This work was supported by Merit Review awards (BX002332 and BX000640) from the Biomedical Laboratory Research and Development Service of the Veterans Affairs Office of Research and Development, National Institutes of Health (R15HL129140), and funds from Institutional Research and Improvement account. The project is supported in part by the National Institutes of Health grant C06RR0306551.

All surgical procedures performed on animals should be carried out in accordance with Guide for the Care and Use of Laboratory Animals13 or other appropriate ethical guidelines. Protocols should be approved by the animal welfare committee at the appropriate institution before proceeding.
1. Preparing for the ECG
NOTE: Before beginning, don personal protective equipment including gloves, eyewear and a clean laboratory coat or disposable gown.
<ol>
	<li>Clean the ECG pad using a non-alcohol and non-bleach based decontamination solution. Gently use a delicate task wipe to blot off excess solution to ensure that the electrode pad does not become damaged.</li>
	<li>If the ECG pad has a heating feature, use it. Anesthetized mice tend to lose body heat rapidly. Heat the pad to 40-42 &#176;C to maintain normothermic body temperature of 37 &#176;C throughout the surgery14. Monitor body temperature using a rectal thermometer. Adjust pad temperature as necessary to maintain the body temperature ~37 &#176;C.</li>
	<li>As most thoracotomies are performed with the mouse lying on its back (supine), ensure that the toggle is flipped to the &#34;supine&#34; setting. Many ECG pads have a function to toggle between prone and supine positions. Failure to select the right orientation can result in misrepresentation of electrocardiographic events.</li>
	<li>Anesthetize mouse using 5% inhaled isoflurane and 1 L/min oxygen. Once mouse is anesthetized, transfer mouse to ECG pad equipped with an anesthesia nose-cone and reduce isoflurane to 2% and 1 L/min oxygen. Confirm proper anesthesia by ensuring mouse does not react when the mouse&#39;s foot is pinched with forceps.</li>
	<li>Apply a thin coat of eye lubrication ointment over the mouse&#39;s eyes to prevent dryness and corneal damage while anesthetized.</li>
	<li>Clean the mouse&#39;s paws with a wet wipe to remove all visible bedding that may be stuck to the paws or may interfere with the transmission of electrical impulses from the paws to the ECG pad. Dry paws with a wipe.</li>
	<li>Apply a small amount (slightly smaller than a USD dime) of highly conductive electrolyte gel to each of the four metallic electrodes on the ECG pad. 
	Note: Be sure to only apply a small amount of gel as too much gel makes it difficult to restrain the paws to the pad using tape. Additionally, paws are likely to slip out of the restraint during surgery if they were wet before applying tape.</li>
	<li>With the mouse in supine position, use clear medical tape to restrain each paw to its corresponding electrode (Figure 1). First press each paw to its piece of tape and then adhere the tape to the ECG pad. Ensure that each restrained paw is in contact with the electrolyte gel and the electrode.</li>
</ol>
2. Acquiring the ECG
<ol>
	<li>Depending on the equipment used for ECG acquisition, configure the machine so that the ECG waveform can be visualized in real time. For ECG recordings using the physiologic monitoring settings on an echocardiography machine, a live B-mode image will have the ECG waveform running along the bottom of the screen. 
	Note: See individual machine user guides to determine how best to configure that equipment.</li>
	<li>Enable real time visualization of ECG waveform by pressing the B-mode key on an echocardiography machine or the equivalent on other ECG recording devices.
	<ol>
		<li>Adjust the resolution to account for differences in amplitude. If the peak of the R-wave or the trough (valley) of the Q-wave are out of the visual frame, adjust the resolution until the entire height of the waveform can be observed. 
