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.
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.
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.
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
2. Thoracotomy
3. Application of hilar clamp
4. Release of hilar clamp
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.
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.
7. Histology (H&E) staining
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 antibodies18,24. Representative plots show that ~75% of neutrophils are intravascular (APC-Ly6G+FITC-Ly6G+) and ~25% are interstitial (APC-Ly6G+FITC-Ly6G–) after hilar clamping of the left lung (Figure 7C, n=4). These results demonstrate a reliable way to assess neutrophil extravasation and can be adapted to other cell types depending on the antibodies used (i.e., anti-Ly6C for monocytes, anti-CD19 for B cells). This technique is useful for the study of cell dynamics following IRI.
Figure 1: Mouse intubation. (A) Mouse with left chest and abdomen shaved. (B) Endotracheal tube connected to ventilator tubing. (C) Once intubation is confirmed, tape the tube circumferentially to the nose of the mouse. (D) Intubated mouse with inflow and outflow tubing from and to the small animal ventilator. Please click here to view a larger version of this figure.
Figure 2: Left thoracotomy. (A) Skin incision over 4th intercostal space (ICS). (B) Muscle layers divided over 4th intercostal space. (C) Making small incision just above 5th rib. (D) Extending incision anteriorly and posteriorly in 4th ICS. (E) Lung visualized in 4th 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 4th rib and (C) one bite just above 6th 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: Introducer for endotracheal tube. Please click here to download this File.
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 ischemia4,5,6. Furthermore, transplant centers are increasingly adopting DCD lung transplants, which are accompanied by much longer warm ischemic times than traditional DBD lung transplants7. Thus, detailed study of the events during warm ischemia and the subsequent reperfusion period is critical and can be accomplished with this hilar clamp model.
Many variations of the hilar clamp technique have been created over the last couple decades. These include the use of an atraumatic microvascular clamp8,9,10,11,12,13, a Rumel tourniquet14,15, and a tie with a slipknot around the hilum16. We describe our adaptation of the slipknot technique in mice. The main advantage this technique has over other hilar clamp techniques is that it allows for closure of the chest during the period of ischemia, which minimizes insensible fluid losses particularly in protocols with longer periods of ischemia. This cannot be accomplished when a clamp or tourniquet is placed on the hilum.
Regarding animal selection, we chose to develop this model in mouse as opposed to rat or large animal models, such as pig. The reasoning is two-fold – our laboratory has extensive experience in mouse microsurgery with orthotopic mouse lung transplantation17. Secondly, while mice are smaller and more technically challenging to operate on than rats, there are presently more commercially available transgenic mouse strains than rat strains26,27. Mice also breed more quickly, allowing for faster development of new strains, which is particularly useful when investigating immune mechanisms of IRI. For these reasons, we have chosen to develop this technique in mice.
To our knowledge, this is the first video-based article using the technique of left lung hilar clamp in mouse with a slipknot. Saito et al have published a video-based article demonstrating hilar clamp in rat using a microvascular clamp22. In mouse, Liao et al published a lung IRI model with pulmonary artery (PA) clamping with a slipknot, however the lung remains ventilated in this model as the bronchus is excluded from the clamp28. The PA clamping technique has been used as a model to study pulmonary embolism29, pulmonary hypertension and/or right ventricular failure30,31. Since the area of study is lung IRI in transplantation, we elected to clamp the entire hilum including the bronchus to minimize barotrauma on top of the ischemic injury.
Of note, 30 min of warm ischemia was chosen followed by 1 h of reperfusion because we are interested in the dynamics of lung-infiltrating cells in the very early period post-IRI. For example, studies have reported upregulation of cytoprotective genes, such as heme oxygenase 1, after just 30 min of ischemia followed by 2 h of reperfusion13. Neutrophils were previously thought to be the central cell type infiltrating the lung after IRI32, however we showed that monocytes precede neutrophils in their arrival to the lung18,24. More recently, we found that B cells precede even monocytes and enter the lung within 1 h of reperfusion (unpublished data). When adapting this technique, the durations of warm ischemia and reperfusion can be adjusted according to the study question and hypothesis. Longer periods of warm ischemia are possible without concern for excessive fluid and heat loss since the chest can be closed during this period.
Several complications can occur perioperatively, with the final common outcome being perioperative demise. We have observed that the incidence of animal death during surgery decreases with repetitions of the technique by the surgeon. We estimate that surgeons require at least 2 months of practice and/or at least 20 cases to reach a plateau in skill. At this point, we estimate that intraoperative mortality occurs in approximately 1 in 50 cases. Death in the anesthetized mouse is often heralded by agonal breaths that involve forceful contractions of the diaphragm and chest wall musculature. Upon the onset of these agonal breaths, death will occur in 3-5 min if no intervention is taken. There are several causes that can precipitate intraoperative demise, many of which are recognizable and preventable. We will highlight below the critical steps of this surgery and troubleshooting techniques to achieve a successful outcome.
Intubation
The first critical step is careful intubation and securing of the airway to prevent inadvertent extubation. In a survival surgery, it is particularly important to ensure that the vocal cords are not damaged during initial intubation, as the mouse will not be able to breathe following extubation. Advancement of the ETT past the cords should be done under direct visualization through the microscope. If resistance is met during intubation, the ETT should not be advanced blindly. While keeping the oropharynx open with the mosquito clamp, care should be taken to not pierce the oral mucosa as blood will obscure view of the airway. The number of attempts at intubation should be limited to less than five.
Accidental extubation
If at any point during the procedure, the lungs do not appear to be inflating appropriately, it is possible that the ETT has dislodged. Another manifestation of accidental extubation that goes unnoticed for a prolonged period is the mouse suddenly developing agonal breathing. Following initial intubation, care should be taken to secure the ETT well to the nose circumferentially with silk tape, and adjustment of the mouse'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 4th and 5th ribs should be brought together when the lung is at maximum end-inspiration. This will minimize the amount of air remaining in the pleural cavity, which would result in pneumothorax once the ribs are closed. The muscle layers and skin should be closed in an airtight manner to avoid development of tension pneumothorax.
Obtaining ABGs
Finally, when obtaining ABGs, it is important to first clamp the right hilum so that the ABG measurement will be reflective of only left lung function22,23. Without the right hilar clamp, the normally ventilating right lung will more than compensate for the damaged ischemic left lung and ABG measurements will appear normal.
In summary, the hilar clamp technique that we describe above is a valuable tool for the study of lung transplantation, and importantly can isolate the study of warm IRI without the introduction of allogenicity, surgical anastomosis-related trauma, or cold ischemic storage. Notably, this described technique of hilar clamping is easily taught and widely adoptable and can lead to reliable and reproducible results.
The authors have nothing to disclose.
This work received no specific grant from any funding agency in the public, commercial, or not-for-profit sectors.
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 |