Murine Left Pulmonary Hilar Clamp Model of Lung Ischemia Reperfusion Injury

Abstract

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 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.

Introduction

IRI during organ transplantation is a major risk factor for primary graft dysfunction and later episodes of graft rejection^1,2. 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 ischemia^4,5,6, 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 clamp^{8,9,10,11,12,13}, a Rumel tourniquet^14,15, 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 receptor^14,18,19,20. In the following protocol, we present a reliable, teachable, and reproducible method of hilar clamping as a tool for studying lung IRI.

Protocol

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, 8^th 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

1. Select a mouse that weighs at least 25 g. This will facilitate easier intubation due to the larger orifice between the vocal cords.
2. 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 ½ cc syringe with a ½ inch 29G needle. Wait about 5 min for ketamine to take effect - the mouse should not exhibit spontaneous movement which confirms adequate anesthesia, however, it may still twitch with a paw pinch.
3. Inject buprenorphine (dose 0.05 mg/kg of body weight) subcutaneously prior to surgery for additional pain control.
4. Apply non-medicated ophthalmic ointment over both eyes to avoid corneal desiccation during surgery.
5. 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).
6. Once the mouse is anesthetized, perform the intubation and remainder of the procedure under a microscope, over a warmed mat at ~37 °C (to maintain normothermia in the mouse).
7. Suspend the mouse vertically by applying an atraumatic clamp on the tip of its tongue, with its back facing toward the surgeon. The rump of the mouse can rest on the mat (see Figure 1B and Supplementary Figure 1A).
8. Pull on the whiskers to expose the oropharynx. Use the prongs of a curved mosquito clamp to depress the tongue and spread the oropharynx in the anterior-posterior axis. When the oropharynx is sufficiently opened, there should be a clear view of the vocal cords (through the microscope objective). The vocal cords of an anesthetized mouse should still open and close spontaneously.
9. Insert a homemade introducer (see Supplementary Figure 1B) with a curve on the end into a 1 inch 20G angiocatheter, which is used as the endotracheal tube (ETT). Use the non-dominant hand to keep the oropharynx open with a curved mosquito (see Figure 1C). With a clear view of the vocal cords, use the dominant hand to 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).
10. 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.
11. Once confirmed to be in position, secure the ETT with a 5 cm strip of 1 inch silk tape around the mouse nose, ensuring there is adequate contact with the ETT and nose so that the ETT will not slide out of the mouth (see Figure 1E).
12. To maintain adequate anesthesia throughout the surgery, administer 1%-1.5% isoflurane in-line with the oxygen flow.

2. Thoracotomy

1. 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).
2. Apply alcohol (70%) over the left chest prior to incision.
3. Perform an incision over the 4^th intercostal space from the anterior axillary line to the posterior axillary line, using scissors to open the overlying skin (see Figure 2A). The 4^th intercostal space is located approximately 1 cm below the axilla in the mid-axillary line.
4. To minimize blood loss, coagulate visible blood vessels within the subcutaneous and muscle layers using a handheld cautery pen.
5. Divide the latissimus dorsi and serratus anterior muscles sharply using scissors along the length of the skin incision (see Figure 2B).
6. Using fine curved forceps, carefully elevate the 4^th rib (taking care not to injure the underlying lung) and make a pinpoint snip with fine scissors just above the 5^th rib to enter the 4^th intercostal space (see Figure 2C). Perform the thoracotomy above the 5^th rib rather than below the 4^th rib to avoid injuring the intercostal neurovascular bundle.
7. 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 4^th intercostal space just above the 5^th 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.
8. 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).

3. Application of hilar clamp

1. 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.
2. 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).
3. Then, flip the left lung posteriorly to lay over the tie and pull both ends of the tie anteriorly (see Figure 3D).
4. 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).
   1. 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).
   2. 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.
5. 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.
6. 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.

4. Release of hilar clamp

1. 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).
2. During the reperfusion period, close the chest in three layers to minimize insensible losses. First, close the 4^th rib space with a single stitch with 6-0 nylon suture, with one bite just above the 4^th rib and one bite just above the 6^th rib (see Figure 4B-C). Tie this at maximum lung inflation, to minimize risk of an iatrogenic pneumothorax (see Figure 4D-E).
3. Next, close the muscle layer using a simple running stitch with 6-0 nylon suture.
4. 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.
5. 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.
6. 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.
   1. For survival surgery, inject 1 mL of warm saline subcutaneously to account for fluid losses from surgery. Administer additional pain control with carprofen (dose 5 mg/kg of body weight) subcutaneously every 6-8 h and buprenorphine (dose 0.05 mg/kg of body weight) subcutaneously every 8-12 h, for a total duration of 48 h.