		Note: This can be done under the physiological settings tab on an echocardiography machine. Click the increase or decrease arrows until the entire waveform is visible.</li>
	</ol>
	</li>
	<li>Any time that an image is to be obtained, clear the ECG pad of tools. Touching the mouse during ECG recording with forceps or fingers will disturb the waveform. Ensure that the mouse is still and untouched on the ECG pad before recording any ECGs.</li>
	<li>Use the machine&#39;s &#34;record&#34; or &#34;store&#34; feature before making any surgical incisions on the mouse. This image will be used as a baseline for comparison later on.</li>
</ol>
3. Surgical Procedure and Recording ECG
<ol>
	<li>Inject anesthetized mouse with analgesic (buprenorphine, 1.5 &#181;g, intraperitoneal) before beginning. The details of the ischemia/reperfusion surgical procedure can also be found elsewhere5.</li>
	<li>Remove hair around the surgical site chemically or mechanically and disinfect the area with betadine solution. Use a scalpel to make a vertical incision parallel to the esophagus and trachea. Gently move the lymph nodes to each side of the incision until the thin tissue covering the trachea is exposed. Using forceps, gently separate the tissue until the white cartilage rings of the trachea are visible. 
	Note: We generally use Nair hair removal lotion. The lotion is applied on the surgical site for ~1 min. Nair is then thoroughly washed off using saline or water. This method is preferred in our laboratory because high resolution echocardiography (which can detect hair follicles) is performed before and after surgery. However, caution should be taken to avoid sensitive areas such as genitals, and then thoroughly washing the Nair off to avoid potential skin burns.</li>
	<li>Quickly remove the mouse&#39;s nose from the nose-cone and insert ventilation tubing into the mouth of the mouse and towards the throat. When the tip of the ventilation tubing is visible through the exposed neck area, align the tube with the start of the trachea. Gently wiggle the tube side to side while applying upward pressure until the tubing slides into the trachea which can be confirmed visually through the translucent trachea.</li>
	<li>Ensure that the mouse remains anesthetized during the intubation procedure. Pause from intubation and return the mouse to the nose-cone if it begins to stir.</li>
	<li>Using a loop of string, hook the mouse&#39;s two front teeth through the loop and tape the string ends to the ECG pad to steady the head of the mouse and to ensure the ventilation tubing does not move during surgery. Quickly attach ventilation tubing to rodent ventilator and adjust ventilation settings according to the weight of the mouse. Tape ventilation tubing in place.</li>
	<li>Cover the mouse&#39;s exposed trachea with a gauze soaked in warm saline to keep tissue from drying.</li>
	<li>Make a vertical incision using a scalpel along the left side of the sternum.</li>
	<li>Using forceps, gently separate the fascia layer from the muscle layer. Carefully cut the underlying muscle layers without cutting visible blood vessels.</li>
	<li>Using forceps, grab the third rib and pull upwards gently. Maintain grip on the rib with one hand and use surgical scissors to carefully cut the intercostal tissue between the third and fourth rib. Ensure that the lungs are not damaged. 
	Note: Lungs will retract deep into the chest cavity almost immediately after the chest cavity is punctured by the surgical incision due to the loss of the pressure gradient. Wait until the lungs have retracted before continuing.</li>
	<li>Use forceps to grab and gently separate the thin layer of pericardium which surrounds the heart.</li>
	<li>Insert retractors or manually use forceps as rib retractors to move the ribs into a position where the heart is visible between the ribs. 
