Name: murine left pulmonary hilar clamp model of lung ischemia reperfusion injury
Uploaded: 2024-04-12T13:10:00.000+00:00
Duration: 6 min 45 s
Description: Watch this Scientific Journal Video about Author Spotlight: Insights into the Effect of Ischemia Reperfusion on Lung Transplantation at JoVE.com

This work received no specific grant from any funding agency in the public, commercial, or not-for-profit sectors.

All studies were approved by the Institutional Animal Care and Use Committee at Washington University School of Medicine. Animals received humane care in compliance with the Guide for the care and use of laboratory animals, 8th edition21 prepared by the National Academy of Sciences and published by the National Institutes of Health, and the Principles of laboratory animal care formulated by the National Society for Medical Research.
1. Anesthesia and intubation
<ol>
	<li>Select a mouse that weighs at least 25 g. This will facilitate easier intubation due to the larger orifice between the vocal cords.</li>
	<li>Inject the mouse intraperitoneally with a mixture of ketamine (dose 100 mg/kg of body weight) and xylazine (dose 10 mg/kg of body weight). This mixture should be loaded into a &#189; cc syringe with a &#189; inch 29G needle. Wait about 5 min for ketamine to take effect - the mouse should not exhibit spontaneous movement and does not respond to a toe pinch, which confirms adequate anesthesia.</li>
	<li>Inject buprenorphine (dose 0.05 mg/kg of body weight) subcutaneously prior to surgery for additional pain control.</li>
	<li>Apply non-medicated ophthalmic ointment over both eyes to avoid corneal desiccation during surgery.</li>
	<li>Shave the mouse with a clipper over the left chest and back (see Figure 1A), extending onto the abdomen if laparotomy is planned (see step 5.3).</li>
	<li>Once the mouse is anesthetized, perform the intubation using a preferred intubation set-up and the remainder of the procedure under a microscope over a warmed mat at ~37 &#176;C (to maintain normothermia in the mouse).</li>
	<li>After achieving an adequate view of the vocal cords, insert a homemade introducer (see Supplementary Figure 1) with a curve on the end into a 1-inch 20G angiocatheter, which is used as the endotracheal tube (ETT). Guide the tip of the introducer and then the ETT between and past the vocal cords. 
	NOTE: It is important to visualize the ETT going through the vocal cords to avoid false intubation into the esophagus. Care should be taken to avoid injury to the vocal cords during intubation (i.e., the number of attempts of intubation should be limited to 5 and the ETT should not be advanced if resistance is met).</li>
	<li>Once the ETT is inserted, remove the introducer, and connect the ETT to a small animal ventilator. Observe for symmetric chest rise to confirm correct endotracheal intubation. The ventilator settings should be 100-105 breaths per minute and fraction of inspired oxygen of 100%. Tidal volume is 0.35 mL and positive end-expiratory pressure is 1 cm H2O. 
	NOTE: Ensure that the chest rather than abdomen is rising as accidental esophageal intubation will lead to death if not promptly corrected.</li>
	<li>Once confirmed to be in position, secure the ETT with a 5 cm strip of 1 inch silk tape around the mouse nose, ensuring&#160;adequate contact with the ETT and nose so that the ETT will not slide out of the mouth (see Figure 1B-C).</li>
	<li>To maintain adequate anesthesia throughout the surgery, administer 1%-1.5% isoflurane in-line with the oxygen flow.</li>
</ol>
2. Thoracotomy
<ol>
	<li>Position the mouse in right lateral decubitus position, with both front paws taped to the top left corner, the right hind paw taped to the bottom left corner, and the left hind paw taped to the bottom right corner (see Figure 2G for taping).</li>
	<li>Disinfect with skin over the left chest with at least 3 alternating rounds of povidone iodine followed by 70% alcohol application.</li>
	<li>Perform an incision over the 4th intercostal space from the anterior axillary line to the posterior axillary line, using scissors to open the overlying skin (see Figure 2A). The 4th intercostal space is located approximately 1 cm below the axilla in the mid-axillary line.</li>
	<li>To minimize blood loss, coagulate visible blood vessels within the subcutaneous and muscle layers using a handheld cautery pen.</li>
	<li>Divide the latissimus dorsi and serratus anterior muscles sharply using scissors along the length of the skin incision (see Figure 2B).</li>
	<li>Using fine curved forceps, carefully elevate the 4th rib (taking care not to injure the underlying lung) and make a pinpoint snip with fine scissors just above the 5th rib to enter the 4th intercostal space (see Figure 2C). Perform the thoracotomy above the 5th rib rather than below the 4th rib to avoid injuring the intercostal neurovascular bundle.</li>
	<li>Once the negative pleural cavity pressure is lost and the lung retracts away from the chest wall, extend the thoracotomy anteriorly and posteriorly in the 4th intercostal space just above the 5th rib (see Figure 2D-E). The entire length of the incision should be long enough to expose the entire lung, usually ~1 cm. 
	NOTE: Internal mammary artery bleeding can occur if the thoracotomy is extended too far anteriorly - if encountered, this should be cauterized to avoid excessive blood loss.</li>
	<li>Apply two rib retractors to spread open the rib space, allowing for a working window of at least 1 cm2 for adequate visualization (see Figure 2F-G).</li>
</ol>
3. Application of hilar clamp
<ol>
	<li>Using two-pointed cotton-tipped applicators, mobilize the left lung by elevating it cephalad and then bluntly dividing the translucent inferior pulmonary ligament (see Figure 3A-B). 
	NOTE: If this is challenging to perform without tearing the lung, the inferior pulmonary ligament can also be divided sharply using fine scissors.</li>
	<li>Reflect the lung anteriorly so that the posterior left pulmonary hilum can be visualized. Cut a piece of 6-0 silk tie to ~10 cm and place the midpoint of this tie posterior to the hilum (see Figure 3C).</li>
	<li>Then, flip the left lung posteriorly to lay over the tie and pull both ends of the tie anteriorly (see Figure 3D).</li>
	<li>With the two free ends (A and B) of the tie, instrument-tie a reversible slipknot with forceps and a curved mosquito clamp. See detailed description in steps 3.4.1 and 3.4.2. The warm ischemic time starts upon tying of the slipknot. The left lung should no longer be inflating with each ventilator breath once the knot is tied. It should become pale white implying cessation of perfusion. 
	NOTE: Care should be taken to make sure the knot sits in the middle of the hilum, avoiding inadvertently snaring the left atrium centrally and the lung parenchyma laterally (see Figure 3F).
	<ol>
		<li>With the clamp in the dominant hand and forceps in the non-dominant hand, hold the end of A with forceps and loop the midpoint of A around the closed clamp once (see Figure 3E).</li>
		<li>Grasp the midpoint of B with the clamp and pull with both hands to cinch the slipknot. Do not let the end of B pull through the knot, which would render the knot irreversible. After tying the knot, B should have a loop emanating from the knot, while A should be straight (see Figure 3F). The knot can be released by pulling the end of B.</li>
	</ol>
	</li>
	<li>Confirm adequate occlusion of the bronchus by manually occluding the outflow tubing from the ETT to the ventilator, which should cause the left lung to not inflate (see Figure 3G). Vascular occlusion is assumed with bronchial occlusion given the increased collapsibility of the pulmonary artery and veins compared to the cartilaginous bronchus.</li>
	<li>After application of the hilar clamp, close the skin incision with one simple interrupted stitch using 6-0 nylon suture, to minimize insensible fluid losses during the period of warm ischemia.</li>
</ol>
4. Release of hilar clamp
<ol>
	<li>After the desired period of warm ischemia, release the slipknot by gently pulling on the short free end of the silk tie (B from Figure 3F and step 3.4). Start the reperfusion time upon release of the slipknot. 
	NOTE: Following release of the tie, lung ventilation can be confirmed by again manual occlusion of the outflow of the ETT, which should now cause expansion of the left lung. Again, perfusion is implied with inflation of the lung, however, it can also be confirmed by the lung pinking up (see Figure 4A).</li>
	<li>During the reperfusion period, close the chest in three layers to minimize insensible losses. First, close the 4th rib space with a single stitch with 6-0 nylon suture, with one bite just above the 4th rib and one bite just above the 6th rib (see Figure 4B-C). Tie this at maximum lung inflation, to minimize risk of an iatrogenic pneumothorax (see Figure 4D-E).</li>
	<li>Next, close the muscle layer using a simple running stitch with 6-0 nylon suture.</li>
