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.
1. Mouse Intubation Protocol
<ol>
	<li>Anesthesia and preparation for intubation
	<ol>
		<li>Wipe the mouse abdomen with an ethanol swab. Anesthetize the mouse with an intraperitoneal injection of tribromoethanol (250-400 mg/kg). Assess the appropriate depth of anesthesia by the lack of pedal withdrawal reflex. Place eye lubricating ointment now or later (step 2.1.4). 
		NOTE: For this procedure, tribromoethanol (and etomidate as an alternative option) provides a stable anesthetic plane without affecting the hemodynamic conditions required for this surgery. This anesthetic is only used once to avoid the risk of peritoneal adhesions. Isoflurane could also be used, but we do not use it here. The practitioner is free to use whatever anesthetic recipe they see fit.</li>
		<li>Place the anesthetized mouse on an intubation stand or plastic support in a supine position, suspended by its upper incisors on looped 4-0 sutures (silk or other) across two support anchors.</li>
		<li>To keep the mouse immobilized during the intubation procedure, loosely tape the lower part of the chest (or both upper limbs) to the platform.</li>
		<li>Place the fiberoptic flexible light gently on the trachea of the mouse, slightly below the vocal cords. Adjust the level of illumination so that only a dark field is visible when looking into the mouse oropharynx except for red light emanating from below the vocal cords, demonstrating the target for the eventual placement of the endotracheal tube. Note that vocal cord movements should be visible with the naked eye or, if needed, under magnification.</li>
	</ol>
	</li>
	<li>Intubation procedure
	<ol>
		<li>Hold the tweezers with the dominant hand and use them to gently grip and draw the tongue out of the oral cavity.</li>
		<li>Open the lower jaw using forceps held by the non-dominant hand, and then push the forceps into the larynx to gently lift the epiglottis. At this time, release the tongue from the tweezers.</li>
		<li>Look for the vocal cords. They should open and close according to each breath. Holding the cannula with the guide wire pre-loaded, insert the tip of the wire through the vocal cords.</li>
		<li>Being very careful not to move the wire by holding a portion of it that is outside the cannula but just above the vocal cords, withdraw the cannula, leaving just the wire in place with its distal end within the trachea.</li>
		<li>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.</li>
		<li>Hold the wire outside the mouth with the curved forceps in the left hand, stabilized against a hard surface, and carefully advance the 20G catheter with tape wings over the wire.</li>
		<li>Once the distal end of the wire emerges from the back end of the 20G catheter or endotracheal tube, hold that end with the curved forceps and smoothly advance the 20G catheter into the trachea.</li>
		<li>Carefully remove the wire from the distal end of the 20G catheter with the curved forceps without dislodging the placement of the catheter.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Post-intubation
	<ol>
		<li>Disconnect the catheter from the ventilator. Fix the tape wings (attached to the catheter) through the lower lip of the mouse using a 4-0 vicryl suture to firmly secure the endotracheal tube (ETT) to the mouse during all subsequent procedures/manipulations. 
		NOTE: Alternatively, silk tape or other tape can be used to secure the ETT, however care should be taken to avoid dislodgement of the ETT during movement of the animal from the intubation sled to the surgical surface.</li>
		<li>Carefully remove the mouse from the intubation sled. Briefly connect the catheter to the ventilator set at a tidal volume 0.2-0.225 mL and a respiratory rate of 120-150 breaths per min to confirm correct tracheal placement of the orotracheal tube and then disconnect with the mouse breathing spontaneously through the orotracheal tube.</li>
		<li>Do not leave the animal unattended from this point onward until it has regained sufficient consciousness to maintain sternal recumbency at the end of the procedure.</li>
	</ol>
	</li>
</ol>
2. Lung ischemia and reperfusion (IR) surgery protocol
<ol>
	<li>Analgesia and preparation of the surgical site
	<ol>
		<li>Wipe the mouse abdomen with an ethanol swab and inject buprenorphine (0.05-0.1 mg/kg) intraperitoneally.</li>
		<li>Shave the hair over the left thorax area up to the left scapula. Remove excess shaved hair using alcohol swabs. 
		NOTE: Steps 2.1.1 and 2.1.2 can also be performed before intubation if there concern for dislodgement of the ETT when secured with silk tape.</li>