	Note: It is common practice to move the mouse&#39;s lower left paw so that it is overlapping the lower right paw during the placement of the ligature. This helps to position the heart so that the left atrial appendage, or auricle, is easily visible during placement of the ligature. Be aware that valid ECG waveforms will not be obtained while the lower left paw is off of the electrode. For this reason it is advisable to return the paw to its electrode after the suturing ligature is passed through the myocardial tissue but before a knot is tightened.</li>
	<li>Locate the LAD visually, beneath the left auricle. Swiftly insert a 7-0 silk tapered suturing needle into the myocardium deeply enough to pass under the LAD but not so deep as to penetrate the LV cavity. Pull the suturing ligature through until there is about 4 cm of suturing silk left on the free (non-needle) end of the suturing ligature.</li>
	<li>Begin to tie a simple suture knot. Once the free end of the suturing silk has been pulled through the loops to form the knot, pause.</li>
	<li>Holding both the free and needle ends of the suturing silk with forceps, insert a ~1 cm section of PE-10 tubing underneath the forming knot and atop the epicardial surface.</li>
	<li>If the mouse&#39;s left paw is crossed, return the paw to its proper electrode. Tighten knot so that the PE-10 tubing is sutured to the heart. Release all physical contact with the mouse to allow ECG to be recorded.</li>
	<li>Allow ECG waveform to cycle through for ~10 sec. Check ECG waveform visually and record waveform as &#34;Time of Occlusion&#34;. If the T-wave does not increase in amplitude within 1 min, reassess placement of ligature.
	<ol>
		<li>If the T-wave amplitude does not increase, either discard the animal from the study or attempt to correct the ligature placement.</li>
	</ol>
	</li>
	<li>Visually check the color of the myocardium to confirm ischemic paling of the LV.</li>
	<li>If ECG changes and myocardial color changes indicate ischemia, double knot the suture around the PE-10 tubing.</li>
	<li>Cover the open chest cavity with warm saline gauze.</li>
	<li>Record ECG every 5-10 min for the duration of the ischemic period.</li>
</ol>
4. Confirmation of Reperfusion Using ECG
<ol>
	<li>Remove saline gauze covering the chest cavity and visualize the heart.</li>
	<li>Use a blade to cut the suturing silk atop the PE-10 tubing. Once the ligature is cut, remove the section of PE-10 tubing and gently remove the suturing ligature from the myocardium.</li>
	<li>Release all physical contact with the mouse and allow the ECG waveform ~10 sec to cycle. Record waveform as &#34;Time of Reperfusion.&#34; Continue to record ECG waveforms every 5-10 min until the desired experimental time point is reached.</li>
	<li>Adjust resolution for changes in amplitude as needed. If the T-wave does not change upon removal of the PE-10 tubing and ligature, reperfusion is not confirmed.
	<ol>
		<li>If the T-wave does not change upon removal of tubing, either discard the animal from the study or attempt to correct the ligature placement.</li>
	</ol>
	</li>
	<li>Visually inspect myocardium to additionally confirm reperfusion by return to red color.</li>
	<li>Close chest cavity by suturing the intercostal space with a 5-0 silk suture while applying gentle pressure to the mouse&#39;s chest to expel excess air that has entered during surgery. Then suture the muscle layers and finally, skin. Note: Applying pressure to the chest cavity may not be sufficient to evacuate the chest cavity of air in all mice.&#160; Therefore, the syringe and needle method of evacuation should be employed to ensure that all air has been expelled.</li>
	<li>Record the last ECG before turning inhaled anesthesia off and removing the mouse&#39;s paws from the electrodes. Increase oxygen to 2 L/min and maintain ventilation until the mouse regains consciousness.</li>
	<li>Allow mouse to recover in a constant temperature controlled environment, e.g. heating pad or warm incubator, to avoid infarct variability. Treat mouse with buprenorphine 24 hr after the surgery and then as needed as indicated by the mouse grimace scale. 
	Note: Procedure for reperfusion is also discussed in detail by Xu et al.5</li>
</ol>

