This work was funded by departmental support from the Department of Anesthesia and Perioperative Care, University of California San Francisco and San Francisco General Hospital, as well as by an NIH R01 award (to AP): 1R01HL146753.

All procedures and steps described below were approved by the institutional animal care and use committee (IACUC) at the University of California San Francisco. Any mouse strain can be used, though some strains have a more robust lung IR inflammatory response compared to others12. Mice that are approximately 12-15 weeks of age (30-40 g) or older tolerate and survive the lung IR surgery better than younger mice. Both male and female mice can be used for these surgeries.
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<ol>
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Lung Cellular and Molecular Physiology. 294 (4), 714-723 (2008).</li><li>Prakash, A., Kianian, F., Tian, X., Maruyama, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ferroptosis+mediates+inflammation+in+lung+ischemia-reperfusion+(IR)+sterile+injury+in+mice.">Ferroptosis mediates inflammation in lung ischemia-reperfusion (IR) sterile injury in mice.</a> American Journal of Respiratory and Critical Care Medicine. 201, (2020).</li><li>Tian, X., 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=NLRP3+inflammasome+mediates+dormant+neutrophil+recruitment+following+sterile+lung+injury+and+protects+against+subsequent+bacterial+pneumonia+in+mice.">NLRP3 inflammasome mediates dormant neutrophil recruitment following sterile lung injury and protects against subsequent bacterial pneumonia in mice.</a> Frontiers in Immunology. 8, 1337 (2017).</li><li>Tian, X., Hellman, J., Prakash, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Elevated+gut+microbiome-derived+propionate+levels+are+associated+with+reduced+sterile+lung+inflammation+and+bacterial+immunity+in+mice.">Elevated gut microbiome-derived propionate levels are associated with reduced sterile lung inflammation and bacterial immunity in mice.</a> Frontiers in Microbiology. 10, 159 (2019).</li><li>Liu, Q., Tian, X., Maruyama, D., Arjomandi, M., Prakash, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lung+immune+tone+via+gut-lung+axis:+Gut-derived+LPS+and+short-chain+fatty+acids'+immunometabolic+regulation+of+lung+IL-1&#946;,+FFAR2,+and+FFAR3+expression.">Lung immune tone via gut-lung axis: Gut-derived LPS and short-chain fatty acids' immunometabolic regulation of lung IL-1&#946;, FFAR2, and FFAR3 expression.</a> American Journal of Physiology. Lung Cellular and Molecular Physiology. 321 (1), 65-78 (2021).</li><li>Dodd-o, J. 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=Interactive+effects+of+mechanical+ventilation+and+kidney+health+on+lung+function+in+an+in+vivo+mouse+model.">Interactive effects of mechanical ventilation and kidney health on lung function in an in vivo mouse model.</a> American Journal of Physiology. Lung Cellular and Molecular Physiology. 296 (1), 3-11 (2009).</li></ol>

The authors declare that they have no competing financial interests.

This manuscript details the steps involved in performing the ventilated lung IR model developed by Dodd-o et al.9. This model has helped identify molecular pathways involved in the generation and resolution of inflammation from lung IR in isolation14,15,16,17, lung IR in combination with co-existing infection18, and lung IR in relation to the gut-...

