NOTE: All Procedures were approved by the Lifespan Institutional Animal Care and Use Committee and all animals and procedures according to the National research Council Guide for Care and Use of Laboratory Animals10.
1. Balloon Fabrication and Left Ventricular Pressure Monitoring Circuit
<ol>
	<li>Construct LV balloons according to Miller et al.11 In a beaker with a stir bar, mix 9.5 ml of distilled water, 14.2 ml of light corn syrup and 33.8 g of sucrose, and heat on a hot plate, mix until the sugar is dissolved. Continue heating solution until it reaches approximately 150 &#186;C. 
	NOTE: Numerous methods exist to construct LV balloons including modification of condom tips and cling film construction9,12. We find the above method to be relatively easy to construct leak-free balloons, however an advantage of cling film balloons includes previous characterization for appropriate frequency-response relationships9.</li>
	<li>Break dry spaghetti strands into pieces about 5 cm in length and dip one side of each piece about 1 cm deep into the sugar solution and slowly remove.</li>
	<li>Place the dry end of the pasta strand into a polystyrene foam block and suspend so the sugar mix drips down and forms a teardrop shape mold. Leave O/N in a desiccator so the mold hardens.</li>
	<li>The following day, dip the molds into silicone dispersion gel. Place pasta strands back into polystyrene foam block and place in a a 37 &#186;C oven for 2 hr or until dry. Repeat this step once more so two layers of silicone are applied.</li>
	<li>Once dry, place in water for several hours to assist in removal of balloon from the mold. Store balloons in 0.02% sodium azide solution.</li>
	<li>Use a 23 G blood collection set to produce a customized balloon cannula by cutting a needle to create a blunt tip and placing notches in the needle.</li>
	<li>Connect the tubing to a pressure transducer and flush with water while keeping tubing and cannula completely submerged to prevent air from entering the system. Place a balloon that has been filled with water onto the cannula and use 2-0 silk sutures to tie onto cannula. Test balloon by ability to maintain pressure (~100 mmHg for at least 1 hr)</li>
</ol>
2. Preparation of the Isolated Heart Perfusion System
<ol>
	<li>Initially, wash and warm the system. Turn on a 37 &#186;C warm water bath circulator that has been connected to the Langendorff apparatus, and fill buffer reservoir with distilled water. Connect a new glass fiber filter to tubing flowing into apparatus and turn on pump to flush out the system. During prep time, monitor the temperature of water expelled from the Langendorff block using a temperature probe, to ensure that water bath is set accurately. Also, turn on a refrigerated circulator pump and set to 20 &#186;C that will be used during cardioplegic arrest of the heart.</li>
	<li>In the meantime prepare the following solutions. Prepare 1 L of cardioplegia solution (110 mM NaCl, 16 mM KCl, 16 mM MgCl2, 1.5 mM CaCl2, 10 mM NaHCO3) and filter. Also, prepare 2 L of KHB (118 mM NaCl, 4.8 mM KCl, 1.2 mM KH2PO4, 1.7 mM MgSO4, 2 mM Na-pyruvate, 6 mM dextrose, 24.9 mM NaHCO3 (pre-aerated with CO2),1.4 mM CaCl2 (added last). Once dissolved, remove precipitates by filtering into a flask using a 5 &#181;m glass fritted funnel. Place some into a small dish on ice to be used during surgery and isolation.</li>
	<li>Oxygenate the perfusate (KHB) with 95% O2/5% CO2 for at least 30 min prior to use. Perfuse Langendorff system with KHB to remove any remaining water from system and buffer reservoir. For mice, set the initial pump speed to ~2.0 ml/min. Place 2-0 and 4-0 silk sutures near perfusion cannula. Run the pump until the tubing and apparatus are filled with perfusate and ensure the bubble strup is filled with perfusate. 
	NOTE: For long periods of perfusion and stable function it is highly recommended to install an in-line glass fiber filter (~1 &#181;m) in the perfusion circuit to collect any precipitates that could form clots in the heart</li>
</ol>
3. Mouse Surgery
3.1) Mouse Anesthesia and Handling
<ol>
	<li>Prepare syringe with anesthetic dose of 80 mg/kg ketamine and 5 mg/ml xylazine mixture, and add sterile 0.9% saline to bring up volume to 0.2 ml. Inject Heparin I.P. (50 &#181;l of 1,000 U/ml solution).</li>
	<li>Place mouse back into transport container (~10-20 min) and wait for it to lose consciousness as heparin takes effect. Periodically do a toe pinch to monitor pain reflex.</li>
</ol>
3.2) Remove the Heart
<ol>
	<li>Once mouse is fully unconscious and does not respond to toe pinch, secure it to an appropriate operating table using pins or 25 G syringe tips through the extremities. Perform a thoracotomy to expose the heart. Cut a small hole just below the sternum and extend the cut to the sides of the mouse avoiding the diaphragm.</li>
	<li>Quickly cut the diaphragm and then rapidly cut up the sides of the rib cage. Flip back the rib cage like a clam shell to expose the thoracic cavity. Gently take hold of the heart, place scissors underneath and remove the heart. 