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The authors have nothing to disclose.

Using ECG changes as a supplemental method for confirming myocardial ischemia and reperfusion ensures the accurate placement of the occluding ligature. Accuracy of ligature placement is critical to reducing data variability among animals. The LAD in a mouse heart is a difficult artery to visualize. Therefore, supplementing visual pallor with electrocardiographic changes will help ensure the correct placement of the ligature and resulting tissue damage.
Since the ECG pad offers a non-invasive v...

Heart disease remains the leading cause of death worldwide1,2. Not only is the left ventricle (LV) the most muscular chamber, responsible for pumping blood from the heart to the entire body3, it is a common cardiac injury site post-myocardial infarction4. Left ventricular tissue death often results in systolic heart failure. Animal models of heart disease are imperative for the advancement of biomedical cardiovascular research. The C57Bl/6 strain of mice have been a popular choice for animal models due to their quick breeding time, low cost and ease in genetic alterations. Most murine surgical models for the study of heart disease involve occlusion of the LAD branch of the left coronary artery. The LAD is sometimes called the left obtuse marginal5,6. The LAD supplies blood to the left ventricular anterior and antero-lateral walls. LAD occlusion studies are aimed at inducing anterior infarctions, sometimes extending into the inferior and lateral wall regions7.
Two models that are used frequently for myocardial infarction studies include chronic occlusion myocardial infarction and myocardial ischemia/reperfusion injury. The chronic occlusion is created by surgically suturing around and permanently blocking blood flow through the LAD. The ischemia/reperfusion injury is created much in the same way only with a transient, usually 30-60 min, ischemic period. To achieve transient ischemia, the occluding suture ties around the LAD and a small PE-10 tube which is placed parallel to the LAD on the epicardial surface of the heart, followed by a reperfusion period where the tubing and occluding suture is removed and blood is allowed to once again flow through the artery and into the myocardium. The ischemia/reperfusion surgery has been deemed to be clinically relevant due to the nature of reperfusion injury paralleling the treatment of human infarctions which includes prompt coronary angioplasty and stenting of the artery, or coronary artery bypass. Typically, during these surgeries, ischemia of the LV in a mouse heart is confirmed by visible pallor of the myocardial wall. However, by simply performing the surgeries on an electrocardiogram (ECG) pad under constant monitoring conditions, visible changes can be observed in the ECG waveform, thereby confirming ischemia and reperfusion of the mouse myocardium.
Although the murine heart is similar to the human heart in many respects, including its four-chambered structure, the hearts also have differences. One obvious difference is the average resting heart rate of adult mice is 600 - 700 beats per min (bpm) whereas that of adult humans is ~60-100 bpm8,9. Additionally, in mice the repolarization waves, J and T, often merge with the depolarization QRS-complex making a clear ST-segment difficult to discern10. To complicate the process of electrocardiographically confirming myocardial ischemia, it is the elevation of the T-wave and the ST-segment which are used as markers for the diagnosis of ischemia and myocardial infarction injury in humans, clinically referred to as ST elevation myocardial infarction or STEMI. One of the key differences between human and murine waveforms is that S-wave is immediately followed be a J-wave that transfers directly into a negative T-wave. During acute myocardial ischemia in mice the amplitude of S-wave decreases and is directly followed by an abnormal J-wave and an inverted T-wave11. The T-wave does not seem to represent a significant portion of the repolarization in mice11. Despite nomenclature and mouse vs. human differences, ECG confirmation of murine myocardial ischemia and reperfusion is still feasible and relatively simple. For the sake of simplifying waveform interpretation, the segment between the S-J-T is referred to as ST-segment herein.
STEMI guidelines published in 2013 recommend a patient door-to-balloon time of less than 90 min12.This means that the time frame from the identification of the patient&#39;s coronary artery occlusion until the artery is reopened should be less than 90 min. The beating heart is constantly working and therefore, has a high oxidative metabolism and a high level of oxygen consumption3. To provide for this, a network of capillaries is available to each myocyte3. It only takes a heart a few beats to exhaust its oxygen and nutrient supply. In a 90 min window, an ischemic heart region in a human will have been blocked from receiving between 5,400 and 9,000 heart beats worth of oxygen-rich blood. In that same 90 min window, a mouse would have 54,000 to 63,000 heart beats. Experimental time points for murine ischemia/reperfusion injury are typically between 30 and 60 min.
The importance of developing a supplemental method of confirming myocardial ischemia and reperfusion in a murine model has profound implications on the consistency and reproducibility of data in myocardial ischemia/reperfusion studies. The current practice of visually observing the heart for a change in tissue color is not adequate as a stand-alone diagnostic. Additionally, reperfusion after removal of the tubing and suture is not guaranteed. Although the artery is no longer tied off, the artery may have sustained damage during the procedure and may become impossible to reperfuse. It would be beneficial to have a record of electrocardiographic changes to confirm reperfusion rather than relying on observations of myocardial pallor and rubor (red color). Hearts that do not show the markers of ischemia/reperfusion injury can then quickly be flagged and a decision on how to proceed can be made by the investigators.
Lastly, establishing a record of ECG changes from baseline throughout the ischemic and reperfusion periods allows investigators to continue to monitor the heart after the initial surgery. Investigators currently lose sight of the heart as soon as the surgery is completed. ECG is a simple way to gain insight into changes occurring in the myocardium hours to days after the surgery. ECG recorded at time points after surgery could reveal late-developing Q-waves indicating continued or worsening tissue death. However, to effectively gage new or worsening electrocardiographic markers, a baseline ECG must be available for comparison.
This protocol will demonstrate how to prepare, obtain, and interpret the ECG to confirm ischemia and reperfusion of the mouse heart using 8 - 12 week old male C57Bl/6 mice.