	<li>Finally, close the skin incision with a simple interrupted stitch with 6-0 nylon suture (see Figure 4F). Avoid running stitches on the skin due to the risk of awake animals picking at their incision and leading to complete wound dehiscence.</li>
	<li>From this point, the lung can be directly harvested at the end of the desired reperfusion time. If ABG evaluation is desired, see step 5. If additional intravenous injection is desired prior to sacrifice, see step 6. This is a terminal surgery.</li>
	<li>If reperfusion time is intended to be greater than a couple of hours, turn off the isoflurane to allow the mouse to awaken from anesthesia and extubate. Extubation criteria include twitching with paw pinch and spontaneous respirations and movement. Waking a mouse from ketamine and isoflurane anesthesia typically takes 30-45 min. This is a survival surgery.
	<ol>
		<li>For survival surgery, inject 1 mL of warm saline subcutaneously to account for fluid losses from surgery. Inject buprenorphine (dose 0.05 - 0.1 mg/kg of body weight) subcutaneously prior to surgery for pain control. Repeat anesthesia every 4-6 h&#160;for at least 3 days after surgery. Consider a bupivacaine block along the incision for additional pain control.</li>
	</ol>
	</li>
</ol>
5. ABG evaluation
NOTE: If ABG measurement is desired, this is best obtained via arterial blood aspiration from the left ventricle. To ascertain that the ABG reflects only left lung function, this arterial blood should be obtained after about 4 min of the right hilum being clamped22,23, during which time only the left lung is performing oxygenation and ventilation.
<ol>
	<li>If the mouse has been awoken from anesthesia after hilar clamp, re-anesthetize and intubate the mouse according to step 1.</li>
	<li>About 15-20 min prior to the end of the desired reperfusion time, position the mouse supine, securing all four limbs with tape. 
	NOTE: Ample time should be allotted for the following steps to be performed prior to the end of reperfusion. Specific timing to open the abdomen and chest will vary depending on the surgeon and their experience.</li>
	<li>Perform midline laparotomy from pubic bone to xiphoid, incising with scissors the skin followed by the abdominal wall along the linea alba (see Figure 5A-B).</li>
	<li>At the xiphoid, extend the laparotomy left and right to the anterior axillary line, following the curve of the inferior-most rib (see Figure 5C).</li>
	<li>From the abdomen, incise the anterior diaphragm in the midline to enter the chest, taking care not to go too deep to avoid injuring the heart (see Figure 5D-E). Then, extend the anterior diaphragm incision left and right to the anterior axillary line, along the inferior-most rib (see white dotted line in Figure 5D).</li>
	<li>Divide the bilateral ribs along the anterior axillary line, extending upwards towards the axilla, to create a clamshell thoracotomy. Then, reflect the anterior chest wall (sternum and bilateral anterior ribs) cephalad, which allows for full exposure of the heart and bilateral lungs (see white dotted lines in Figure 5F).</li>
	<li>Apply a clamp onto the anterior chest wall that is flipped cephalad to facilitate retraction. Retract the diaphragm downward with a curved mosquito clamp in the midline to improve visualization of the chest (see Figure 5F).</li>
	<li>To fully mobilize the right lung (which has 4 lobes), return the accessory lobe that extends past midline into the left chest back into the right chest (see Figure 5G-H). There is a thin ligament attaching this lobe to the left chest - divide this either bluntly with cotton-tipped applicators or sharply with scissors. 
	NOTE: This accessory lobe crosses the midline posterior to the inferior vena cava (IVC), so care must be taken not to injure the IVC during this maneuver.</li>
	<li>Once all lobes are returned to the right chest, reflect the entire right lung anteriorly and place another 6-0 silk tie (cut to ~10 cm) posterior to the right hilum. Then, replace the right lung back into the chest over the tie.</li>
	<li>Tie another slipknot around the right hilum using the same technique as described in step 3.4, taking care to encircle all four lobes of the right lung. This right hilar clamp step should be timed to be approximately 4 min prior to the end of reperfusion.</li>
	<li>Coat a &#189; inch 31G needle on a 1 cc tuberculin syringe with about 200 &#181;L of heparin 1000 unit/mL. To do this, draw up the volume of heparin and repeatedly pull the plunger back and forth 3-4 times to allow for the heparin to coat the entire inside of the syringe. This is done to minimize the risk of aspirated blood coagulating during transport from the surgical station to the ABG machine .</li>
	<li>After 4 min of right hilar clamping, aspirate arterial blood from the left ventricle into the heparin-coated syringe (see Figure 5I). Take care to avoid puncturing the ventricular septum and inadvertently aspirating venous right ventricular blood. There is a clear color difference between the darker right ventricle and brighter left ventricle (see Figure 5I inset). Angle the needle toward the left neck. Multiple punctures may be required to obtain sufficient blood to run an ABG (~150 &#181;L). 
	NOTE: During the 4 min of right hilar clamp, the mouse may start to exhibit agonal breathing which heralds impending death. If this occurs, arterial blood should be expeditiously aspirated prior to cardiac arrest. It is not possible to aspirate blood from a non-beating heart.</li>
	<li>Run the arterial blood on an ABG machine to obtain oxygen saturation, partial pressure of oxygen, partial pressure of carbon dioxide, among other measurements. Euthanize the mouse after ABG collection and/or antibody treatment (see step 6).</li>
</ol>
6. Intravenous antibody injection for measurement of cell extravasation
NOTE: This technique can be used to determine cell extravasation via injection of fluorochrome-labelled antibodies intravenously into the IVC prior to sacrifice followed by flow cytometric analysis, as previously published18. In brief, intravascular neutrophils can be distinguished from interstitial neutrophils using neutrophil specific anti-Ly6G antibodies. Fluorescein isothiocyanate-labelled (FITC-labeled) anti-Ly6G (clone 1A8) is injected intravenously 5 minutes prior to sacrifice, which labels the circulating intravascular neutrophils. The concentration of FITC-Ly6G antibody used is 100 ng diluted in 200 &#181;L of phosphate-buffered saline. Then, after preparation of single cell suspension from the left lung for flow cytometry, all neutrophils are labeled with allophycocyanin-labeled (APC-labeled) anti-Ly6G (clone 1A8). Thus, APC-Ly6G+FITC-Ly6G+ neutrophils are intravascular while APC-Ly6G+FITC-Ly6G- are extravascular or interstitial. This technique can be adapted to monocytes with anti-Ly6C antibodies, B cells with anti-CD19 antibodies, for example.
<ol>
	<li>Load the desired antibody into a 3/10 cc syringe with 5/16 inch 31G needle, taking care to minimize air bubbles within the syringe. Use this for intravenous injection into the IVC in step 6.5.</li>
	<li>Perform laparotomy according to steps 5.3-5.4.</li>
	<li>Following laparotomy, perform right medial visceral rotation bluntly using two pointed-cotton-tipped applicators. In a mouse, this is done by eviscerating all the bowel to the left of the abdomen, which will allow a clear view of the IVC (see Figure 6A).</li>
	<li>Clear the fat overlying the IVC bluntly with cotton-tipped applicators.</li>
	<li>Inject the antibody solution into the IVC via venipuncture (see Figure 6B). Upon extraction of the needle, immediately apply gentle pressure with a cotton swab to the site of venipuncture until it is hemostatic (usually about 2-3 min).</li>
	<li>Return the eviscerated bowel into the abdomen over the cotton swab to continue to apply pressure over the IVC (see Figure 6C).</li>
	<li>Perform clamshell thoracotomy according to steps 5.5-5.7 to harvest the left lung. Allow the antibodies to have at least 5 min to circulate systemically prior to sacrifice and harvest. Euthanize the mouse after ABG collection and/or antibody treatment.</li>
</ol>
7. Histology (H&#38;E) staining
<ol>
	<li>Following formalin-fixation and paraffin-embedding of the left lung and sectioning to a thickness of 5 &#181;m, deparaffinize the slide in xylene (two 10 min washes).</li>
	<li>Dehydrate in sequential ethanol washes: 2x 5&#160;min washes with 100%, 1x 2 min wash with 95%, and 1x 2 min wash with 70% ethanol. Then, rinse in deionized water.</li>
	<li>Stain the slide with hematoxylin for 2-4 min, then wash with deionized water for 5 min or until it is running clear. The precise duration of hematoxylin staining will need to be optimized to the tissue and desired nuclear staining intensity.</li>
	<li>Wash in defining solution for 30 s to differentiate stains, then wash in water for 2 min. Wash in blue color forming solution for 30 s, then wash in water for 2 min.</li>
	<li>Dehydrate further by dipping in 95% ethanol 15x. Stain in eosin for 1 min.</li>
	<li>Dehydrate with 2 min ethanol washes (2x), followed by 2 min xylene washes (2x). Apply mounting media and cover slip.</li>
</ol>

