligation, and intramyocardial injection studies.

Mouse models are a valuable tool for studying acute injury and chronic remodeling of the myocardium in vivo.  With the advent of genetic modifications to ...

A Murine Closed-chest Model of Myocardial Ischemia and Reperfusion

Surgical trauma by thoracotomy in open-chest models of coronary ligation induces an immune response which modifies different mechanisms involved in ischemia and reperfusion. Immune response includes cytokine expression and release or secretion of endogenous ligands of innate immune receptors. Activation of innate immunity can potentially modulate infarct size. We have modified an existing murine closed-chest model using hanging weights which could be useful for studying myocardial pre- and postconditioning and the role of innate immunity in myocardial ischemia and reperfusion. This model allows animals to recover from surgical trauma before onset of myocardial ischemia.
Volatile anesthetics have been intensely studied and their preconditioning effect for the ischemic heart is well known. However, this protective effect precludes its use in open chest models of coronary artery ligation. Thus, another advantage could be the use of the well controllable volatile anesthetics for instrumentation in a chronic closed-chest model, since their preconditioning effect lasts up to 72 hours. Chronic heart diseases with intermittent ischemia and multiple hit models are other possible applications of this model.
For the chronic closed-chest model, intubated and ventilated mice undergo a lateral blunt thoracotomy via the 4th intercostal space. Following identification of the left anterior descending a ligature is passed underneath the vessel and both suture ends are threaded through an occluder. Then, both suture ends are passed through the chest wall, knotted to form a loop and left in the subcutaneous tissue. After chest closure and recovery for 5 days, mice are anesthetized again, chest skin is reopened and hanging weights are hooked up to the loop under ECG control.
At the end of the ischemia/reperfusion protocol, hearts can be stained with TTC for infarct size assessment or undergo perfusion fixation to allow morphometric studies in addition to histology and immunohistochemistry.

Surgical trauma induces an inflammatory response. Cytokines and endogenous ligands are known to modulate myocardial infarct size following ischemia and reperfusion. We present a modified closed-chest model of murine ischemia and reperfusion using hanging weights to minimize effects of thoracotomy.

Surgical trauma by thoracotomy in open-chest models of coronary ligation induces an immune response which modifies different mechanisms involved in ...

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 ...

Murine Isolated Heart Model of Myocardial Stunning Associated with Cardioplegic Arrest

The following protocol is of use to evaluate impaired cardiac function or myocardial stunning following moderate ischemic insults. The technique is useful for modeling ischemic injury associated with numerous clinically relevant phenomenon including cardiac surgery with cardioplegic arrest and cardiopulmonary bypass, off-pump CABG, transplant, angina, brief ischemia, etc. The protocol presents a general method to model hypothermic hyperkalemic cardioplegic arrest and reperfusion in rodent hearts focusing on measurement of myocardial contractile function. In brief, a mouse heart is perfused in langendorff mode, instrumented with an intraventricular balloon, and baseline cardiac functional parameters are recorded. Following stabilization, the heart is then subject to brief infusion of a cardioprotective hypothermic cardioplegia solution to initiate diastolic arrest. Cardioplegia is delivered intermittently over 2 hr. The heart is then reperfused and warmed to normothermic temperatures and recovery of myocardial function is monitored. Use of this protocol results in reliable depressed cardiac contractile function free from gross myocardial tissue damage in rodents.

The goal of this protocol to assess myocardial stunning following ischemic cardioplegic arrest in rodents.

The following protocol is of use to evaluate impaired cardiac function or myocardial stunning following moderate ischemic insults. The technique is useful ...

A Chronic Cardiac Ischemia Model in Swine Using an Ameroid Constrictor

Cardiovascular disease remains the number one cause of mortality in the United States. There are numerous approaches to treating these diseases, but regardless of the approach, an in vivo model is needed to test each treatment. The pig is one of the most used large animal models for cardiovascular disease. Its heart is very similar in anatomy and function to that of a human. The ameroid placement technique creates an ischemic area of the heart, which has many useful applications in studying myocardial infarction. This model has been used for surgical research, pharmaceutical studies, imaging techniques, and cell therapies.
There are several ways of inducing an ischemic area in the heart. Each has its advantages and disadvantages, but the placement of an ameroid constrictor remains the most widely used technique. The main advantages to using the ameroid are its prevalence in existing research, its availability in various sizes to accommodate the anatomy and size of the vessel to be constricted, the surgery is a relatively simple procedure, and the post-operative monitoring is minimal, since there are no external devices to maintain. This paper provides a detailed overview of the proper technique for the placement of the ameroid constrictor.

The purpose of this protocol is to demonstrate the placement of a delayed constricting device (an ameroid constrictor) around a coronary artery in a swine model. This device creates an ischemic area of the heart that is useful for studying new diagnostic imaging techniques and new methods of treatment.

Cardiovascular disease remains the number one cause of mortality in the United States. There are numerous approaches to treating these diseases, but ...

Murine Myocardial Infarction Model using Permanent Ligation of Left Anterior Descending Coronary Artery

Myocardial infarction (MI) and acute coronary diseases are among the most prominent causes of death in population with western lifestyle. The murine models of MI with permanent ligation of left-anterior descending (LAD) coronary artery closely mimics MI in humans. Murine models benefit from the extensive genetic engineering available nowadays. Here we propose a reproducible murine surgical model of myocardial infarction by permanent LAD coronary ligation. Our technique comprises anesthesia with ketamine/xylazine that can be rapidly reversed by administration of an antagonist, intubation without tracheotomy for mechanical-assisted ventilation, ventilation with application of extrinsic positive end-expiratory pressure (PEEP) to avoid alveolar collapse, a thoracotomy method limiting to the minimum surgical lesions made to skeletal muscles, and lung inflation without thoracentesis. This method is sparsely invasive, reproducible and reduces post-surgery mortality and complications.