<table><tbody><tr><td>Vevo 1100</td><td>Fujifilm&lt;br/&gt; Visual Sonics</td><td></td><td>Echocardiography Machine</td></tr><tr><td>Mouse Handling Plate</td><td>Fujifilm&lt;br/&gt; Visual Sonics</td><td></td><td>Heated ECG plate</td></tr><tr><td>Signa-Gel&amp;nbsp;</td><td></td><td></td><td>Highly Conductive Multi-</td></tr><tr><td>Electrode Gel</td><td>Parker</td><td>15-25</td><td>Purpose Electrolyte</td></tr><tr><td>Transpore Medical Tape</td><td>3M</td><td>1527-0</td><td></td></tr><tr><td>PI-Spray II</td><td>Pharmaceutical Innovations</td><td>NDC 36-2013-25</td><td>Cleaning agent for ECG plate</td></tr><tr><td>C57Bl6 Mice</td><td>The Jackson Laboratory</td><td>000664</td><td>Male, 8 - 12 wk</td></tr><tr><td>IsoThesia-Isoflurane</td><td>Henry Schein</td><td>NDC 1169-0500-1</td><td></td></tr><tr><td>Excel</td><td>Microsoft</td><td></td><td></td></tr><tr><td>Systane Nighttime Lubricant Eye Ointment</td><td>Alcon</td><td>65050935</td><td></td></tr><tr><td>7-0 Perma-Hand Silk Sutures</td><td>Ethicon</td><td>640.O32</td><td></td></tr><tr><td>5-0 Perma-Hand Silk Sutures</td><td>Ethicon</td><td>K809.O32</td><td></td></tr><tr><td>Surgical Scissors</td><td>ROBOZ</td><td>RS-5881</td><td></td></tr><tr><td>Forceps</td><td>Fine Science Tools</td><td>11052-10</td><td></td></tr><tr><td>Gauze</td><td>Bio Nuclear Diagnostics Inc</td><td>DIS-022B</td><td></td></tr><tr><td>Needle Holder</td><td>Fine Science Tools</td><td>12565-14</td><td></td></tr><tr><td>Buprenex CIII&amp;nbsp;</td><td>Patterson Veterinary</td><td>0-891-9756</td><td>Buprenorphine Hydrochloride Analgesic&amp;nbsp;</td></tr><tr><td>Betadine</td><td>Purdue Products</td><td>67618-150-08</td><td></td></tr><tr><td>Nair</td><td>Church and Dwight Co.</td><td>NRSL-22339-05</td><td></td></tr></tbody></table>

confirmation of myocardial ischemia and reperfusion injury in mice using surface pad electrocardiography

Many animal models have been established for the study of myocardial remodeling and heart failure due to its status as the number one cause of mortality worldwide. In humans, a pathologic occlusion forms in a coronary artery and reperfusion of that occluded artery is considered essential to maintain viability of the myocardium at risk. Although essential for myocardial recovery, reperfusion of the ischemic myocardium creates its own tissue injury. The physiologic response and healing of an ischemia/reperfusion injury is different from a chronic occlusion injury. Myocardial ischemia/reperfusion injury is gaining recognition as a clinically relevant model for myocardial infarction studies. For this reason, parallel animal models of ischemia/reperfusion are vital in advancing the knowledge base regarding myocardial injury. Typically, ischemia of the mouse heart after left anterior descending (LAD) coronary artery occlusion is confirmed by visible pallor of the myocardium below the occlusion (ligature). However, this offers only a subjective way of confirming correct or consistent ligature placement, as there are multiple major arteries that could cause pallor in different myocardial regions. A method of recording electrocardiographic changes to assess correct ligature placement and resultant ischemia as well as reperfusion, to supplement observed myocardial pallor, would help yield consistent infarct sizes in mouse models. In turn, this would help decrease the number of mice used. Additionally, electrocardiographic changes can continue to be recorded non-invasively in a time-dependent fashion after the surgery. This article will demonstrate a method of electrocardiographically confirming myocardial ischemia and reperfusion in real time.

A normal murine ECG is displayed in Figure 2 with alphabetic markers for electrical events P, Q, R, S, J and T. P is the initial atrial depolarization. QRS is the wave of depolarization over the ventricles. J is early repolarization and T represents heterogeneous repolarization also known as recovery11. It should be noted that many labs do not use the J-wave nomenclature and instead refer to the SJT-segment as the ST-segment10,15-17. Here, results an...

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Confirmation of Myocardial Ischemia and Reperfusion Injury in Mice Using Surface Pad Electrocardiography

During murine myocardial ischemia/reperfusion surgery, correct placement of the occluding ligature is typically confirmed by visible observation of myocardial pallor. Herein, a method of electrocardiographically confirming ischemia and reperfusion, to supplement observed myocardial pallor, is demonstrated in male C57Bl/6 mice.