Mouse Model for Studying Lung Ischemia-Reperfusion Injury

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E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=NADPH+oxidase+mediates+synergistic+effects+of+IL-17+and+TNF-alpha+on+CXCL1+expression+by+epithelial+cells+after+lung+ischemia-reperfusion.">NADPH oxidase mediates synergistic effects of IL-17 and TNF-alpha on CXCL1 expression by epithelial cells after lung ischemia-reperfusion.</a> Am J Physiol Lung Cell Mol Physiol. 306 (1), L69-L79 (2014).</li><li>Zanotti, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Novel+critical+role+of+Toll-like+receptor+4+in+lung+ischemia-reperfusion+injury+and+edema.">Novel critical role of Toll-like receptor 4 in lung ischemia-reperfusion injury and edema.</a> Am J Physiol Lung Cell Mol Physiol. 297 (297), L52-L63 (2009).</li><li>National Research Council. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Guide for the care and use of laboratory animals. , (2011).</li><li>Saito, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pirfenidone+alleviates+lung+ischemia-reperfusion+injury+in+a+rat+model.">Pirfenidone alleviates lung ischemia-reperfusion injury in a rat model.</a> J Thorac Cardiovasc Surg. 158 (1), 289-296 (2019).</li><li>Tanaka, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Protective+effects+of+Imatinib+on+ischemia/reperfusion+injury+in+rat+lung.">Protective effects of Imatinib on ischemia/reperfusion injury in rat lung.</a> Ann Thorac Surg. 102 (5), 1717-1724 (2016).</li><li>Kreisel, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+two-photon+imaging+reveals+monocyte-dependent+neutrophil+extravasation+during+pulmonary+inflammation.">In vivo two-photon imaging reveals monocyte-dependent neutrophil extravasation during pulmonary inflammation.</a> Proc Natl Acad Sci U S A. 107 (42), 18073-18078 (2010).</li><li>Koletsis, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+situ+cooling+in+a+lung+hilar+clamping+model+of+ischemia-reperfusion+injury.">In situ cooling in a lung hilar clamping model of ischemia-reperfusion injury.</a> Exp Biol Med (Maywood). 231 (8), 1410-1420 (2006).</li><li>Hermsen, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genomic+landscape+of+rat+strain+and+substrain+variation.">Genomic landscape of rat strain and substrain variation.</a> BMC Genomics. 16 (1), 357 (2015).</li><li>Beck, J. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genealogies+of+mouse+inbred+strains.">Genealogies of mouse inbred strains.</a> Nat Genet. 24 (1), 23-25 (2000).</li><li>Liao, W. I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+mouse+model+of+orotracheal+intubation+and+ventilated+lung+ischemia+reperfusion+surgery.">A mouse model of orotracheal intubation and ventilated lung ischemia reperfusion surgery.</a> J Vis Exp. (187), 64383 (2022).</li><li>Murata, T., Nakazawa, H., Mori, I., Ohta, Y., Yamabayashi, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reperfusion+after+a+two-hour+period+of+pulmonary+artery+occlusion+causes+pulmonary+necrosis.">Reperfusion after a two-hour period of pulmonary artery occlusion causes pulmonary necrosis.</a> Am Rev Respir Dis. 146 (4), 1048-1053 (1992).</li><li>Ukita, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+large+animal+model+for+pulmonary+hypertension+and+right+ventricular+failure:+Left+pulmonary+artery+ligation+and+progressive+main+pulmonary+artery+banding+in+sheep.">A large animal model for pulmonary hypertension and right ventricular failure: Left pulmonary artery ligation and progressive main pulmonary artery banding in sheep.</a> J Vis Exp. (173), 62694 (2021).</li><li>Wang, Q., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Induction+of+right+ventricular+failure+by+pulmonary+artery+constriction+and+evaluation+of+right+ventricular+function+in+mice.">Induction of right ventricular failure by pulmonary artery constriction and evaluation of right ventricular function in mice.</a> J Vis Exp. (147), 59431 (2019).</li><li>Welbourn, C. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pathophysiology+of+ischaemia+reperfusion+injury:+central+role+of+the+neutrophil.">Pathophysiology of ischaemia reperfusion injury: central role of the neutrophil.</a> Br J Surg. 78 (6), 651-655 (1991).</li></ol>

The authors report no relevant disclosures.