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 clamped^22,23, during which time only the left lung is performing oxygenation and ventilation.

1. If the mouse has been awoken from anesthesia after hilar clamp, re-anesthetize and intubate the mouse according to step 1.
2. 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.
3. 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).
4. 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).
5. 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).
6. 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).
7. 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).
8. 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.
9. 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.
10. 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.
11. Coat a ½ inch 31G needle on a 1 cc tuberculin syringe with about 200 µL 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 .
12. 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 µL).
    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.
13. Run the arterial blood on an ABG machine to obtain oxygen saturation, partial pressure of oxygen, partial pressure of carbon dioxide, among other measurements.

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 µL 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.

1. 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.
2. Perform laparotomy according to steps 5.3-5.4.
3. 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).
4. Clear the fat overlying the IVC bluntly with cotton-tipped applicators.
5. 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).
6. Return the eviscerated bowel into the abdomen over the cotton swab to continue to apply pressure over the IVC (see Figure 6C).
7. 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.

7. Histology (H&E) staining

1. Following formalin-fixation and paraffin-embedding of the left lung and sectioning to a thickness of 5 µm, deparaffinize the slide in xylene (two 10 min washes).
2. Dehydrate in sequential ethanol washes: 2x 5 min washes with 100%, 1x 2 min wash with 95%, and 1x 2 min wash with 70% ethanol. Then, rinse in deionized water.
3. 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.
4. 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.
5. Dehydrate further by dipping in 95% ethanol 15x. Stain in eosin for 1 min.
6. Dehydrate with 2 min ethanol washes (2x), followed by 2 min xylene washes (2x). Apply mounting media and cover slip.

Representative Results

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&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 naïve 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 antibodies^18,24. 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.

Figure 1: Mouse intubation. (A) Mouse with left chest and abdomen shaved. (B) Clamp the tongue with an atraumatic clamp, allowing the mouse to sit vertically. (C) Under microscope guidance, open the oropharynx with a mosquito clamp to visualize the vocal cords and intubate the mouse. (D) Endotracheal tube connected to ventilator tubing. (E) Once intubation is confirmed, tape the tube circumferentially to the nose of the mouse. (F) Intubated mouse with inflow and outflow tubing from and to small animal ventilator. Please click here to view a larger version of this figure.

Figure 2: Left thoracotomy. (A) Skin incision over 4^th intercostal space (ICS). (B) Muscle layers divided over 4^th intercostal space. (C) Making small incision just above 5^th rib. (D) Extending incision anteriorly and posteriorly in 4^th ICS. (E) Lung visualized in 4^th ICS. (F-G) Rib retractors placed. Please click here to view a larger version of this figure.

Figure 3: Left hilar clamp. (A) Left lung mobilized with cotton swabs. (B) Inferior pulmonary ligament (IPL) visualized. (C) With left lung flipped anteriorly, silk tie is placed posterior to the hilum. (D) With left flipped posteriorly over silk tie, anterior hilum is visualized. (E) Instrument tie of slipknot over left hilum. (F) Slipknot on hilum. (G) With manual occlusion of outflow, left lung does not inflate, indicating successful hilar clamp. Please click here to view a larger version of this figure.

Figure 4: Release of left hilar clamp and left chest closure. (A) With slipknot released, the left lung is reinflated. (B) Rib closure with one bite just above 4^th rib and (C) one bite just above 6^th rib. (D-E) Chest closed with one stitch tied down at end-inspiration. (F) Skin closure with interrupted stitches (after muscle layer closure with running stitch). Please click here to view a larger version of this figure.

Figure 5: Laparotomy, bilateral clamshell thoracotomy, and arterial blood collection. (A) Mouse adjusted to supine position. (B) Skin incision made over midline abdomen. (C) Laparotomy extending from xiphoid to pubis, with extension left and right along the most inferior rib. (D-E) Opening of the diaphragm to enter the chest cavity, with extension of diaphragm incision left and right along most inferior rib (dotted line). (F) Bilateral clamshell thoracotomy with incisions in anterior axillary lines (dotted line) toward apex of chest. (G) Accessory lobe of right lung extending into the left chest posterior to the inferior vena cava. (H) 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). Please click here to view a larger version of this figure.

Figure 6: Intravenous injection into the inferior vena cava. (A) Right medial visceral rotation to expose inferior vena cava (IVC). (B) Injection of solution into IVC. (C) Applying pressure over IVC to achieve hemostasis. Please click here to view a larger version of this figure.