	NOTE: It is essential to quickly remove the heart once it is exposed in the thoracic cavity and the lungs are non-functional.</li>
</ol>
3.3) Clean the Heart
<ol>
	<li>Place the heart into the dish containing ice cold KHB and trim off any large pieces of lung tissue attached. Use forceps to pick up the heart and locate the aorta. Gently squeeze the heart and look for any blood that appears, as this should be the open end of the aorta. Use fine tip tweezers to hold the heart by the open end of the aorta.</li>
</ol>
4. Mounting the Heart, Starting Perfusion, and Placing the Balloon
4.1) Mounting and Cannulating the Heart
<ol>
	<li>Before transporting the heart to the Langendorff apparatus, turn on the pump ensuring that the perfusion pressure is low (~20 mmHg). Hold the heart by the aorta just under the cannula. Using another set of fine tip tweezers, carefully open the aorta and slide heart up onto the cannula. Hold it in place with one set of tweezers and then use a soft edged clip to temporarily secure it to cannula. 
	NOTE: Quickly mounting the heart to the perfusion cannula is critical for a good preparation. With experience, the procedures from opening the thoracic cavity until mounting the heart should take between 1 and 2 min.</li>
	<li>Tie and initiate perfusion as follows, using 4-0 silk suture secure aorta to the cannula directly below the clip. Ensure the tie is also around the metal aortic cannula, so the aorta does not get tied off when tightening the knot. Once one tie is firmly placed, remove the clip.</li>
	<li>Use additional silk sutures to fasten the aorta securely to the cannula making sure to tie off below any vessel branches that may be coming off the aorta. These can often be spotted by perfusate leaking or shooting out from the aorta. 
	NOTE: During the initial stages of perfusion the blood should be washing from the heart and the entire heart should appear a soft pink color. Dark discolorations that do not wash out indicate ischemic areas likely due to air emboli or clots and the heart should not be used.</li>
</ol>
4.2) Establishing Perfusion and LVP Measurement
<ol>
	<li>Slowly increase the perfusion pressure and increase pump speed until the perfusion pressure reaches 70 mmHg. 
	NOTE: Standard perfusion pressures for mouse perfusion can be 70-90 mmHg, but must be kept constant from animal to animal within an experiment.</li>
	<li>Clean the remaining tissue (lung, thyroid etc.) that may still be attached to the heart. Use scissors to cut off the left atria in order to create an opening into the left ventricle.</li>
	<li>Place the balloon onto the holder and deflate. Position the balloon cannula near the aortic cannula, directly over the opening into the left ventricle. Carefully insert balloon downward into the left ventricle while holding the heart in place so the aorta does not tear.</li>
	<li>Once it is in place slowly start to inflate the balloon until the LVEDP reaches ~8 mmHg. Place the temperature probe against the bottom of the heart so it measures the temperature of the effluent.</li>
</ol>
4.3) Basal Measurement
<ol>
	<li>Seal the heart in a water jacketed perfusion chamber. During this time continuously perfuse the heart with KHB, and ensure that the temperature continues to rise. Monitor the temperature and adjust the water bath accordingly until it reaches approximately ~37 &#186;C. 
	NOTE: Monitoring temperature in the initial phases of perfusion is critical as coronary flows and subsequently temperature can vary from heart to heart. Temperature of the effluent perfusate is monitored for desired myocardial temperature via a temperature probe placed at the apex of the heart. In addition, in comparison to other Langendorff perfusion protocols, the heart is not submerged in CP or KHB during perfusion or arrest, this is mainly done to quickly adjust temperature as well as provide efficient CP delivery without diffusion.</li>
	<li>Begin to continuously record functional measurement using a data acquisition system attached to the appropriate sensors, including perfusion pressure, left ventricular pressure (pressure transducer attached to LVP balloon), temperature, and electrophysiological parameters (EKG, MAP if so equipped) (attachment will be according to manufacturers and specific to the individual sensors and data acquisition system). Once the effluent KHB reaches ~37 &#186;C for at least 15 min and cardiac functional parameters are stable, note the time for baseline measurements. 