We describe a hilar clamp technique that involves application of a slipknot on the left hilum which occludes the pulmonary artery and veins and bronchus to induce warm ischemia followed by reperfusion. After hilar clamping, the left lung can be harvested for a variety of experimental techniques such as histology, flow cytometry, bulk or single cell sequencing, and quantitative polymerase chain reaction. Additionally, the blood and spleen may be used to study systemic effects, while the non-ischemic right lung may serve as an internal control. In this protocol, we also describe detailed methods to obtain ABG measurements and/or determine cell extravasation.
The murine hilar clamping technique is critical to the study of lung IRI and to the wider field of lung transplantation. For example, using this hilar clamp model, our group found that spleen derived CCR2+ classical monocytes mediate neutrophil extravasation into the lung following IRI18. Another group showed that type II alveolar epithelial cells produce CXCL1 (a neutrophil chemoattractant) in a NADPH oxidase-IL-17-TNF&#945;-dependent manner using a hilar clamp model19. TLR4 was found to play a role in the development of edema in a murine model of hilar clamping20.
While orthotopic murine lung transplantation is more clinically applicable and can simulate both warm and cold ischemia, it also introduces surgical trauma from vascular and bronchial anastomoses and allogeneic stimuli from donor-recipient mismatch. Thus, the main advantage of the hilar clamp technique is its ability to isolate the effects of lung IRI from these other factors. Additionally, for the inexperienced mouse microsurgeon, the hilar clamp procedure&#160;is technically less demanding and can be more quickly mastered compared to lung transplantation. One limitation of the hilar clamp model is that cold ischemia is difficult to achieve. In situ cold ischemia has been performed in pigs with antegrade flush perfusion via the left pulmonary artery25, however this is much more challenging in the mouse due to its smaller size and the need to maintain normothermia during surgery. Nevertheless, the study of isolated warm ischemia is important as it is generally more deleterious for organ function than cold ischemia4,5,6. Furthermore, transplant centers are increasingly adopting DCD lung transplants, which are accompanied by much longer warm ischemic times than traditional DBD lung transplants7. Thus, detailed study of the events during warm ischemia and the subsequent reperfusion period is critical and can be accomplished with this hilar clamp model.
Many variations of the hilar clamp technique have been created over the last couple decades. These include the use of an atraumatic microvascular clamp8,9,10,11,12,13, a Rumel tourniquet14,15, and a tie with a slipknot around the hilum16. We describe our adaptation of the slipknot technique in mice. The main advantage this technique has over other hilar clamp techniques is that it allows for closure of the chest during the period of ischemia, which minimizes insensible fluid losses particularly in protocols with longer periods of ischemia. This cannot be accomplished when a clamp or tourniquet is placed on the hilum.
Regarding animal selection, we chose to develop this model in mouse as opposed to rat or large animal models, such as pig. The reasoning is two-fold - our laboratory has extensive experience in mouse microsurgery with orthotopic mouse lung transplantation17. Secondly, while mice are smaller and more technically challenging to operate on than rats, there are presently more commercially available transgenic mouse strains than rat strains26,27. Mice also breed more quickly, allowing for faster development of new strains, which is particularly useful when investigating immune mechanisms of IRI. For these reasons, we have chosen to develop this technique in mice.
To our knowledge, this is the first video-based article using the technique of left lung hilar clamp in mouse with a slipknot. Saito et al have published a video-based article demonstrating hilar clamp in rat using a microvascular clamp22. In mouse, Liao et al published a lung IRI model with pulmonary artery (PA) clamping with a slipknot, however the lung remains ventilated in this model as the bronchus is excluded from the clamp28. The PA clamping technique has been used as a model to study pulmonary embolism29, pulmonary hypertension and/or right ventricular failure30,31. Since the area of study is lung IRI in transplantation, we elected to clamp the entire hilum including the bronchus to minimize barotrauma on top of the ischemic injury.
Of note, 30 min of warm ischemia was chosen followed by 1 h of reperfusion because we are interested in the dynamics of lung-infiltrating cells in the very early period post-IRI. For example, studies have reported upregulation of cytoprotective genes, such as heme oxygenase 1, after just 30 min of ischemia followed by 2 h of reperfusion13. Neutrophils were previously thought to be the central cell type infiltrating the lung after IRI32, however we showed that monocytes precede neutrophils in their arrival to the lung18,24. More recently, we found that B cells precede even monocytes and enter the lung within 1 h of reperfusion (unpublished data). When adapting this technique, the durations of warm ischemia and reperfusion can be adjusted according to the study question and hypothesis. Longer periods of warm ischemia are possible without concern for excessive fluid and heat loss since the chest can be closed during this period.
Several complications can occur perioperatively, with the final common outcome being perioperative demise. We have observed that the incidence of animal death during surgery decreases with repetitions of the technique by the surgeon. We estimate that surgeons require at least 2 months of practice and/or at least 20 cases to reach a plateau in skill. At this point, we estimate that intraoperative mortality occurs in approximately 1 in 50 cases. Death in the anesthetized mouse is often heralded by agonal breaths that involve forceful contractions of the diaphragm and chest wall musculature. Upon the onset of these agonal breaths, death will occur in 3-5 min if no intervention is taken. There are several causes that can precipitate intraoperative demise, many of which are recognizable and preventable. We will highlight below the critical steps of this surgery and troubleshooting techniques to achieve a successful outcome.
Intubation 
The first critical step is careful intubation and securing of the airway to prevent inadvertent extubation. In a survival surgery, it is particularly important to ensure that the vocal cords are not damaged during initial intubation, as the mouse will not be able to breathe following extubation. Advancement of the ETT past the cords should be done under direct visualization through the microscope. If resistance is met during intubation, the ETT should not be advanced blindly. While keeping the oropharynx open with the mosquito clamp, care should be taken to not pierce the oral mucosa as blood will obscure view of the airway. The number of attempts at intubation should be limited to less than five.
Accidental extubation 
If at any point during the procedure, the lungs do not appear to be inflating appropriately, it is possible that the ETT has dislodged. Another manifestation of accidental extubation that goes unnoticed for a prolonged period is the mouse suddenly developing agonal breathing. Following initial intubation, care should be taken to secure the ETT well to the nose circumferentially with silk tape, and adjustment of the mouse&#39;s neck and torso should be limited. If neither the left nor right lung is inflating, the ETT may have come out of the trachea. Attempts can be made to push the ETT back in, however the mouse may need to be re-intubated in an expeditious manner to avoid premature death of the animal. If only one lung is inflating when both are expected to, the ETT may be too deep and should be carefully drawn back and re-secured.
Lung mobilization 
Left lung mobilization should be done using cotton-tipped applicators with gentle pressure on the lung parenchyma, as compression injury may damage alveoli and decrease compliance of the lung. The lungs should not be grasped with forceps or clamps as this will damage the lung parenchyma.
Application of hilar clamp 
The left hilar clamp should be placed under direct visualization, taking care not to lay the knot down too centrally (as to not catch the left atrium) or too peripherally (as to not catch any lung parenchyma). Catching the left atrium with the hilar clamp will likely precipitate atrial fibrillation and subsequent&#160;death. Visualization of the hilum can be improved by ensuring adequate rib retraction to expand the field of view. After tying down the slipknot, it is useful to ensure that the clamp is completely occlusive by performing a recruitment maneuver - by manually occluding the outflow tubing connecting the ETT to the ventilator and ensuring that the left lung does not expand with positive pressure. Vascular occlusion is presumed with confirmation of bronchial occlusion, given the increased collapsibility of the pulmonary artery and veins compared to the cartilaginous bronchus. Additionally, the hilar clamp should not be tied with excessive force as it will crush the cartilaginous portions of the left mainstem bronchus and prevent expansion of the left lung following release of the hilar clamp. Once the hilar clamp is released, it is important to check that the lung is still able to be ventilated by performing another recruitment maneuver.
Chest closure 
When closing the chest, the 4th and 5th ribs should be brought together when the lung is at maximum end-inspiration. This will minimize the amount of air remaining in the pleural cavity, which would result in pneumothorax once the ribs are closed. The muscle layers and skin should be closed in an airtight manner to avoid development of tension pneumothorax.
Obtaining ABGs 
Finally, when obtaining ABGs, it is important to first clamp the right hilum so that the ABG measurement will be reflective of only left lung function22,23. Without the right hilar clamp, the normally ventilating right lung will more than compensate for the damaged ischemic left lung and ABG measurements will appear normal.
In summary, the hilar clamp technique that we describe above is a valuable tool for the study of lung transplantation, and importantly can isolate the study of warm IRI without the introduction of allogenicity, surgical anastomosis-related trauma, or cold ischemic storage. Notably, this described technique of hilar clamping is easily taught and widely adoptable and can lead to reliable and reproducible results.

