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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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=Neutrophil+extracellular+traps+are+pathogenic+in+primary+graft+dysfunction+after+lung+transplantation.">Neutrophil extracellular traps are pathogenic in primary graft dysfunction after lung transplantation.</a> Am J Respir Crit Care Med. 191 (4), 455-463 (2015).</li><li>Okazaki, 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=A+mouse+model+of+orthotopic+vascularized+aerated+lung+transplantation.">A mouse model of orthotopic vascularized aerated lung transplantation.</a> Am J Transplant. 6 (7), 1672-1679 (2007).</li><li>Hsiao, H. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spleen-derived+classical+monocytes+mediate+lung+ischemia-reperfusion+injury+through+IL-1beta.">Spleen-derived classical monocytes mediate lung ischemia-reperfusion injury through IL-1beta.</a> J Clin Invest. 128 (7), 2833-2847 (2018).</li><li>Sharma, A. K., Mulloy, D. P., Le, L. T., Laubach, V. 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 a...

<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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Medications</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>Tools and Instruments</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.&#160;</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&#956;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&#38;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&#38;gclid=Cj0KCQjwmICoBhDx 
			ARIsABXkXlInZX--R3ezBkc304nS_GVGI9Z2T3Esr33 
			2aM8WGPiUVhicPQZ 
			Wj2AaAqhDEALw_wcB&#160;&#160;</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>Supplies</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>&#189; cc syringe with &#189; inch 29G needle</td>
			<td>McKesson</td>
			<td>942665</td>
			<td>For injecting ketamine/xylazine intraperitoneally</td>
		</tr>
		<tr>
			<td>&#189; 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 ce...

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

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1.2K 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

Normothermic Cardiac Arrest and Cardiopulmonary Resuscitation: A Mouse Model of Ischemia-Reperfusion Injury

Acute Kidney Injury (AKI) is a common, highly lethal, complication of critical illness which has a high mortality1-4 and which is most 
frequently caused by whole-body hypoperfusion.5,6 Successful reproduction of whole-body hypoperfusion in rodent models has been fraught with 
difficulty.7-9,9,10 Models which employ focal ischemia have repeatedly demonstrated results which do not translate to the clinical setting, and larger 
animal models which allow for whole body hypoperfusion lack access to the full toolset of genetic manipulation possible in the mouse.11,12 However, in recent 
years a mouse model of cardiac arrest and cardiopulmonary resuscitation has emerged which can be adapted to model AKI.13 This model reliably reproduces 
physiologic, functional, anatomic, and histologic outcomes seen in clinical AKI, is rapidly repeatable, and offers all of the significant advantages of a murine surgical 
model, including access to genetic manipulative techniques, low cost relative to large animals, and ease of use. Our group has developed extensive experience 
with use of this model to assess a number of organ-specific outcomes in AKI.14,15

A powerful model for perioperative and critical care related acute kidney injury is presented. Using whole body hypoperfusion induced by cardiac arrest it is possible to nearly replicate the histologic and functional changes of clinical AKI.

Acute Kidney Injury (AKI) is a common, highly lethal, complication of critical illness which has a high mortality1-4 and which is most 
frequently caused ...

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 ...

Ischemia-reperfusion Model of Acute Kidney Injury and Post Injury Fibrosis in Mice