Ischemia reperfusion (IR) injury can occur when blood flow is restored to an organ or tissue bed after some period of interruption. In the lung, IR can occur in isolation or in association with other injurious processes such as infection, hypoxia, atelectasis, volutrauma (from high tidal volumes during mechanical ventilation), barotrauma (high peak or sustained pressures during mechanical ventilation), or blunt (non-penetrating) lung contusion injury1,2,3. There remain several gaps in our knowledge about the mechanisms of LIRI and the impact of concurrent processes (e.g., infection) on LIRI outcomes, and also the treatment options for LIRI are limited. An in vivo model of pure LIRI is required to identify the pathophysiology of lung IR injury in isolation and to study its contribution to any multi-hit process of which lung injury is a component.
Murine lung IR models can be used to study the lung-specific pathophysiology of multiple processes, including lung transplantation3, pulmonary embolism4, and lung injury following hemorrhagic trauma with resuscitation5. Currently used models include surgical lung transplantation6, hilar clamping7, ex vivo lung perfusion8, and ventilated lung IR9. Here, we provide a detailed protocol for a murine ventilated lung IR model of sterile lung injury. There are multiple benefits of this approach (Figure 2), including that it induces minimal hypoxia and minimal atelectasis, and it is a survival surgery model that allows for long-term studies.
Reasons to choose this model of LIRI over other models such as the hilar clamping and ex vivo perfusion models are the following: this model minimizes the inflammatory contributions of atelectasis, mechanical ventilation, and hypoxia; it preserves cyclical ventilation; it maintains an intact in vivo circulatory immune system that can respond to the IR injury; and finally, as a survival procedure, it permits the longer-term analysis of the mechanisms of secondary injury generation (2-hit models) and injury resolution. Overall, we believe this ventilated lung IR model provides the &#34;purest&#34; form of IR injury that can be studied experimentally.
Other publications have described the use of orotracheal intubation of mice to perform IT injections or installations10,11, but not as the starting point for a survival surgery as it is in this model. The placement of an orotracheal tube permits the performance of lung surgery by allowing the collapse of the operative lung. It also allows for the reinflation of the lung at the end of the procedure, which is critical for the pneumothorax and for the ability of the mouse to return to spontaneous ventilation at the conclusion of the procedures. Finally, the removal of the secured orotracheal tube is a simple procedure that, unlike an invasive tracheotomy, is compatible with a survival surgery. This allows for longer term research studies focused on understanding the progression and resolution of LIRI and associated disorders, as well as the creation of chronic injury models.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Equipment</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fiber Optic Light Pipe</td>
			<td>Cole-Parmer</td>
			<td>UX-41720-65</td>
			<td>Fiberoptic light pipe</td>
		</tr>
		<tr>
			<td>Fiber Optic Light Source</td>
			<td>AmScope</td>
			<td>SKU: CL-HL250-B</td>
			<td>Light source for fiberoptic lights</td>
		</tr>
		<tr>
			<td>Germinator 500</td>
			<td>Cell Point Scientific, Inc.</td>
			<td>No.5-1450</td>
			<td>Bead Sterilizer</td>
		</tr>
		<tr>
			<td>Heating Pad</td>
			<td>AIMS</td>
			<td>14-370-223</td>
			<td>Alternative option</td>
		</tr>
		<tr>
			<td>Lithium.Ion Grooming Kits(hair clipper)</td>
			<td>WAHL home products</td>
			<td>SKU 09854-600B</td>
			<td>To remove mouse hair on surgical site</td>
		</tr>
		<tr>
			<td>Microscope</td>
			<td>Nikon</td>
			<td>SMZ-10</td>
			<td>Other newer options available at the company website</td>
		</tr>
		<tr>
			<td>MiniVent Ventilator</td>
			<td>Havard Apparatus</td>
			<td>Model 845</td>
			<td>Mouse ventilator</td>
		</tr>
		<tr>
			<td>Ultrasonic Cleaner</td>
			<td>Cole-Parmer</td>
			<td>UX-08895-05</td>
			<td>Clean tools that been used in operation</td>
		</tr>
		<tr>
			<td>Warming Pad</td>
			<td>Kent Scientific</td>
			<td>RT-0501</td>
			<td>To keep mouse warm while recovering from surgery</td>
		</tr>
		<tr>
			<td>Weighing Scale</td>
			<td>Cole-Parmer</td>
			<td>UX-11003-41</td>
			<td>Weighing scale</td>
		</tr>
		<tr>
			<td>Surgery Tools</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>4-0 Silk Suture</td>
			<td>Ethicon</td>
			<td>683G</td>
			<td>For closing muscle layer</td>
		</tr>
		<tr>
			<td>7-0 Prolene Suture</td>
			<td>Ethicon Industry</td>
			<td>EP8734H</td>
			<td>Using for making a slip knot of left pulmonary artery</td>
		</tr>
		<tr>
			<td>Bard-Parker (11) Scalpel (Rib-Back Carbon Steel Surgical Blade, sterile, single use)</td>
			<td>Aspen Surgical</td>
			<td>372611</td>
			<td>For entering thoracic cavity (option 1)</td>
		</tr>
		<tr>
			<td>Bard-Parker (12) Scalpel</td>
			<td>Aspen Surgical</td>
			<td>372612</td>
			<td>For entering thoracic cavity (option 2)</td>
		</tr>
		<tr>
			<td>Extra Fine Graefe Forceps</td>
			<td>FST</td>
			<td>11150-10</td>
			<td>Muscle/rib holding forceps</td>
		</tr>
		<tr>
			<td>Magnetic Fixator Retraction System</td>
			<td>FST</td>
			<td>1. Base Plate (Nos. 18200-03) 
			2. Fixators (Nos. 18200-01) 
			3. Retractors (Nos. 18200-05 through 18200-12) 
			4. Elastomer (Nos.18200-07) 5. Retractor(No.18200-08)</td>
			<td>Small Animal Retraction System</td>
		</tr>
		<tr>
			<td>Monoject Standard Hypodermic Needle</td>
			<td>COVIDIEN</td>
			<td>05-561-20</td>
			<td>For medication delivery IP</td>
		</tr>
		<tr>
			<td>Narrow Pattern Forceps</td>
			<td>FST</td>
			<td>11002-12</td>
			<td>Skin level forceps</td>
		</tr>
		<tr>
			<td>Needle holder/Needle driver</td>
			<td>FST</td>
			<td>12565-14</td>
			<td>for holding needles</td>
		</tr>
		<tr>
			<td>Needles</td>
			<td>BD</td>
			<td>305110</td>
			<td>26 gauge needle for externalizing slipknot (24 or 26 gauge needle okay too)</td>
		</tr>
		<tr>
			<td>PA/Vessel Dilating forceps</td>
			<td>FST</td>
			<td>00125-11</td>
			<td>To hold PA; non-damaging gripper</td>
		</tr>
		<tr>
			<td>Scissors</td>
			<td>FST</td>
			<td>14060-09</td>
			<td>Used for incision and cutting into the muscular layer durging surgery</td>
		</tr>
		<tr>
			<td>Ultra Fine Dumont micro forceps</td>
			<td>FST</td>
			<td>11295-10 (Dumont #5 forceps, Biology tip, tip dimension:0.05*0.02mm,11cm)</td>
			<td>For passing through the space between the left pulmonary artery and bronchus</td>
		</tr>
		<tr>
			<td>Reagents</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>0.25% Bupivacaine</td>
			<td>Hospira, Inc.</td>
			<td>0409-1159-02</td>
			<td>Topical analgesic used during surgical wound closure</td>
		</tr>
		<tr>
			<td>Avertin (2,2,2-Tribromoethanol)</td>
			<td>Sigma-Aldrich</td>
			<td>T48402-25G</td>
			<td>Anesthetic, using for anesthetize the mouse for IR surgery, the concentration used in IR surgery is 250-400 mg/kg.</td>
		</tr>
		<tr>
			<td>Buprenorphine</td>
			<td>Covetrus North America</td>
			<td>59122</td>
			<td>Analgesic: concentration used for surgery is 0.05-0.1 mg/kg</td>
		</tr>
		<tr>
			<td>Eye Lubricant</td>
			<td>BAUSCH+LOMB</td>
			<td>Soothe Lubricant Eye Ointment</td>
			<td>Relieves dryness of the eye</td>
		</tr>
		<tr>
			<td>Povidone-Iodine 10% Solution</td>
			<td>MEDLINE INDUSTRIES INC</td>
			<td>SKU MDS093944H (2 FL OZ, topical antiseptic)</td>
			<td>Topical liquid applied for an effective first aid antiseptic at beginning of surgery</td>
		</tr>
		<tr>
			<td>Materials</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Alcohol Swab</td>
			<td>BD brand</td>
			<td>&#160;BD 326895</td>
			<td>for sterilzing area of injection and surgery</td>
		</tr>
		<tr>
			<td>Plastic film</td>
			<td>KIRKLAND</td>
			<td>Stretch-Tite premium</td>
			<td>Alternative for covering the sterilized surgical field (more cost effective)</td>
		</tr>
		<tr>
			<td>Rodent Surgical Drapes</td>
			<td>Stoelting</td>
			<td>50981</td>
			<td>Sterile field or drape for surgical field</td>
		</tr>
		<tr>
			<td>Sterile Cotton Tipped Application</td>
			<td>Pwi-Wnaps</td>
			<td>703033</td>
			<td>used for applying eye lubricant</td>
		</tr>
		<tr>
			<td>Top Sponges</td>
			<td>Dukal Corporaton</td>
			<td>Reorder # 5360</td>
			<td>Stopping bleeding from skin/muscle</td>
		</tr>
	</tbody>
</table>