<a href="https://app.jove.com/t/6603">This corrects</a> the article <a href="https://dx.doi.org/10.3791/66232">10.3791/66232</a>

An erratum was issued for:&#160;<a href="https://www.jove.com/t/66232">Murine Left Pulmonary Hilar Clamp Model of Lung Ischemia Reperfusion Injury</a>. The Protocol and Representative Results&#160;sections were&#160;updated.

Erratum: Murine Left Pulmonary Hilar Clamp Model of Lung Ischemia Reperfusion Injury

IRI during organ transplantation is a major risk factor for primary graft dysfunction and later episodes of graft rejection1,2. During transplantation, warm ischemia time is defined as the period of time from donor aortic cross-clamp to initiation of cold perfusion and from organ removal from ice to organ implantation. Cold storage time is defined as the period of time from the start of cold perfusion to the removal of the organ from ice3. Warm ischemia is more deleterious for later organ function than cold ischemia4,5,6, and its underlying mechanisms warrant further study in pre-clinical models. Additionally, organ transplantation from donation after cardiac death (DCD) is associated with longer warm ischemic times than traditional donation after brain death (DBD)7. While the use of DCD donors can expand the donor pool and increase lung utilization, further pre-clinical studies to evaluate the effects of warm ischemia on post-transplant lung function are needed. Below, we describe a model of warm IRI in mice via the left pulmonary hilar clamp.
Several animal models of lung hilar clamping have been developed and adapted over the last several years and may include the use of an atraumatic microvascular clamp8,9,10,11,12,13, a Rumel tourniquet14,15, or suture ligation16 as the hilar clamp. The crux of the hilar clamp is that it must be reversible and cause minimal or no damage to the hilar structures so that reperfusion can be achieved. Here, we describe our hilar clamp technique in mice that involves reversible suture ligation of the left pulmonary hilum with a slipknot. This method occludes the pulmonary arterial inflow, venous outflow, and airflow in and out of the mainstem bronchus. The main benefit of a slipknot over a vascular clamp, clip, or tourniquet is that the chest can be closed during prolonged periods of ischemia, thereby minimizing insensible fluid and heat loss in the mouse. We provide a protocol for obtaining reliable arterial blood gas (ABG) measurements and for measuring cell extravasation after hilar clamping.
This hilar clamp technique holds an important place in the wider study of lung transplantation. Compared to small animal models of orthotopic lung transplantation, the hilar clamp technique can isolate the effects of IRI without the addition of surgical anastomotic trauma or allogenicity17. Additionally, the hilar clamp technique can be more easily and quickly mastered than the mouse lung transplant. In fact, using hilar clamp techniques, several important mechanisms in the pathogenesis of IRI have been identified in the past decade, such as TLR4, NADPH oxidase, and adenosine A2A receptor14,18,19,20. In the following protocol, we present a reliable, teachable, and reproducible method of hilar clamping as a tool for studying lung IRI.