Ischemia-reperfusion induced acute kidney injury (IR-AKI) is widely used as a model of AKI in mice, but results are often quite variable with high, often unreported mortality rates that may confound analyses. Bilateral renal pedicle clamping is commonly used to induce IR-AKI, but differences between effective clamp pressures and/or renal responses to ischemia between kidneys often lead to more variable results. In addition, shorter clamp times are known to induce more variable tubular injury, and while mice undergoing bilateral injury with longer clamp times develop more consistent tubular injury, they often die within the first 3 days after injury due to severe renal insufficiency. To improve post-injury survival and obtain more consistent and predictable results, we have developed two models of unilateral ischemia-reperfusion injury followed by contralateral nephrectomy. Both surgeries are performed using a dorsal approach, reducing surgical stress resulting from ventral laparotomy, commonly used for mouse IR-AKI surgeries. For induction of moderate injury BALB/c mice undergo unilateral clamping of the renal pedicle for 26 min and also undergo simultaneous contralateral nephrectomy. Using this approach, 50-60% of mice develop moderate AKI 24 hr after injury but 90-100% of mice survive. To induce more severe AKI, BALB/c mice undergo renal pedicle clamping for 30 min followed by contralateral nephrectomy 8 days after injury. This allows functional assessment of renal recovery after injury with 90-100% survival. Early post-injury tubular damage as well as post injury fibrosis are highly consistent using this model.

We describe models of moderate and severe ischemia-reperfusion-induced kidney injury in which the mice undergo unilateral renal pedicle clamping followed by simultaneous or delayed contralateral nephrectomy, respectively. These models consistently give rise to renal dysfunction and post-injury fibrosis, but injury severity and survival are dependent on mouse background, age and surgical equipment.

Ischemia-reperfusion induced acute kidney injury (IR-AKI) is widely used as a model of AKI in mice, but results are often quite variable with high, often ...

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 Model of Intestinal Ischemia-reperfusion Injury

Intestinal ischemia is a life-threatening condition associated with a broad range of clinical conditions including atherosclerosis, thrombosis, hypotension, necrotizing enterocolitis, bowel transplantation, trauma and chronic inflammation. Intestinal ischemia-reperfusion (IR) injury is a consequence of acute mesenteric ischemia, caused by inadequate blood flow through the mesenteric vessels, resulting in intestinal damage. Reperfusion following ischemia can further exacerbate damage of the intestine. The mechanisms of IR injury are complex and poorly understood. Therefore, experimental small animal models are critical for understanding the pathophysiology of IR injury and the development of novel therapies.
Here we describe a mouse model of acute intestinal IR injury that provides reproducible injury of the small intestine without mortality. This is achieved by inducing ischemia in the region of the distal ileum by temporally occluding the peripheral and terminal collateral branches of the superior mesenteric artery for 60 min using microvascular clips. Reperfusion for 1 hr, or 2 hr after injury results in reproducible injury of the intestine examined by histological analysis. Proper position of the microvascular clips is critical for the procedure. Therefore the video clip provides a detailed visual step-by-step description of this technique. This model of intestinal IR injury can be utilized to study the cellular and molecular mechanisms of injury and regeneration.

Here we describe the detailed procedure of intestinal ischemia-reperfusion in mice which results in reproducible injury without mortality to encourage the standardization of this technique across the field. This model of intestinal ischemia-reperfusion injury can be utilized to study the cellular and molecular mechanisms of injury and regeneration.

Intestinal ischemia is a life-threatening condition associated with a broad range of clinical conditions including atherosclerosis, thrombosis, ...

Evaluation of Coronary Flow Reserve After Myocardial Ischemia Reperfusion in Rats

Coronary artery disease is the leading cause of death worldwide. After an acute myocardial infarction, early and successful myocardial intervention via recanalization of the coronary artery is the most effective strategy for reducing the size of ischemic myocardium. The coronary microvasculature cannot be visualized and imaged&#160;in vivo, but there are several invasive and noninvasive techniques that can be used to assess parameters which depend directly on coronary microvascular function. The endothelial function after ischemia reperfusion can be assessed also at the level of the coronary circulation via the coronary flow reserve (CFR).&#160;In this study, peak velocity of left anterior descending (LAD) coronary arteries was measured in rats in vivo via Transthoracic Doppler Echocardiography during resting and stress challenge (induced by Dobutamine). A normal heart can increase its coronary blood flow up to four times above the resting values during stress induction. Following ischemia reperfusion, we found a significantly diminished CFR, which can be used as a marker of coronary microvascular dysfunction. CFR has opened a window on the importance of microvascular dysfunction and has been shown to predict cardiovascular risk independent of whether the severe obstructive disease is present.