a mouse model of orotracheal intubation and ventilated lung ischemia reperfusion surgery

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

Inflammation generated by unilateral ventilated sterile lung ischemia reperfusion (IR) injury: Following 1 h of ischemia, we observed increased levels of cytokines in the serum and within the lung tissue by both ELISA and qRT-PCR that peaked at 1 h following reperfusion and rapidly returned to baseline within 12-24 h after reperfusion13. For samples collected at 3 h following reperfusion, we observed intense neutrophil infiltration within the left lung tissue and noted that the intensity of the in...

Watch this Scientific Journal Video about A Mouse Model of Orotracheal Intubation and Ventilated Lung Ischemia Reperfusion Surgery at JoVE.com

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

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

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

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.

a-mouse-model-orotracheal-intubation-ventilated-lung-ischemia

Research

JoVE Journal

Immunology and Infection

3.4K vistas.  University of California San Francisco and San Francisco General Hospital. La cirugía de isquemia-reperfusión pulmonar ventilada se puede utilizar para estudiar la fisiopatología específica del pulmón de múltiples procesos, incluido el trasplante de pulmón, la embolia pulmonar y la lesión pulmonar después de un traumatismo hemorrágico con reanimación. Este modelo minimiza las contribuciones inflamatorias de la atelectasia, la ventilación mecánica y la hipoxia. Mantiene intacto el sistema inmune circulatorio in vivo y permite los estudios a largo plazo.