<table><tbody><tr><td>&lt;strong&gt;Medications&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>10% povidone-iodine solution</td><td>Aplicare</td><td>NDC 52380-0126-2</td><td>For disinfectant</td></tr><tr><td>Buprenorphine 1.3 mg/mL</td><td>Fidelis Animal Health</td><td>NDC 86084-100-30</td><td>For pain control</td></tr><tr><td>Carprofen</td><td>Cronus Pharma</td><td>NDC 69043-027-18</td><td>For pain control</td></tr><tr><td>Heparin 1000 units/mL</td><td>Sagent</td><td>NDC 25021-404-01</td><td>For obtaining arterial blood</td></tr><tr><td>Isoflurane 1%-1.5%</td><td>Sigma Aldrich</td><td>26675-46-7</td><td>For anesthesia</td></tr><tr><td>Ketamine hydrochloride 100 mg/mL</td><td>Vedco</td><td>NDC 50989-996-06</td><td>For anesthesia</td></tr><tr><td>Puralube Vet eye ointment</td><td>Medi-Vet.com</td><td>11897</td><td>To prevent eye dessiccation</td></tr><tr><td>Xylazine 20 mg/mL</td><td>Akorn</td><td>NDC 59399-110-20</td><td>For pain control</td></tr><tr><td>&lt;strong&gt;Tools and Instruments&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Argent High Temp Fine Tip Cautery Pen</td><td>McKesson</td><td>231</td><td>To coagulate blood vessels</td></tr><tr><td>Curved mosquito clamp</td><td>Fine Science Tools</td><td>13009-12</td><td>For surgical procedure</td></tr><tr><td>Fine curved forceps</td><td>Fine Science Tools</td><td>11274-20</td><td>For surgical procedure</td></tr><tr><td>Fine scissors</td><td>Fine Science Tools</td><td>15040-11</td><td>For surgical procedure</td></tr><tr><td>Intubation clamp set-up</td><td>Fine Science Tools</td><td>18374-44, 18144-30</td><td>For holding mouse vertically by the tongue during intubation. See Supplementary Figure 1A.&amp;nbsp;</td></tr><tr><td>Magnetic rib retractors</td><td>Fine Science Tools</td><td>18200-01, 18200-10</td><td>For retraction of thoracotomy. Magnetic fixator and retractor should be connected by micro latex tubing below.</td></tr><tr><td>Optical Grade Plastic Optical Fiber Unjacketed, 500&amp;mu;m</td><td>Edmund Optics</td><td>02-532</td><td>To make the introducer for the endotracheal tube. See Supplemental Figure 1B. A 1.5-inch length of this optical fiber should have a piece of silk tape secured to one end. It can then be used as an introducer for the endotracheal tube. The end of the introducer should be curves slightly.</td></tr><tr><td>Power Pro Ultra clipper</td><td>Oster</td><td>078400-020-001</td><td>To clip hair</td></tr><tr><td>Scissors</td><td>Fine Science Tools</td><td>14370-22</td><td>For surgical procedure</td></tr><tr><td>Small animal heating pad</td><td>K&amp;amp;H Pet Products</td><td>Thermo-Peep Heated Pad</td><td>To maintain normothermia</td></tr><tr><td>Small animal ventilator</td><td>Harvard Apparatus</td><td>55-0000</td><td>For ventilation (TV 0.35 cc, PEEP 1 cm H2O, RR 100-105/min, FiO2 100%)</td></tr><tr><td>Spearit Micro Latex Rubber Tubing (1/8 in outside diameter, 1/16 in inside diameter)</td><td>Amazon.com</td><td>https://www.amazon.com/Rubber-Tubing-CONTINUOUS-Select-Length/dp/B00H4MT7V0?th=1</td><td>For retraction of thoracotomy</td></tr><tr><td>Stat Profile Prime Critical Care Blood Gas Analyzer</td><td>Nova Biomedical</td><td>https://novabiomedical.com/prime-plus-critical-care-blood-gas-analyzer/index.php?gad=1&amp;amp;gclid=Cj0KCQjwmICoBhDx&lt;br/&gt; ARIsABXkXlInZX--R3ezBkc304nS_GVGI9Z2T3Esr33&lt;br/&gt; 2aM8WGPiUVhicPQZ&lt;br/&gt; Wj2AaAqhDEALw_wcB&amp;nbsp;&amp;nbsp;</td><td>For retraction of thoracotomy</td></tr><tr><td>Straight clamp</td><td>Fine Science Tools</td><td>13008-12</td><td>For surgical procedure</td></tr><tr><td>Straight forceps</td><td>Fine Science Tools</td><td>91113-10</td><td>For surgical procedure</td></tr><tr><td>Surgical microscope</td><td>Wild Heerbrugg</td><td>no longer produced</td><td>For intubation and surgical procedure; recommend replacement with Leica surgical microscopes</td></tr><tr><td>&lt;strong&gt;Supplies&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>&amp;frac12; cc syringe with &amp;frac12; inch 29G needle</td><td>McKesson</td><td>942665</td><td>For injecting ketamine/xylazine intraperitoneally</td></tr><tr><td>&amp;frac12; inch 31G needle on a 1 cc tuberculin syringe</td><td>McKesson</td><td>16-SNT1C2705</td><td>For aspiration of arterial blood from left ventricle</td></tr><tr><td>1-inch 20G IV catheter</td><td>Terumo</td><td>SROX2025CA</td><td>For endotracheal tube (ETT)</td></tr><tr><td>1-inch silk tape</td><td>Durapore</td><td>3M ID 7100057168</td><td>To tape ETT to nose and to secure limbs</td></tr><tr><td>3/10 cc syringe with 5/16 inch 31G needle</td><td>McKesson</td><td>102-SN310C31516P</td><td>For antibody injection into the inferior vena cava</td></tr><tr><td>6-0 monofilament suture on a P-10 needle</td><td>McKesson</td><td>S697GX</td><td>For closure of thoracotomy, muscle layer, and skin</td></tr><tr><td>6-0 silk tie</td><td>Surgical Specialties Look</td><td>SP102</td><td>To make slipknot for hilar clamp</td></tr><tr><td>Pointed cotton-tipped applicators</td><td>Solon</td><td>56225</td><td>To manipulate lung and for blunt dissection</td></tr></tbody></table>

murine left pulmonary hilar clamp model of lung ischemia reperfusion injury

Ischemia reperfusion injury (IRI) during lung transplantation is a major risk factor for post-transplant complications, including primary graft dysfunction, acute and chronic rejection, and mortality. Efforts to study the underpinnings of IRI led to the development of a reliable and reproducible mouse model of left lung hilar clamping. This model involves a surgical procedure performed in an anesthetized and intubated mouse. A left thoracotomy is performed, followed by careful lung mobilization and dissection of the left pulmonary hilum. The hilar clamp involves reversible suture ligation of the pulmonary hilum with a slipknot, which stops the arterial inflow, venous outflow, and airflow through the left mainstem bronchus. Reperfusion is initiated by careful removal of the suture. Our laboratory uses 30 min of ischemia and 1 h of reperfusion for the experimental model in the current investigations. However, these time periods can be modified depending on the specific experimental question. Immediately prior to sacrifice, arterial blood gas can be obtained from the left ventricle after a 4 min period of right hilar clamping to ensure that the PaO2 values obtained are attributed to the injured left lung alone. We also describe a method to measure cell extravasation with flow cytometry, which involves intravenous injection of a fluorochrome-labeled antibody specific for the cell(s) to be studied prior to sacrifice. The left lung can then be harvested for flow cytometry, frozen or fixed, paraffin-embedded immunohistochemistry, and quantitative polymerase chain reaction. This hilar clamp technique allows for&#160;detailed study of the cellular and molecular mechanisms underlying IRI. Representative results reveal decreased left lung oxygenation and histologic evidence of lung injury following hilar clamping. This technique can be readily learned and reproduced by personnel with and without microsurgical experience, leading to reliable and consistent results and serving as a widely adoptable model for studying lung IRI.