Coronary flow reserve (CFR), is defined as the ratio of maximal coronary blood flow to the resting coronary blood flow. We present a protocol for evaluating CFR in rats via ultrasound, which offers the opportunity to predict cardiovascular risk factors in the absence of obstructive coronary disease.

Coronary artery disease is the leading cause of death worldwide. After an acute myocardial infarction, early and successful myocardial intervention via ...

Delayed Intramyocardial Delivery of Stem Cells after Ischemia Reperfusion Injury in a Murine Model

There is significant interest in the use of stem cells (SCs) for the recovery of cardiac function in individuals with myocardial injuries. Most commonly, cardiac stem cell therapy is studied by delivering SCs concurrently with the induction of myocardial injury. However, this approach presents two significant limitations: the early hostile pro-inflammatory ischemic environment may affect the survival of transplanted SCs, and it does not represent the subacute infarction scenario where SCs will likely be used. Here we describe a two-part series of surgical procedures for the induction of ischemia-reperfusion injury and delivery of mesenchymal stem cells (MSCs). This method of stem cell administration may allow for the longer viability and retention around damaged tissue by circumventing the initial immune response. A model of ischemia reperfusion injury was induced in mice accompanied by the delivery of mesenchymal stem cells (3.0 x 105), stably expressing the reporter gene firefly luciferase under the constitutively expressed CMV promoter, intramyocardially 7 days later. The animals were imaged via ultrasound and bioluminescent imaging for confirmation of injury and injection of cells, respectively. Importantly, there was no added complication rate when performing this two-procedure approach for SC delivery. This method of stem cell administration, collectively with the utilization of state-of-the-art reporter genes, may allow for the in vivo study of viability and retention of transplanted SCs in a situation of chronic ischemia commonly seen clinically, while also circumventing the initial pro-inflammatory response. In summary, we established a protocol for the delayed delivery of stem cells into the myocardium, which can be used as a potential new approach in promoting regeneration of the damaged tissue.

Stems cells are continuously investigated as potential treatments for individuals with myocardial damage, however, their decreased viability and retention within injured tissue can impact their long-term efficacy. In this manuscript we describe an alternative method for stem cell delivery in a murine model of ischemia reperfusion injury.

There is significant interest in the use of stem cells (SCs) for the recovery of cardiac function in individuals with myocardial injuries. Most commonly, ...

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 ...

An Effective Mouse Model of Unilateral Renal Ischemia-Reperfusion Injury

Ischemia-reperfusion injury (IRI) is the leading cause of acute renal failure and is a significant contributor to delayed graft function. Animal models are the only available resources that mimic the complexities of the IRI-associated damage encountered in vivo. This paper describes an effective mouse model of unilateral renal IRI that delivers highly reproducible data. Ischemia is induced by occluding the right renal pedicle for 30 min followed by reperfusion. In addition to the surgical procedure, a sequential overview of the expected physiological and histopathological changes following renal IRI will be provided by comparing data from seven different reperfusion times (4 h, 8 h, 16 h, 1 day, 2 days, 4 days, and 7 days). Critical data for planning experiments ahead, such as mean surgical time, average anesthetic consumption, and body weight changes over time, will be shared. This work will help researchers implement a reliable renal IRI model and select the appropriate reperfusion time that aligns with their intended investigative goals.

Renal ischemia-reperfusion injury is associated with high morbidity and mortality in hospitalized patients. Here, we present a simple and effective mouse model of unilateral renal ischemia-reperfusion injury and provide a sequential overview of representative pathological changes observed in the kidney.

Ischemia-reperfusion injury (IRI) is the leading cause of acute renal failure and is a significant contributor to delayed graft function. Animal models ...

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 ...