		<li>Place the mouse on a warming pad in a left lateral or 3/4 turned position and connect the tracheal tube on the ventilator with a tidal volume of 0.2-0.225 mL (~8 mg/kg) and a respiratory rate of 120-150 breaths per min. Do not use supplemental oxygen for this procedure.</li>
		<li>Apply eye lubricant with a sterile cotton-tip swab. Turn the mouse to 3/4 left side up and immobilize all four limbs and the tail with laboratory tape.</li>
		<li>Disinfect the shaved skin area and surrounding fur with povidone-iodine and wait for the solution to dry. Then cover the surgical field with a sterile drape or clear plastic film and create a rectangular opening in the drape or plastic film for the surgical field.</li>
	</ol>
	</li>
	<li>Surgical procedure
	<ol>
		<li>Confirm the appropriate level of anesthesia (provided by the administration of tribromoethanol and buprenorphine as described earlier) by testing response to toe pinch.</li>
		<li>Using a pair of sharp scissors and a pair of larger forceps (narrow pattern forceps or similar), make a 2 cm transverse skin incision below the inferior angle of the scapula in the left lateral thorax. Use the scissors and a finer pair of forceps (extra fine graefe forceps or similar) to cut into the muscular layer and dissect down to the ribs.</li>
		<li>Identify the second intercostal space and hold the second rib with the extra fine forceps. Pulling the rib upward, use a sterile #11 or #12 (curved) scalpel blade (no handle necessary) to enter the pleural space by separating and cutting across the 2nd-3rd space&#39;s intercostal muscles. Consider pausing ventilation to reduce injury to the left lung apex.</li>
		<li>Insert three sterilized retractors. Use the smallest/narrowest retractor cephalad along the orientation of the ribs, the medium size retractor to the left along the 2nd rib, and the largest retractor to the right along the surface of the 3rd rib.</li>
		<li>Open the chest with slow and progressive retraction using the elastic retractor cords. Expose and identify the left pulmonary artery (PA) by moving the left lung apex away with a sterile cotton-tip swab.</li>
		<li>Use the micro forceps, ultrafine 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.</li>
		<li>Using the PA forceps, pick up the left PA and pull gently but firmly upward and cephalad to visualize the transparent bronchus below. Increase magnification on the dissection microscope (see equipment list for more details) at this point to maximum (2x). 
		NOTE:&#160;Sterilize all equipment before use. Additionally, to maintain sterility, only the tips of surgical instruments should enter the sterile surgical field.</li>
		<li>While retracting the PA away from the bronchus, carefully pass the closed ultrafine 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 bronchus (below).</li>
		<li>Encircle the left PA by tying a slipknot to create an occlusion in the PA. Blood flow interruption is easily visualized under the microscope. This marks the initiation of the ischemic period.</li>
		<li>Externalize the free end of the knot through a different entry point in the anterior left thorax using a 24G-28G needle and secure the end of the suture with a small piece of tape for easier identification later on.</li>
		<li>Reinflate the lung to expel as much air out of the chest cavity as possible using a PEEP valve/tubing on the rodent ventilator. Then, close the ribcage with two interrupted 4-0 nylon sutures.</li>
		<li>Close the muscle and subcutaneous layer with a running 4-0 nylon suture. Then apply two or three drops of topical bupivacaine (0.5%) to the incision. Use a 4-0 nylon suture to close the skin layer with a running suture.</li>
	</ol>
	</li>
	<li>Post-operative care
	<ol>
		<li>When spontaneous ventilation has resumed, disconnect the endotracheal tube from the ventilator and extubate the mouse.</li>
		<li>Place the mouse on the warming pad to maintain body temperature during early post-anesthesia recovery.</li>
		<li>Carefully monitor the mouse while recovering from general anesthesia. Pull the externalized slipknot gently at the end of the ischemic period (30 min or 1 h).</li>
		<li>Move the mouse from the warming pad to a cage once it has exhibited signs of recovery: self-righting and/or movement.</li>
		<li>After the period of reperfusion (1 h or 3 h), euthanize the animal and collect blood by cardiac puncture and lung tissue for further analysis. For 1 h reperfusion, collect plasma for ELISA, tissue for RNA, and protein analysis; for 3 h reperfusion, additionally collect tissue for histology.</li>
	</ol>
	</li>
</ol>