After left hilar clamping, partial pressure of oxygenation in the arterial blood (PaO2) attributed to the left lung is ~100 mmHg, significantly lower compared to the ~500 mmHg following sham thoracotomy (Figure 7A, n=6-7). Of note, sham thoracotomies were performed in B6 mice with ABG measurement taken after 4 min of right hilar clamping, representing values attributed to the left lung alone. H&#38;E staining of the hilar clamped left lung demonstrated infiltrating inflammatory cells filling the airways and engorgement of blood vessels with erythrocytes, while these findings are absent in the na&#239;ve left lung (Figure 7B, n=2). These results demonstrate the immediate tissue injury and cell infiltration following a period of ischemia and reperfusion.
As discussed in step 6, intravascular and interstitial neutrophils can be differentiated with intravenous injection of FITC-labeled anti-Ly6G antibody 5 min prior to sacrifice, followed by labeling of all neutrophils with APC-Ly6G antibodies18,24. Representative plots show that ~75% of neutrophils are intravascular (APC-Ly6G+FITC-Ly6G+) and ~25% are interstitial (APC-Ly6G+FITC-Ly6G-) after hilar clamping of the left lung (Figure 7C, n=4). These results demonstrate a reliable way to assess neutrophil extravasation and can be adapted to other cell types depending on the antibodies used (i.e., anti-Ly6C for monocytes, anti-CD19 for B cells). This technique is useful for the study of cell dynamics following IRI.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66232/66232fig1v2.jpg" /> 
Figure 1: Mouse intubation. (A) Mouse with left chest and abdomen shaved. (B) Endotracheal tube connected to ventilator tubing. (C) Once intubation is confirmed, tape the tube circumferentially to the nose of the mouse. (D) Intubated mouse with inflow and outflow tubing from and to the small animal ventilator. <a href="https://www.jove.com/files/ftp_upload/66232/66232fig1v2large.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/66232/66232fig02.jpg" /> 
Figure 2: Left thoracotomy. (A) Skin incision over 4th intercostal space (ICS). (B)&#160;Muscle layers divided over 4th intercostal space. (C)&#160;Making small incision just above 5th rib. (D)&#160;Extending incision anteriorly and posteriorly in 4th ICS. (E) Lung visualized in 4th ICS. (F-G)&#160;Rib retractors placed. <a href="https://www.jove.com/files/ftp_upload/66232/66232fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66232/66232fig03.jpg" /> 
Figure 3: Left hilar clamp. (A) Left lung mobilized with cotton swabs. (B) Inferior pulmonary ligament (IPL) visualized. (C)&#160;With left lung flipped anteriorly, silk tie is placed posterior to the hilum. (D)&#160;With left flipped posteriorly over silk tie, anterior hilum is visualized. (E)&#160;Instrument tie of slipknot over left hilum. (F)&#160;Slipknot on hilum. (G)&#160;With manual occlusion of outflow, left lung does not inflate, indicating successful hilar clamp. <a href="https://www.jove.com/files/ftp_upload/66232/66232fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/66232/66232fig04.jpg" /> 
Figure 4: Release of left hilar clamp and left chest closure. (A) With slipknot released, the left lung is reinflated. (B)&#160;Rib closure with one bite just above 4th rib and (C)&#160;one bite just above 6th rib. (D-E)&#160;Chest closed with one stitch tied down at end-inspiration. (F)&#160;Skin closure with interrupted stitches (after muscle layer closure with running stitch). <a href="https://www.jove.com/files/ftp_upload/66232/66232fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/66232/66232fig05.jpg" /> 
Figure 5: Laparotomy, bilateral clamshell thoracotomy, and arterial blood collection. (A) Mouse adjusted to supine position. (B)&#160;Skin incision made over midline abdomen. (C)&#160;Laparotomy extending from xiphoid to pubis, with extension left and right along the most inferior rib. (D-E)&#160;Opening of the diaphragm to enter the chest cavity, with extension of diaphragm incision left and right along most inferior rib (dotted line). (F)&#160;Bilateral clamshell thoracotomy with incisions in anterior axillary lines (dotted line) toward apex of chest. (G)&#160;Accessory lobe of right lung extending into the left chest posterior to the inferior vena cava. (H)&#160;Accessory lobe mobilized and returned to the right chest. (I) Aspiration of arterial blood from left ventricle (inset: color differential between right and left ventricles). <a href="https://www.jove.com/files/ftp_upload/66232/66232fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/66232/66232fig06.jpg" /> 
Figure 6: Intravenous injection into the inferior vena cava. (A) Right medial visceral rotation to expose inferior vena cava (IVC). (B)&#160;Injection of solution into IVC. (C)&#160;Applying pressure over IVC to achieve hemostasis. <a href="https://www.jove.com/files/ftp_upload/66232/66232fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/66232/66232fig07.jpg" /> 
Figure 7: Representative results of lung oxygenation, histology, and neutrophil extravasation following left hilar clamp. (A) Partial pressure of oxygen in the arterial blood of a C57BL/6 mouse that underwent sham thoracotomy vs. hilar clamp. Results are presented as mean &#177; SEM, n=6-7. p values calculated by Mann-Whitney U test. **p &#60;0.01. (B) H&#38;E staining of formalin-fixed, paraffin-embedded lung section of a C57BL/6 mouse that underwent sham thoracotomy vs. hilar clamp. The scale bar represents 250 &#181;m. (C) Representative figure of distribution of intravascular (APC-Ly6G+FITC-Ly6G+) vs. extravascular (APC-Ly6G+FITC-Ly6G-) Ly6G+ neutrophils in a C57BL/6 mouse following hilar clamp. <a href="https://www.jove.com/files/ftp_upload/66232/66232fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary Figure 1:&#160;Introducer for endotracheal tube. <a href="https://www.jove.com/files/ftp_upload/66232/Supplemental_Figure_1_updated.pdf" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Author Spotlight: Insights into the Effect of Ischemia Reperfusion on Lung Transplantation at JoVE.com

Author Spotlight: Insights into the Effect of Ischemia Reperfusion on Lung Transplantation

The protocol outlines a feasible, reliable, and reproducible method of left pulmonary hilar clamping that can be used to study lung ischemia-reperfusion injury in mouse models.

Murine Left Pulmonary Hilar Clamp Model of Lung Ischemia Reperfusion Injury

Ischemia reperfusion injury (IRI) during lung transplantation is a major risk factor for post-transplant complications, including primary graft ...

murine-left-pulmonary-hilar-clamp-model-lung-ischemia-reperfusion

Nanyang Technological University

Research

JoVE Journal

Medicine

1.3K Views.  Washington University, St. Louis. The protocol outlines a feasible, reliable, and reproducible method of left pulmonary hilar clamping that can be used to study lung ischemia-reperfusion injury in mouse models.

Video: Author Spotlight: Insights into the Effect of Ischemia Reperfusion on Lung Transplantation

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 ...

Murine Model of Metastatic Liver Tumors in the Setting of Ischemia Reperfusion Injury

Liver ischemia and reperfusion (I/R) injury, a common clinical challenge, remains an inevitable pathophysiological process that has been shown to induce multiple tissue and organ damage. Despite recent advances and therapeutic approaches, the overall morbidity has remained unsatisfactory especially in patients with underlying parenchymal abnormalities. In the context of aggressive cancer growth and metastasis, surgical I/R is suspected to be the promoter regulating tumor recurrence. This article aims to describe a clinically relevant murine model of liver I/R and colorectal liver metastasis. In doing so, we aim to assist other investigators in establishing and perfecting this model for their routine research practice to better understand the effects of liver I/R on promoting liver metastases.