Many animal models have been established for the study of myocardial remodeling and heart failure due to its status as the number one cause of mortality ...

confirmation-myocardial-ischemia-reperfusion-injury-mice-using

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JoVE Journal

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13.2K Views.  East Tennessee State University. During murine myocardial ischemia/reperfusion surgery, correct placement of the occluding ligature is typically confirmed by visible observation of myocardial pallor. Herein, a method of electrocardiographically confirming ischemia and reperfusion, to supplement observed myocardial pallor, is demonstrated in male C57Bl/6 mice.

Video: Confirmation of Myocardial Ischemia and Reperfusion Injury in Mice Using Surface Pad Electrocardiography

Programmed Electrical Stimulation in Mice

Genetically-modified mice have emerged as a preferable animal model to study the molecular mechanisms underlying conduction abnormalities, atrial and ventricular arrhythmias, and sudden cardiac death.1 Intracardiac pacing studies can be performed in mice using a 1.1F octapolar catheter inserted into the jugular vein, and advanced into the right atrium and ventricle. Here, we illustrate the steps involved in performing programmed electrical stimulation in mice. Surface ECG and intracardiac electrograms are recorded simultaneously in the atria, atrioventricular junction, and ventricular myocardium, whereas intracardiac pacing of the atrium is performed using an external stimulator. Thus, programmed electrical stimulation in mice provides unique opportunities to explore molecular mechanisms underlying conduction defects and cardiac arrhythmias.

Programmed electrical stimulation provides the ability to determine conduction properties of the heart, and the possibility to induce and terminate cardiac arrhythmias using various pacing protocols. Using a transvenous catheter, intracardiac electrogram recordings can be obtained in mice following programmed electrical stimulation protocols to identify arrhythmogenic substrates.

Genetically-modified mice have emerged as a preferable animal model to study the molecular mechanisms underlying conduction abnormalities, atrial and ...

High-frequency High-resolution Echocardiography: First Evidence on Non-invasive Repeated Measure of Myocardial Strain, Contractility, and Mitral Regurgitation in the Ischemia-reperfused Murine Heart

Ischemia-reperfusion (IR) was surgically performed in murine hearts which were then subjected to repeated imaging to monitor temporal changes in functional parameters of key clinical significance. Two-dimensional movies were acquired at high frame rate (8 kHz) and were utilized to estimate high-quality myocardial strain. Two-dimensional elastograms (strain images), as well as strain profiles, were visualized. Results were powerful in quantitatively assessing IR-induced changes in cardiac events including left-ventricular (LV) contraction, LV relaxation and isovolumetric phases of both pre-IR and post-IR beating hearts in intact mice. In addition, compromised sector-wise wall motion and anatomical deformation in the infarcted myocardium were visualized.  The elastograms were uniquely able to provide information on the following parameters in addition to standard physiological indices that are known to be affected by myocardial infarction in the mouse: internal diameters of mitral valve orifice and aorta, effective regurgitant orifice, myocardial strain (circumferential as well as radial), turbulence in blood flow pattern as revealed by the color Doppler movies and velocity profiles, asynchrony in LV sector, and changes in the length and direction of vectors demonstrating slower and asymmetrical wall movement. This work emphasizes on the visual demonstration of how such analyses are performed. 

High-frequency High-resolution Echocardiography: First Evidence on Non-invasive Repeated Measure of Myocardial Strain, Contractility, and Mitral Regurgitation in the Ischemia-reperfused Murine Heart 

High frequency Doppler ultrasound is a novel technology for assessing regional myocardial function. This work presents first evidence demonstrating applicability of this versatile imaging platform for the repeated measure of myocardial strain, dp/dt, and mitral regurgitation in the ischemia-reperfused (IR) murine heart.

Ischemia-reperfusion (IR) was surgically performed in murine hearts which were then subjected to repeated imaging to monitor temporal changes in ...