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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><tbody><tr><td>&lt;strong&gt;Equipment&lt;/strong&gt;</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>&lt;strong&gt;Surgery Tools&lt;/strong&gt;</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)&lt;br/&gt; 2. Fixators (Nos. 18200-01)&lt;br/&gt; 3. Retractors (Nos. 18200-05 through 18200-12)&lt;br/&gt; 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>&lt;strong&gt;Reagents&lt;/strong&gt;</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>&lt;strong&gt;Materials&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Alcohol Swab</td><td>BD brand</td><td>&amp;nbsp;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...

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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 ...

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

Research

JoVE Journal

Immunology and Infection

3.6K Views.  University of California San Francisco and San Francisco General Hospital. Beademde longischemie-reperfusiechirurgie kan worden gebruikt om de longspecifieke pathofysiologie van meerdere processen te bestuderen, waaronder longtransplantatie, longembolie en longletsel na hemorragisch trauma met reanimatie. Dit model minimaliseert de inflammatoire bijdragen van atelectase, mechanische ventilatie en hypoxie. Het handhaaft een intact in vivo bloedsomloop immuunsysteem en maakt de studies op langere termijn mogelijk.Dit model zou inzicht kunnen geven in hoe sterie...

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

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

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

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

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

Medicine

A Murine Closed-chest Model of Myocardial Ischemia and Reperfusion

Surgical trauma by thoracotomy in open-chest models of coronary ligation induces an immune response which modifies different mechanisms involved in ischemia and reperfusion. Immune response includes cytokine expression and release or secretion of endogenous ligands of innate immune receptors. Activation of innate immunity can potentially modulate infarct size. We have modified an existing murine closed-chest model using hanging weights which could be useful for studying myocardial pre- and postconditioning and the role of innate immunity in myocardial ischemia and reperfusion. This model allows animals to recover from surgical trauma before onset of myocardial ischemia.
Volatile anesthetics have been intensely studied and their preconditioning effect for the ischemic heart is well known. However, this protective effect precludes its use in open chest models of coronary artery ligation. Thus, another advantage could be the use of the well controllable volatile anesthetics for instrumentation in a chronic closed-chest model, since their preconditioning effect lasts up to 72 hours. Chronic heart diseases with intermittent ischemia and multiple hit models are other possible applications of this model.
For the chronic closed-chest model, intubated and ventilated mice undergo a lateral blunt thoracotomy via the 4th intercostal space. Following identification of the left anterior descending a ligature is passed underneath the vessel and both suture ends are threaded through an occluder. Then, both suture ends are passed through the chest wall, knotted to form a loop and left in the subcutaneous tissue. After chest closure and recovery for 5 days, mice are anesthetized again, chest skin is reopened and hanging weights are hooked up to the loop under ECG control.
At the end of the ischemia/reperfusion protocol, hearts can be stained with TTC for infarct size assessment or undergo perfusion fixation to allow morphometric studies in addition to histology and immunohistochemistry.

Surgical trauma induces an inflammatory response. Cytokines and endogenous ligands are known to modulate myocardial infarct size following ischemia and reperfusion. We present a modified closed-chest model of murine ischemia and reperfusion using hanging weights to minimize effects of thoracotomy.

Surgical trauma by thoracotomy in open-chest models of coronary ligation induces an immune response which modifies different mechanisms involved in ...

A Murine Model of Myocardial Ischemia-reperfusion Injury through Ligation of the Left Anterior Descending Artery

Acute or chronic myocardial infarction (MI) are cardiovascular events resulting in high morbidity and mortality. Establishing the pathological mechanisms at work during MI and developing effective therapeutic approaches requires methodology to reproducibly simulate the clinical incidence and reflect the pathophysiological changes associated with MI. Here, we describe a surgical method to induce MI in mouse models that can be used for short-term ischemia-reperfusion (I/R) injury as well as permanent ligation. The major advantage of this method is to facilitate location of the left anterior descending artery (LAD) to allow for accurate ligation of this artery to induce ischemia in the left ventricle of the mouse heart. Accurate positioning of the ligature on the LAD increases reproducibility of infarct size and thus produces more reliable results. Greater precision in placement of the ligature will improve the standard surgical approaches to simulate MI in mice, thus reducing the number of experimental animals necessary for statistically relevant studies and improving our understanding of the mechanisms producing cardiac dysfunction following MI. This mouse model of MI is also useful for the preclinical testing of treatments targeting myocardial damage following MI.