Herein we describe a surgical procedure showing how to achieve permanent ligation of the left-anterior descending coronary artery in mice. This model is of high relevance to investigate the pathophysiology of myocardial infarction and the concomitant biological processes.

Myocardial infarction (MI) and acute coronary diseases are among the most prominent causes of death in population with western lifestyle. The murine ...

Improved Rodent Model of Myocardial Ischemia and Reperfusion Injury

Myocardial ischemia and reperfusion injury (MIRI), induced by coronary heart disease (CHD), causes damage to the cardiomyocytes. Furthermore, evidence suggests that thrombolytic therapy or primary percutaneous coronary intervention (PPCI) does not prevent reperfusion injury. There is still no ideal animal model for MIRI. This study aims to improve the MIRI model in rats to make surgery easier and more feasible. A unique method for establishing MIRI is developed by using a soft tube during a key step of the ischemic period. To explore this method, thirty rats were randomly divided into three groups: sham group (n = 10); experimental model group (n = 10); and existing model group (n = 10). Findings of triphenyltetrazolium chloride staining, electrocardiography, and percent survival are compared to determine the accuracies and survival rates of the operations. Based on the study results, it has been concluded that the improved surgery method is associated with a higher survival rate, elevated ST-T segment, and larger infarct size, which is expected to mimic the pathology of MIRI better.

Myocardial ischemia-reperfusion model of rat heart is improved by using a self-made retractor, polyvinyl chloride tube, and a unique knotting method. Electrocardiogram, triphenyltetrazolium chloride and histological staining, and percent survival analysis results showed that the improved model group has higher success and survival rates than the already existing model group.

Myocardial ischemia and reperfusion injury (MIRI), induced by coronary heart disease (CHD), causes damage to the cardiomyocytes. Furthermore, evidence ...

Rodent Working Heart Model for the Study of Myocardial Performance and Oxygen Consumption

Isolated working heart models have been used to understand the effects of loading conditions, heart rate and medications on myocardial performance in ways that cannot be accomplished in vivo. For example, inotropic medications commonly also affect preload and afterload, precluding load-independent assessments of their myocardial effects in vivo. Additionally, this model allows for sampling of coronary sinus effluent without contamination from systemic venous return, permitting assessment of myocardial oxygen consumption. Further, the advent of miniaturized pressure-volume catheters has allowed for the precise quantification of markers of both systolic and diastolic performance. We describe a model in which the left ventricle can be studied while performing both volume and pressure work under controlled conditions. 
In this technique, the heart and lungs of a Sprague-Dawley rat (weight 300-500 g) are removed en bloc under general anesthesia. The aorta is dissected free and cannulated for retrograde perfusion with oxygenated Krebs buffer. The pulmonary arteries and veins are ligated and the lungs removed from the preparation. The left atrium is then incised and cannulated using a separate venous cannula, attached to a preload block. Once this is determined to be leak-free, the left heart is loaded and retrograde perfusion stopped, creating the working heart model. The pulmonary artery is incised and cannulated for collection of coronary effluent and determination of myocardial oxygen consumption. A pressure-volume catheter is placed into the left ventricle either retrograde or through apical puncture. If desired, atrial pacing wires can be placed for more precise control of heart rate. This model allows for precise control of preload (using a left atrial pressure block), afterload (using an afterload block), heart rate (using pacing wires) and oxygen tension (using oxygen mixtures within the perfusate).

Isolated working heart models can be used to measure the effect of loading conditions, heart rate, and medications on myocardial performance and oxygen consumption. We describe methods for preparation of a rodent left heart working model that permits study of systolic and diastolic performance and oxygen consumption under various conditions.

Isolated working heart models have been used to understand the effects of loading conditions, heart rate and medications on myocardial performance in ways ...

Biology

Rodent Model of Intestinal Ischemia-Reperfusion Injury via Occlusion of the Superior Mesenteric Artery

Intestinal ischemia-reperfusion injury (IRI) is associated with a myriad of conditions in both veterinary and human medicine. Intestinal IRI conditions, such as gastric dilatation volvulus (GDV), mesenteric torsion, and colic, are observed in animals such as dogs and horses. An initial interruption of blood flow causes tissues to become ischemic. Although necessary to salvage viable tissue, subsequent reperfusion can induce further injury. The main mechanism responsible for IRI is free radical formation upon reperfusion and reintroduction of oxygen into damaged tissue, but there are many other components involved. The resulting local and systemic effects often impart a poor prognosis. 
Intestinal IRI has been the subject of extensive research over the past 50 years. An in vivo rodent model in which the base of the superior mesenteric artery (SMA) is temporarily ligated is currently the most common method used to study intestinal IRI. Here, we describe a model of intestinal IRI utilizing isoflurane anesthesia in 21% O2 medical air that yields reproducible injury, as demonstrated by consistent histopathology of the small intestines. Tissue injury was also assessed in the colon, liver, and kidneys.

We describe how to generate a widely used surgical model of intestinal ischemia-reperfusion injury (IRI) in rodents. The procedure involves occlusion of the superior mesenteric artery followed by the restoration of blood flow. This model is useful for studies investigating occlusive causes of intestinal IRI in both veterinary and human medicine.

Intestinal ischemia-reperfusion injury (IRI) is associated with a myriad of conditions in both veterinary and human medicine. Intestinal IRI conditions, ...