Immunology and Infection
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A Mouse Model of Orotracheal Intubation and Ventilated Lung Ischemia Reperfusion Surgery
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Summary September 9th, 2022
A mouse surgical model to create left lung ischemia reperfusion (IR) injury while maintaining ventilation and avoiding hypoxia.
Transcript
Ventilated lung ischemia-reperfusion surgery can be used to study the lung specific pathophysiology of multiple processes, including lung transplantation, pulmonary embolism, and lung injury following hemorrhagic trauma with resuscitation. This model minimizes the inflammatory contributions of atelectasis, mechanical ventilation, and hypoxia. It maintains an intact in vivo circulatory immune system and permits the longer-term studies.
This model could provide insight into how sterile inflammation is controlled and regulated within the lung. The microsurgical technique first requires hours of practice. Careful organization and planning are important before beginning.
Also, it can be initially practiced by the euthanized mouse without the distraction of cardiac activity and the movement it causes. The blood stasis in the left PA allows it to be more easily visualized and manipulated. To begin, gently place the fiber optic flexible light on the trachea of an anesthetized mouse, slightly below the vocal cords.
Adjust the illumination levels such that only a dark field is visible when looking into the mouse oral pharynx, except for red light emanating from below the vocal cords. For intubation, hold the tweezers with the dominant hand and use them to gently grip and draw the tongue out of the oral cavity. Open the lower jaw using forceps held by the non-dominant hand and push the forceps into the larynx to lift the epiglottis.
At this point, release the tongue from the tweezers. Observe the vocal chords. They should open and close according to each breath.
Holding the cannula with the guide wire preloaded, insert the tip of the wire through the vocal cords. Next, being very careful not to move the wire by holding a portion of it that is outside of the cannula but just above the vocal cords, withdraw the cannula, leaving just the wire in place with its distal end within the trachea. At this point, perform a second visualization of the vocal cords to confirm that the wire distal tip remains passed through the illuminated vocal cords and into the trachea, and is not in the unlit esophagus.
Next, hold the wire outside the mouth with the curved forceps in the left hand stabilized against a hard surface, and carefully advance the 20 gauge catheter with tape wings over the wire. Once the distal end of the wire emerges from the back end of the catheter or endotracheal tube hold that end with the curved forceps and smoothly advance the catheter into the trachea. Next, carefully remove the wire from the distal end of the catheter with the curved forceps without dislodging the placement of the catheter.
Next, briefly connect the catheter to the ventilator before securing it to confirm proper placement into the trachea and not the esophagus. Confirm tracheal placement by observation of mechanical ventilation dependent bilateral chest wall movements and the absence of inflation of the stomach. After intubation, connect the catheter to the ventilator set at title volume of 0.2 to 0.225 milliliter, and respiratory rate of 120 to 150 breaths per minute to confirm the correct tracheal placement of the orotracheal tube.
Then disconnect with the mouse breathing spontaneously through the orotracheal tube. Shave the mouse hair over the left thorax area up to the left scapula. Remove excess shaved hair using alcohol swabs and disinfect the surgical area.
Then place the mouse on a warming pad in a left lateral or three fourths turned position. Connect the tracheal tube to the ventilator. After making the skin incision insert three sterilized retractors underneath the muscular layer.
Next, identify the second intercostal space and hold the second rib with the extra fine forceps. Then, pulling the rib upward, use a sterile number 12 curved scalpel blade to enter the plural space by separating and cutting across the second to third spaces intercostal muscles. Consider pausing ventilation to reduce injury to the left lung apex.
Use the smallest, narrowest retractor cephalad along the orientation of the ribs, and medium size retractor to the left along the third rib, and the largest retractor to the right along the surface of the second rib. Open the chest with slow and progressive retraction using the elastic retractor cords. Expose and identify the left pulmonary artery, or PA by moving the left lung apex away with a sterile cotton tip swab or a surgical sponge.
Use the micro forceps ultra fine forceps in the right hand and PA or vessel dilating forceps in the left hand to gently expose and create the field in which the left PA and bronchus are both visible. Using the PA forceps, pick up the left PA and pull gently but firmly upward and cephalad to visualize the transparent bronchus below. Increase the magnification on the dissection microscope to four X.While retracting the PA away from the bronchus, carefully pass the closed ultra fine forceps through the space between the left PA and bronchus.
Then use these forceps to hold and pull a 7-0 or 8-0 prolene suture through the space between the left pulmonary artery above and the bronchus below. Next, encircle the left PA by tying a slip knot to create an occlusion in the PA.Blood flow interruption is easily visualized at this point, marking the ischemic period's initiation. Externalize the free end of the knot through a different entry point in the anterior left thorax using a 24 to 28 gauge needle and secure the end of the suture with a small piece of tape for easier identification later.
Next, using a positive and expiratory pressure valve or tubing on the rodent ventilator, reinflate the lung to expel as much air out of the chest cavity as possible. Then, close the ribcage with two interrupted 4-0 nylon sutures, followed by closing the muscle and skin layer, and applying local anesthesia. Disconnect the mouse from the ventilator and carefully place it on the warming pad for postoperative care to maintain body temperature during recovery.
Ensure that the externalized slip knot is controlled and visualized clearly during the movement of the mouse. Pull the externalized slip knot gently at the end of the ischemic period. The histology of lung sections in wild type mice of strains C3H and C57 black six is shown here.
After one hour ischemia and three hour reperfusion, intense neutrophilic infiltration within the lung left tissue was observed in both strains. However, the C3H strain showed markedly greater levels of inflammation compared to C57 black six. Mistakes during the passage of the monofilament between the left PA and the left bronchus can lead to unsalvaged surgery with catastrophic bleeding of the left PA or irreversible injury to the left bronchus.
This is the step that is most technically challenging and requires repeat practicing when first learning this procedure. After mouse recovery from lung ischemia-reperfusion surgery about one hour or less after reperfusion, an intratracheal instillation of live bacteria can be administered at three hours or 24 to 48 hours later to simulate an infection that follows sterile lung injury caused by ischemia-reperfusion. Intratracheal installation of other agents that can modulate the sterile inflammation can also be done after ischemia-reperfusion in a similar manner.
This technique helped discover and confirm the step-wise trafficking and activation of neutrophils in response to sterile injury and infection in the lung.
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