We describe in detail a clinically relevant colorectal cancer liver metastases (CRLM) tumor model and the influence of liver ischemia reperfusion (I/R) in tumor growth and metastasis. This model can help to better understand the mechanisms underlying surgery-induced promotion of liver metastatic growth.

Liver ischemia and reperfusion (I/R) injury, a common clinical challenge, remains an inevitable pathophysiological process that has been shown to induce ...

Cancer Research

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 ...

Method of Direct Segmental Intra-hepatic Delivery Using a Rat Liver Hilar Clamp Model

Major hepatic surgery with inflow occlusion, and liver transplantation, necessitate a period of warm ischemia, and a period of reperfusion leading to ischemia/reperfusion (I/R) injury with myriad negative consequences. Potential I/R injury in marginal organs destined for liver transplantation contributes to the current donor shortage secondary to a decreased organ utilization rate. A significant need exists to explore hepatic I/R injury in order to mediate its impact on graft function in transplantation. Rat liver hilar clamp models are used to investigate the impact of different molecules on hepatic I/R injury. Depending on the model, these molecules have been delivered using inhalation, epidural infusion, intraperitoneal injection, intravenous administration or injection into the peripheral superior mesenteric vein. A rat liver hilar clamp model has been developed for use in studying the impact of pharmacologic molecules in ameliorating I/R injury. The described model for rat liver hilar clamp includes direct cannulation of the portal supply to the ischemic hepatic segment via a side branch of the portal vein, allowing for direct segmental hepatic delivery. Our approach is to induce ischemia in the left lateral and median lobes for 60 min, during which time the substance under study is infused. In this case, pegylated-superoxide dismutase (PEG-SOD), a free radical scavenger, is infused directly into the ischemic segment. This series of experiments demonstrates that infusion of PEG-SOD is protective against hepatic I/R injury. Advantages of this approach include direct injection of the molecule into the ischemic segment with consequent decrease in volume of distribution and reduction in systemic side effects.

A unique rat liver hilar clamp model was developed for studying the impact of pharmacologic molecules in ameliorating ischemia-reperfusion injury. This model includes direct cannulation of the portal supply to the ischemic liver segment via a branch of the portal vein, allowing for direct hepatic delivery.

Major hepatic surgery with inflow occlusion, and liver transplantation, necessitate a period of warm ischemia, and a period of reperfusion leading to ...

Murine Left Anterior Descending (LAD) Coronary Artery Ligation: An Improved and Simplified Model for Myocardial Infarction

Ischemic heart disease (IHD), or acute coronary syndrome (ACS), is one of the leading causes of death in the United States. IHD is characterized by reduced blood supply to the heart, resulting in the loss of oxygen to and the ensuing necrosis of the heart muscle. The MI model has gained popularity for its use as a short-term ischemia-reperfusion model and a long-term permanent ligation model. Below, we describe a reliable method for the permanent ligation of the LAD. With mouse genetic engineering technology becoming more advanced, and with an increasing availability of quality murine surgical instruments, the mouse has become a popular model for MI surgeries. Our surgical model incorporates the use of an easily reversible anesthetic for the rapid recovery of the mouse; a minimally invasive endotracheal intubation without involving a tracheotomy; and a thoracentesis through the original thoracotomy site without creating an additional incision in the chest, as is done in some other methods, to effectively remove excess blood and air from the chest cavity. This method is comparatively less invasive than other methods, which dramatically reduces surgical and post-surgical complications and mortality and improves reproducibility.

Murine Left Anterior Descending LAD Coronary Artery Ligation: An Improved and Simplified Model for Myocardial Infarction

We provide a reliable method for left anterior descending artery (LAD) ligation in a mouse model. This method is comparatively less invasive than other methods, involving endotracheal intubation, a left-sided thoracotomy approach, and thoracentesis. This method can be used as a model for both acute and chronic myocardial infarction (MI).

Ischemic heart disease (IHD), or acute coronary syndrome (ACS), is one of the leading causes of death in the United States. IHD is characterized by ...

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 ...

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

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

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

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

Immunology and Infection

A Rat Lung Transplantation Model of Warm Ischemia/Reperfusion Injury: Optimizations to Improve Outcomes

From our experience with rat lung transplantation, we have found several areas for improvement. Information in the existing literature regarding methods for choosing appropriate cuff sizes for the pulmonary vein (PV), pulmonary artery (PA), or bronchus (Br) are varied, thus making the determination of proper cuff size during rat lung transplantation an exercise of trial and error. By standardizing the cuffing technique to use the smallest effective cuff appropriate for the size of the vessel or bronchus, one can make the transplantation procedure safer, faster, and more successful. Since diameters of the PV, PA, and Br are related to the body weight of the rat, we present a strategy to choosing an appropriate size using a weight-based guide. Since lung volume is also related to body weight, we recommend that this relationship should also be considered when choosing the proper volume of air for donor lung inflation during warm ischemia as well as for the proper volume of PBS to be instilled during bronchoalveolar lavage (BAL) fluid collection. We also describe methods for 4th intercostal space dissection, wound closure, and sample collection from both the native and transplanted lobes.

Here, we present optimizations to a rat lung transplantation model that serve to improve outcomes. We provide a size guide for cuffs based on body weight, a measurement strategy to ascertain the 4th intercostal space, and methods of wound closure and BAL (bronchoalveolar lavage) fluid and tissue collection.

From our experience with rat lung transplantation, we have found several areas for improvement. Information in the existing literature regarding methods ...

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 ...

Left Coronary Artery Ligation: A Surgical Murine Model of Myocardial Infarction

Ischemic heart disease and subsequent myocardial infarction (MI) is one of the leading causes of mortality in the United States and around the world. In order to explore the pathophysiological changes after myocardial infarction and design future treatments, research models of MI are required. Permanent ligation of the left coronary artery (LCA) in mice is a popular model to investigate cardiac function and ventricular remodeling post MI. Here we describe a less invasive, reliable, and reproducible surgical murine MI model by permanent ligation of the LCA. Our surgical model comprises of an easily reversible general anesthesia, endotracheal intubation that does not require a tracheotomy, and a thoracotomy. Electrocardiography and troponin measurement should be performed to ensure MI. Echocardiography at day 28 after MI will discern heart function and heart failure parameters. The degree of cardiac fibrosis can be evaluated by Masson's trichrome staining and cardiac MRI. This MI model is useful for studying the pathophysiological and immunological alterations after MI.

Presented here is a surgical procedure for permanent ligation of the left coronary artery in mice. This model can be used to investigate the pathophysiology and associated inflammatory response after myocardial infarction.

Ischemic heart disease and subsequent myocardial infarction (MI) is one of the leading causes of mortality in the United States and around the world. In ...