	NOTE: Inclusion/exclusion criteria are applied to each heart. A LVDP of &#60;60 mmHg at baseline indicates a heart that should be removed from analysis. In addition, coronary flow &#62;4.5 ml/min, or inability to maintain perfusion pressure at baseline likely indicates a leaking or torn aorta. Also any hearts that have clear ischemic areas which can be confirmed visually (i.e.,&#160;clot) or exhibit ischemia-associated reductions in cardiac function (i.e., increasing LVeDP, extremely arrhythmic) are removed from analysis.</li>
	<li>Measure coronary flow via collection of the coronary effluent in a graduated cylinder for 1 min. Close the chamber and allow temperature and function to return to baseline values. Alternatively measure the coronary flow continuously in the perfusion line via a flow probe placed after the pressure relief circuit.</li>
</ol>
5. Initiating Cardioplegic Arrest
<ol>
	<li>Place 100 ml of cold cardioplegia buffer into another reservoir and. transfer the KHB reservoir cannula to the cardioplegia reservoir.</li>
	<li>Detach the water jacketed housing and the perfusion warming circuit from the warm water bath and connect to the refrigerated circulator using tubing quick disconnects. Use separate heating and cooling circulators to allow rapid temperature changes of the heart and perfusate.</li>
	<li>Follow the air bubble that gets introduced into the tubing during the switch, and once it reaches near the isolated heart start the timing for cardioplegia. Deliver cardioplegia for 2 min. After the initial dose stop the pump. Ensure to arrest the heart in diastole and be close to the desired CP temperature. 
	NOTE: A sufficient bubble trap is necessary to avoid air emboli in the heart. Alternatively a delivery system for both reservoirs switched by a stopcock can be easily constructed, however we find following the small air bubble that is introduced when swapping the cannula between reservoirs, an easy method of timing CP delivery given potentially different pump speeds between experiments (i.e., due to mouse/rat, perfusion pressure, filter resistance, etc.)</li>
	<li>Keep the heart in cardioplegia for 2 hr at ~20 &#186;C. Every half hour during CP turn the pump back on for 1 min&#160;so another dose is administered. 
	NOTE: Intermittent doses of CP every 30 min results in functional impairment of the heart without necrosis of the tissue. Longer periods between doses of CP (i.e., &#62;45 min) can result in necrosis and ischemic contracture and would be more appropriate for a model of necrotic injury associated with unprotected ischemia.</li>
</ol>
6. Reperfusion
<ol>
	<li>At the end of CP, transfer the reservoir cannula back to the oxygenated KHB. Connect the temperature control circuits to the heating circulator and turn the pump on to start reperfusion. At this point the cardioplegia washes out, observe the temperature rise and the heart starts beating again after 2-4 min of reperfusion. When the heart starts the heart beat is slow and often arrhythmic. 
	NOTE: There usually is not a large rise in diastolic LVP (i.e., &#8804;10 mmHg rise) which is often characteristic in pure ischemia models. Arrhythmias can persist well into reperfusion and occasionally the heart will need to be defibrillated. This can be achieved with the use of a stimulator set to higher voltages (~10-50 V) and electrodes placed at the base and apex of the heart.</li>
	<li>Allow the heart to reperfuse for 30 min, and during this time period take coronary flow measurements as necessary. Measure coronary flow by collecting the effluent with a graduated cylinder for 30-60 sec.</li>
</ol>
7. Collecting the Tissue
<ol>
	<li>Deflate the balloon and remove it from the left ventricle. Take the heart off the cannula and weigh it. In initial experiments, at least one transverse slice of the heart should be taken and TTC stained to evaluate necrosis to ensure that the heart is free of any necrotic injury.</li>
	<li>Collect heart slices for microscopy studies, wet/dry weights, etc. The remainder or whole heart should be rapidly frozen by placing into liquid nitrogen. Store frozen hearts in a -80 &#186;C freezer. 
	NOTE: For measurement of high energy phosphate compounds (i.e.,&#160;ATP, creatine phosphate), the hearts should be frozen immediately.</li>
</ol>

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	<li>Kloner, R. a., Jennings, R. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Consequences+of+Brief+Ischemia:+Stunning,+Preconditioning,+and+Their+Clinical+Implications:+Part+1.">Consequences of Brief Ischemia: Stunning, Preconditioning, and Their Clinical Implications: Part 1.</a> Circulation. 104 (24), 2981-2989 (2001).</li><li>Mentzer, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Myocardial+protection+in+heart+surgery.">Myocardial protection in heart surgery.</a> J Cardiovasc Pharmacol Ther. 16 (3-4), 290-297 (2011).</li><li>Chambers, D. J., Fallouh, H. 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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterisation,+utilisation+and+clinical+relevance+of+isolated+perfused+heart+models+of+ischaemia-induced+ventricular+fibrillation.">Characterisation, utilisation and clinical relevance of isolated perfused heart models of ischaemia-induced ventricular fibrillation.</a> Cardiovasc Res. 39 (1), 194-215 (1998).</li><li>Clements, R. T., Feng, J., Cordeiro, B., Bianchi, C., Sellke, F. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=p38+MAPK-dependent+small+HSP27+and+&#945;B-crystallin+phosphorylation+in+regulation+of+myocardial+function+following+cardioplegic+arrest.">p38 MAPK-dependent small HSP27 and &#945;B-crystallin phosphorylation in regulation of myocardial function following cardioplegic arrest.</a> American journal of physiology. 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The authors have nothing to disclose.