We introduce a surgical method to induce experimental ischemia/reperfusion (I/R) injury to simulate myocardial infarction (MI) in mouse models that allows for more clarity in positioning of the ligation on the left anterior descending artery (LAD) to increase the reproducibility of MI experiments in mice.

Acute or chronic myocardial infarction (MI) are cardiovascular events resulting in high morbidity and mortality. Establishing the pathological mechanisms ...

Open Tracheostomy Gastric Acid Aspiration Murine Model of Acute Lung Injury Results in Maximal Acute Nonlethal Lung Injury

Acid pneumonitis is a major cause of sterile acute lung injury (ALI) in humans. Acid pneumonitis spans the clinical spectrum from asymptomatic to acute respiratory distress syndrome (ARDS), characterized by neutrophilic alveolitis, and injury to both alveolar epithelium and vascular endothelium. Clinically, ARDS is defined by acute onset of hypoxemia, bilateral patchy pulmonary infiltrates and non-cardiogenic pulmonary edema. Human studies have provided us with valuable information about the physiological and inflammatory changes in the lung caused by ARDS, which has led to various hypotheses about the underling mechanisms. Unfortunately, difficulties determining the etiology of ARDS, as well as a wide range of pathophysiology have resulted in a lack of critical information that could be useful in developing therapeutic strategies.
Translational animal models are valuable when their pathogenesis and pathophysiology accurately reproduce a concept proven in both in vitro and clinical settings. Although large animal models (e.g., sheep) share characteristics of the anatomy of human trachea-bronchial tree, murine models provide a host of other advantages including: low cost; short reproductive cycle lending itself to greater data acquisition; a well understood immunologic system; and a well characterized genome leading to the availability of a variety of gene deletion and transgenic strains. A robust model of low pH induced ARDS requires a murine ALI that targets mainly the alveolar epithelium, secondarily the vascular endothelium, as well as the small airways leading to the alveoli. Furthermore, a reproducible injury with wide differences between different injurious and non-injurious insults is important.
The murine gastric acid aspiration model presented here using hydrochloric acid employs an open tracheostomy and recreates a pathogenic scenario that reproduces the low pH pneumonitis injury in humans. Additionally, this model can be used to examine interaction of a low pH insult with other pulmonary injurious entities (e.g., food particles, pathogenic bacteria).

This protocol induces acute lung injury in a mouse that has close fidelity to the pathogenesis of acid pneumonitis observed in humans. We generate a maximal acute nonlethal low pH lung injury and account for differences in rodent-human anatomic respiratory structure using an open tracheostomy coupled with circumferential pressure release.

Acid pneumonitis is a major cause of sterile acute lung injury (ALI) in humans. Acid pneumonitis spans the clinical spectrum from asymptomatic to acute ...

Two-vessel Occlusion Mouse Model of Cerebral Ischemia-reperfusion

In this study, a middle cerebral artery (MCA) occlusion mouse model is employed to study cerebral ischemia-reperfusion. A reproducible and reliable mouse model is useful for investigating the pathophysiology of cerebral ischemia-reperfusion and determining potential therapeutic strategies for patients with stroke. Variations in the anatomy of the circle of Willis of C57BL/6 mice affects their infarct volume after cerebral-ischemia-induced injury. Studies have indicated that distal MCA occlusion (MCAO) can overcome this problem and result in a stable infarct size. In this study, we establish a two-vessel occlusion mouse model of cerebral ischemia-reperfusion through the interruption of the blood flow to the right MCA. We distally ligate the right MCA and right common carotid artery (CCA) and restore blood flow after a certain period of ischemia. This ischemia-reperfusion injury induces an infarct of stable size and a behavioral deficit. Peripheral immune cells infiltrate the ischemic brain within the 24 h infiltration period. Additionally, the neuronal loss in the cortical area is less for a longer reperfusion duration. Therefore, this two-vessel occlusion model is suitable for investigating the immune response and neuronal recovery during the reperfusion period after cerebral ischemia.