Modified Technique for Coronary Artery Ligation in Mice

Myocardial infarction (MI) is one of the most important causes of mortality in humans1-3. In order to improve morbidity and mortality in patients with MI we need better knowledge about pathophysiology of myocardial ischemia. This knowledge may be valuable to define new therapeutic targets for innovative cardiovascular therapies4. Experimental MI model in mice is an increasingly popular small-animal model in preclinical research in which MI is induced by means of permanent or temporary ligation of left coronary artery (LCA)5. In this video, we describe the step-by-step method of how to induce experimental MI in mice.
The animal is first anesthetized with 2% isoflurane. The unconscious mouse is then intubated and connected to a ventilator for artificial ventilation. The left chest is shaved and 1.5 cm incision along mid-axillary line is made in the skin. The left pectoralis major muscle is bluntly dissociated until the ribs are exposed. The muscle layers are pulled aside and fixed with an eyelid-retractor. After these preparations, left thoracotomy is performed between the third and fourth ribs in order to visualize the anterior surface of the heart and left lung. The proximal segment of LCA artery is then ligated with a 7-0 ethilon suture which typically induces an infarct size ~40% of left ventricle. At the end, the chest is closed and the animals receive postoperative analgesia (Temgesic, 0.3 mg/50 ml, ip). The animals are kept in a warm cage until spontaneous recovery.

The surgical procedure used to induce experimental myocardial infarction in mice begins with left thoracotomy between the third and the fourth ribs in order to visualize the anterior surface of the heart and left lung. The left coronary artery is ligated, the chest is closed and the mouse is allowed to recover spontaneously.

Myocardial infarction (MI) is one of the most important causes of mortality in humans1-3. In order to improve morbidity and mortality in patients with MI ...

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 ...

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 ...

A Modified Simple Method for Induction of Myocardial Infarction in Mice

Myocardial infarction (MI) represents one of the leading causes of death. MI models are widely used for investigating the pathomechanisms of post-MI remodeling and evaluation of novel therapeutics. Different methods (e.g., isoproterenol treatment, cryoinjury, coronary artery ligation, etc.) have been used to induce MI. Compared with isoproterenol treatment and cryoinjury, coronary artery ligation may better reflect the ischemic response and chronic remodeling after MI. However, traditional methods for coronary ligation in mice are technically challenging. The current study describes a simple and efficient process for induction of MI in mice with readily available materials. The mouse chest skin was cut open under stable anesthesia. The heart was immediately externalized through the intercostal space after blunt separation of the pectoralis major and pectoralis minor. The left anterior descending branch (LAD) was ligated with a 6-0 suture 3 mm from its origin. Following LAD ligation, staining with 2,3,5-Triphenyltetrazolium chloride (TTC) indicated successful induction of MI and temporal changes of post-MI scar size. Meanwhile, survival analysis results showed overt mortality within 7 days after MI, mainly due to cardiac rupture. Moreover, post-MI echocardiographic assessment demonstrated successful induction of contractile dysfunction and ventricular remodeling. Once mastered, an MI model can be established in mice within 2-3 min with readily available materials.

Under adequate anesthesia, the mouse heart was externalized through the intercostal space, and myocardial infarction was successfully induced by ligating the left anterior descending artery (LAD) using materials readily available in most laboratories.

Myocardial infarction (MI) represents one of the leading causes of death. MI models are widely used for investigating the pathomechanisms of post-MI ...

Induction of Myocardial Infarction and Myocardial Ischemia-Reperfusion Injury in Mice

Acute myocardial infarction is a common cardiovascular disease with high mortality. Myocardial reperfusion injury can counteract the beneficial effects of heart reflow and induce secondary myocardial injury. A simple and reproducible model of myocardial infarction and myocardial ischemia-reperfusion injury is a good tool for researchers. Here, a customizable method to create a myocardial infarction (MI) model and MIRI by precision ligation of the left anterior descending coronary artery (LAD) through micromanipulation is described. Accurate and reproducible ligature positioning of the LAD helps obtain consistent results for heart injury. ST-segment changes can help to identify model accuracy. The serum level of cardiac troponin T (cTnT) is used to assess the myocardial injury, cardiac ultrasound is employed to evaluate the myocardial systolic function, and Evans-Blue/triphenyl tetrazolium chloride staining is used to measure infarct size. In general, this protocol reduces procedure duration, ensures controllable infarct size, and improves mouse survival.

Here we describe a simple and reproducible method that can induce myocardial infarction or myocardial ischemia-reperfusion injury in mice by precision ligation of the left anterior descending coronary artery through micromanipulation.

Acute myocardial infarction is a common cardiovascular disease with high mortality. Myocardial reperfusion injury can counteract the beneficial effects of ...