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 ± SEM, n=6-7. p values calculated by Mann-Whitney U test. **p <0.01. (B) H&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 µm. (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. Please click here to view a larger version of this figure.

Supplementary Figure 1: (A) Clamp set-up for intubation. (B) Introducer for endotracheal tube. Please click here to download this File.

Discussion

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-TNFα-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 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 ischemia^4,5,6. 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 clamp^{8,9,10,11,12,13}, a Rumel tourniquet^14,15, 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 strains^26,27. 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 failure^30,31. 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 lung^18,24. 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'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 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 4^th and 5^th 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 function^22,23. 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.

Materials

Name	Company	Catalog Number	Comments
Medications
10% povidone-iodine solution	Aplicare	NDC 52380-0126-2	For disinfectant
Buprenorphine 1.3 mg/mL	Fidelis Animal Health	NDC 86084-100-30	For pain control
Carprofen	Cronus Pharma	NDC 69043-027-18	For pain control
Heparin 1000 units/mL	Sagent	NDC 25021-404-01	For obtaining arterial blood
Isoflurane 1%-1.5%	Sigma Aldrich	26675-46-7	For anesthesia
Ketamine hydrochloride 100 mg/mL	Vedco	NDC 50989-996-06	For anesthesia
Puralube Vet eye ointment	Medi-Vet.com	11897	To prevent eye dessiccation
Xylazine 20 mg/mL	Akorn	NDC 59399-110-20	For pain control
Tools and Instruments
Argent High Temp Fine Tip Cautery Pen	McKesson	231	To coagulate blood vessels
Curved mosquito clamp	Fine Science Tools	13009-12	For surgical procedure
Fine curved forceps	Fine Science Tools	11274-20	For surgical procedure
Fine scissors	Fine Science Tools	15040-11	For surgical procedure
Intubation clamp set-up	Fine Science Tools	18374-44, 18144-30	For holding mouse vertically by the tongue during intubation. See Supplementary Figure 1A.
Magnetic rib retractors	Fine Science Tools	18200-01, 18200-10	For retraction of thoracotomy. Magnetic fixator and retractor should be connected by micro latex tubing below.
Optical Grade Plastic Optical Fiber Unjacketed, 500μm	Edmund Optics	02-532	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.
Power Pro Ultra clipper	Oster	078400-020-001	To clip hair
Scissors	Fine Science Tools	14370-22	For surgical procedure
Small animal heating pad	K&H Pet Products	Thermo-Peep Heated Pad	To maintain normothermia
Small animal ventilator	Harvard Apparatus	55-0000	For ventilation (TV 0.35 cc, PEEP 1 cm H2O, RR 100-105/min, FiO2 100%)
Spearit Micro Latex Rubber Tubing (1/8 in outside diameter, 1/16 in inside diameter)	Amazon.com	https://www.amazon.com/Rubber-Tubing-CONTINUOUS-Select-Length/dp/B00H4MT7V0?th=1	For retraction of thoracotomy
Stat Profile Prime Critical Care Blood Gas Analyzer	Nova Biomedical	https://novabiomedical.com/prime-plus-critical-care-blood-gas-analyzer/index.php?gad=1&gclid=Cj0KCQjwmICoBhDx ARIsABXkXlInZX--R3ezBkc304nS_GVGI9Z2T3Esr33 2aM8WGPiUVhicPQZ Wj2AaAqhDEALw_wcB	For retraction of thoracotomy
Straight clamp	Fine Science Tools	13008-12	For surgical procedure
Straight forceps	Fine Science Tools	91113-10	For surgical procedure
Surgical microscope	Wild Heerbrugg	no longer produced	For intubation and surgical procedure; recommend replacement with Leica surgical microscopes
Supplies
½ cc syringe with ½ inch 29G needle	McKesson	942665	For injecting ketamine/xylazine intraperitoneally
½ inch 31G needle on a 1 cc tuberculin syringe	McKesson	16-SNT1C2705	For aspiration of arterial blood from left ventricle
1-inch 20G IV catheter	Terumo	SROX2025CA	For endotracheal tube (ETT)
1-inch silk tape	Durapore	3M ID 7100057168	To tape ETT to nose and to secure limbs
3/10 cc syringe with 5/16 inch 31G needle	McKesson	102-SN310C31516P	For antibody injection into the inferior vena cava
6-0 monofilament suture on a P-10 needle	McKesson	S697GX	For closure of thoracotomy, muscle layer, and skin
6-0 silk tie	Surgical Specialties Look	SP102	To make slipknot for hilar clamp
Pointed cotton-tipped applicators	Solon	56225	To manipulate lung and for blunt dissection