A mouse model of cerebral ischemia-reperfusion is established to investigate the pathophysiology of stroke. We distally ligate the right middle cerebral artery and right common carotid artery and restore blood flow after 10 or 40 min of ischemia.

In this study, a middle cerebral artery (MCA) occlusion mouse model is employed to study cerebral ischemia-reperfusion. A reproducible and reliable mouse ...

Murine Myocardial Infarction Model using Permanent Ligation of Left Anterior Descending Coronary Artery

Myocardial infarction (MI) and acute coronary diseases are among the most prominent causes of death in population with western lifestyle. The murine models of MI with permanent ligation of left-anterior descending (LAD) coronary artery closely mimics MI in humans. Murine models benefit from the extensive genetic engineering available nowadays. Here we propose a reproducible murine surgical model of myocardial infarction by permanent LAD coronary ligation. Our technique comprises anesthesia with ketamine/xylazine that can be rapidly reversed by administration of an antagonist, intubation without tracheotomy for mechanical-assisted ventilation, ventilation with application of extrinsic positive end-expiratory pressure (PEEP) to avoid alveolar collapse, a thoracotomy method limiting to the minimum surgical lesions made to skeletal muscles, and lung inflation without thoracentesis. This method is sparsely invasive, reproducible and reduces post-surgery mortality and complications.

Herein we describe a surgical procedure showing how to achieve permanent ligation of the left-anterior descending coronary artery in mice. This model is of high relevance to investigate the pathophysiology of myocardial infarction and the concomitant biological processes.

Myocardial infarction (MI) and acute coronary diseases are among the most prominent causes of death in population with western lifestyle. The murine ...

Murine Surgical Model of Topical Elastase Induced Descending Thoracic Aortic Aneurysm

According to the Center for Disease Control, aortic aneurysms (AAs) were considered a leading cause of death in all races and both sexes from 1999-2016. An aneurysm forms as a result of progressive weakening and eventual dilation of the aorta, which can rupture or tear once it reaches a critical diameter. Aneurysms of the descending aorta in the chest, called descending thoracic aortic aneurysms (dTAA), make up a large proportion of aneurysm cases in the United States. Uncontained dTAA rupture is almost universally lethal, and elective repair has a high rate of morbidity and mortality. The purpose of our model is to study dTAA specifically, to elucidate the pathophysiology of dTAA and to search for molecular targets to halt the growth or reduce the size of dTAA. By having a murine model to study thoracic pathology precisely, targeted therapies can be developed to specifically test dTAA. The method is based on the placement of porcine pancreatic elastase (PPE) directly on the outer murine aortic wall after surgical exposure. This creates a destructive and inflammatory reaction, which weakens the aortic wall and allows for aneurysm formation over weeks to months. Though murine models possess limitations, our dTAA model produces robust aneurysms of predictable size. Furthermore, this model can be used to test genetic and pharmaceutical targets which may arrest dTAA growth or prevent rupture. In human patients, interventions such as these could help avoid aneurysm rupture, and difficult surgical intervention.

We describe a surgical protocol to consistently induce robust descending thoracic aortic aneurysms in mice. The procedure involves left thoracotomy, thoracic aorta exposure, and placement of a sponge soaked in porcine pancreatic elastase on the aortic wall.

According to the Center for Disease Control, aortic aneurysms (AAs) were considered a leading cause of death in all races and both sexes from 1999-2016. ...

Improved Rodent Model of Myocardial Ischemia and Reperfusion Injury

Myocardial ischemia and reperfusion injury (MIRI), induced by coronary heart disease (CHD), causes damage to the cardiomyocytes. Furthermore, evidence suggests that thrombolytic therapy or primary percutaneous coronary intervention (PPCI) does not prevent reperfusion injury. There is still no ideal animal model for MIRI. This study aims to improve the MIRI model in rats to make surgery easier and more feasible. A unique method for establishing MIRI is developed by using a soft tube during a key step of the ischemic period. To explore this method, thirty rats were randomly divided into three groups: sham group (n = 10); experimental model group (n = 10); and existing model group (n = 10). Findings of triphenyltetrazolium chloride staining, electrocardiography, and percent survival are compared to determine the accuracies and survival rates of the operations. Based on the study results, it has been concluded that the improved surgery method is associated with a higher survival rate, elevated ST-T segment, and larger infarct size, which is expected to mimic the pathology of MIRI better.