Model of Ischemia and Reperfusion Injury in Rabbits

The protocol here provides a simple, highly replicable methodology to induce in situ acute regional myocardial ischemia in the rabbit for non-survival and survival experiments. New Zealand White adult rabbit is sedated with atropine, acepromazine, butorphanol, and isoflurane. The animal is intubated and placed on mechanical ventilation. An intravenous catheter is inserted into the marginal ear vein for the infusion of medications. The animal is pre-medicated with heparin, lidocaine, and lactated Ringer&#39;s solution. A carotid cut-down is performed to obtain arterial line access for blood pressure monitoring. Select physiologic and mechanical parameters are monitored and recorded by continuous real-time analysis.
With the animal sedated and fully anesthetized, either a fourth intercostal space small left thoracotomy (survival) or midline sternotomy (non-survival) is performed. The pericardium is opened, and the left anterior descending (LAD) artery is located.
A polypropylene suture is passed around the second or third diagonal branch of the LAD artery, and the polypropylene filament is threaded through a small vinyl tube, forming a snare. The animal is subjected to 30 min of regional ischemia, achieved by occluding the LAD by tightening the snare. Myocardial ischemia is confirmed visually by regional cyanosis of the epicardium. Following regional ischemia, the ligature is loosened, and the heart is allowed to re-perfuse.
For both survival and non-survival experiments, the myocardial function can be assessed via an echocardiography (ECHO) measurement of the fractional shortening. For non-survival studies, data from sonomicrometry collected using three digital piezoelectric ultrasonic probes implanted within the ischemic area and the left ventricle developed pressure (LVDP) using an apically inserted left ventricle (LV) catheter can be continuously acquired for evaluating the regional and global myocardial function, respectively.
For survival studies, the incision is closed, a left needle thoracentesis is performed for pleural air evacuation, and postoperative pain control is achieved.

The present study demonstrates a highly reproducible animal model of acute regional myocardial ischemia and reperfusion injury in rabbits using a left mini-thoracotomy for survival cases or a midline sternotomy for non-survival cases.

The protocol here provides a simple, highly replicable methodology to induce in situ acute regional myocardial ischemia in the rabbit for non-survival and ...

Closed-Chest Technique for Murine Myocardial Ischemia-Reperfusion Injury

Remotely Triggered LAD Occlusion Using a Balloon Catheter in Spontaneously Breathing Mice

Acute myocardial infarction (AMI) is a prevalent and high-mortality cardiovascular condition. Despite advancements in revascularization strategies for AMI, it frequently leads to myocardial ischemia-reperfusion injury (IRI), amplifying cardiac damage. Murine models serve as vital tools for investigating both acute injury and chronic myocardial remodeling in vivo. This study presents a unique closed-chest technique for remotely inducing myocardial IRI in mice, enabling the investigation of the very early phase of occlusion and reperfusion using in-vivo imaging such as MRI or PET. The protocol utilizes a remote occlusion method, allowing precise control over ischemia initiation after chest closure. It reduces surgical trauma, enables spontaneous breathing, and enhances experimental consistency. What sets this technique apart is its potential for simultaneous noninvasive imaging, including ultrasound and magnetic resonance imaging (MRI), during occlusion and reperfusion events. It offers a unique opportunity to analyze tissue responses in almost real-time, providing critical insights into processes during ischemia and reperfusion. Extensive systematic testing of this innovative approach was conducted, measuring cardiac necrosis markers for infarction, assessing the area at risk using contrast-enhanced MRI, and staining infarcts at the scar maturation stage. Through these investigations, emphasis was placed on the value of the proposed tool in advancing research approaches to myocardial ischemia-reperfusion injury and accelerating the development of targeted interventions. Preliminary findings demonstrating the feasibility of combining the proposed innovative experimental protocol with noninvasive imaging techniques are presented herein. These initial results highlight the benefit of utilizing the purpose-built animal cradle to remotely induce myocardial ischemia while simultaneously conducting MRI scans.

Author Spotlight: A Novel Standardized Technique for Real-Time Biomedical Imaging of Acute Myocardial Injury

This article presents a unique closed-chest technique for inducing myocardial ischemia-reperfusion injury (IRI) in mice. The presented method allows mice to breathe spontaneously while remotely inducing myocardial ischemia. This provides access to the animal for studying the dynamic processes of ischemia and reperfusion in situ and in real-time via noninvasive imaging.

Acute myocardial infarction (AMI) is a prevalent and high-mortality cardiovascular condition. Despite advancements in revascularization strategies for ...

Confirmation of Myocardial Ischemia and Reperfusion Injury in Mice Using Surface Pad Electrocardiography

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