Myocardial ischemia-reperfusion model of rat heart is improved by using a self-made retractor, polyvinyl chloride tube, and a unique knotting method. Electrocardiogram, triphenyltetrazolium chloride and histological staining, and percent survival analysis results showed that the improved model group has higher success and survival rates than the already existing model group.

Myocardial ischemia and reperfusion injury (MIRI), induced by coronary heart disease (CHD), causes damage to the cardiomyocytes. Furthermore, evidence ...

Left Coronary Artery Ligation: A Surgical Murine Model of Myocardial Infarction

Ischemic heart disease and subsequent myocardial infarction (MI) is one of the leading causes of mortality in the United States and around the world. In order to explore the pathophysiological changes after myocardial infarction and design future treatments, research models of MI are required. Permanent ligation of the left coronary artery (LCA) in mice is a popular model to investigate cardiac function and ventricular remodeling post MI. Here we describe a less invasive, reliable, and reproducible surgical murine MI model by permanent ligation of the LCA. Our surgical model comprises of an easily reversible general anesthesia, endotracheal intubation that does not require a tracheotomy, and a thoracotomy. Electrocardiography and troponin measurement should be performed to ensure MI. Echocardiography at day 28 after MI will discern heart function and heart failure parameters. The degree of cardiac fibrosis can be evaluated by Masson's trichrome staining and cardiac MRI. This MI model is useful for studying the pathophysiological and immunological alterations after MI.

Presented here is a surgical procedure for permanent ligation of the left coronary artery in mice. This model can be used to investigate the pathophysiology and associated inflammatory response after myocardial infarction.

Ischemic heart disease and subsequent myocardial infarction (MI) is one of the leading causes of mortality in the United States and around the world. In ...

Mouse Model for Studying Lung Ischemia-Reperfusion Injury

Murine Left Pulmonary Hilar Clamp Model of Lung Ischemia Reperfusion Injury

Ischemia reperfusion injury (IRI) during lung transplantation is a major risk factor for post-transplant complications, including primary graft dysfunction, acute and chronic rejection, and mortality. Efforts to study the underpinnings of IRI led to the development of a reliable and reproducible mouse model of left lung hilar clamping. This model involves a surgical procedure performed in an anesthetized and intubated mouse. A left thoracotomy is performed, followed by careful lung mobilization and dissection of the left pulmonary hilum. The hilar clamp involves reversible suture ligation of the pulmonary hilum with a slipknot, which stops the arterial inflow, venous outflow, and airflow through the left mainstem bronchus. Reperfusion is initiated by careful removal of the suture. Our laboratory uses 30 min of ischemia and 1 h of reperfusion for the experimental model in the current investigations. However, these time periods can be modified depending on the specific experimental question. Immediately prior to sacrifice, arterial blood gas can be obtained from the left ventricle after a 4 min period of right hilar clamping to ensure that the PaO2 values obtained are attributed to the injured left lung alone. We also describe a method to measure cell extravasation with flow cytometry, which involves intravenous injection of a fluorochrome-labeled antibody specific for the cell(s) to be studied prior to sacrifice. The left lung can then be harvested for flow cytometry, frozen or fixed, paraffin-embedded immunohistochemistry, and quantitative polymerase chain reaction. This hilar clamp technique allows for&#160;detailed study of the cellular and molecular mechanisms underlying IRI. Representative results reveal decreased left lung oxygenation and histologic evidence of lung injury following hilar clamping. This technique can be readily learned and reproduced by personnel with and without microsurgical experience, leading to reliable and consistent results and serving as a widely adoptable model for studying lung IRI.

Author Spotlight: Insights into the Effect of Ischemia Reperfusion on Lung Transplantation

The protocol outlines a feasible, reliable, and reproducible method of left pulmonary hilar clamping that can be used to study lung ischemia-reperfusion injury in mouse models.

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

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