The preceding protocol details methods to evaluate myocardial stunning secondary to global ischemia associated with cardioplegic arrest. In our hands this protocol produces an approximate ~40% reduction in cardiac function (LVDP, +/- dP/dt) with minimal changes in heart rate at the 30 min post-reperfusion time point. As the heart is reperfused and rewarmed all parameters of cardiac function are reduced at initial time points with a greatly reduced heart rate before stabilizing between 20 and 30 min. Coronary flow is generally greatly increased during early reperfusion due to hyperemia, and then falls to ~20-30% less than control levels following 30 min reperfusion.
It is important to remember that myocardial stunning by definition should leave the heart free of cell death and necrosis which is characteristic of pure ischemia/MI models (i.e., &#62;20 min no flow regional ischemia). Initial studies should evaluate tissue histology to ensure a lack of necrotic injury. In addition, although myocardial stunning by definition should result in normal function following prolonged reperfusion (hr to days), It is likely this protocol will not demonstrate fully recovered function due to the ex vivo Langendorff perfusion which is associated with reductons in cardiac function in control perfused hearts over time. Nevertheless changes in the acute recovery of cardiac function in the absence of cell death/necrosis can be used as indices of the severity of stunning The major difference in this protocol vs. classic no-flow ischemia protocols is the use of a cardioprotection strategy, in this case being hyperkalemic cardioplegia. Hyperkalemic cardioplegia solutions provide protection from necrotic injury and cell death by causing diastolic cardiac arrest. Diastolic arrest of the heart promotes preservation of energy reserves. In addition, most clinically used cardioprotection protocols use hypothermia to further limit myocardial injury by reducing metabolic demand. Other factors that can be modulated during the above protocol include different formulations of cardioplegia solutions (hyperpolarizing formulations, Mg++, K+ levels, etc.), strategies (warm vs. cold, &#39;hot shot&#39;, etc.), and various drugs (kinase inhibitors, ion channel modulators, cardioprotective agents, etc.).
Since the heart is in a relatively well protected state, to get reproducible functional impairments in this model requires necessarily longer ischemia times (i.e.,&#160;greater than 2 hr). We have found that rodent hearts are relatively resistant to injury in this model, especially when compared to larger animals (pigs, humans) which reliably display stunned myocardium during much shorter periods (i.e., 30 min). We have also found that intermittent delivery is necessary to protect the heart from severe ischemic damage as a 45-60 min interval following CP delivery can result in gross diastolic dysfunction, ischemic contracture, and tissue injury upon reperfusion. Other readily adaptable components of the protocol can include investigations related to CP constituents and the role of hypo/normothermic arrest as well as different arrest strategies not reliant on K+ (Na channel blockers, hyperpolarizing agents)3.
There are also a number of important limitations to this technique when being used to model stunning associated with clinical cardiac surgery. First, the gross majority of clinical CP solutions are mixed with blood (~4 blood:1 CP ratio). This is generally not feasible in mice due to the perfusion circuitry volume as well as the need for treated tubing and fiber oxygenators. Often for larger animals (guinea pigs / rabbits) donor animals would also be required. In addition, as in all isolated organ models, the influence of peripheral factors (i.e., inflammatory signals, blood reperfusion, etc.) is completely ignored. Nevertheless it is a practical, efficient, and economical model for preliminary studies to test pharmacological additives and different cardioprotection strategies.
The protocol is essentially the same in rat hearts, with the exception of a larger balloon and aortic cannula13,14. In addition, the rat heart preparation requires significantly greater flow rates (12-20 ml/min). Due to its size, the rat heart preparation is considerably easier to learn and reproducibly perform. To assess other types of injury resulting in myocardial stunning the protocol can be easily modified. To emulate reversible ischemic injury, simply stop the perfusion pump for brief periods. In rats ~20 min global ischemia will result in reduced contractile function without gross effects on cell death and infarction.

Myocardial stunning is defined as reversible reduced contractile activity despite restoration of adequate blood flow following a brief period of ischemia or prolonged periods of ischemic insults with cardioprotection1,2,3,4,5. The method presented is specifically used to model clinically relevant ischemic insults which can result in reversible impairments in contractile function (i.e., ischemic insults associated with cardiac surgery utilizing cardioplegic arrest, brief periods of ischemia, angina, etc.). In contrast to severe ischemia studies (myocardial infarction, necrosis) this protocol was developed to evaluate myocardial functional recovery and cardioprotection without tissue injury, remodeling, and cell death. The majority of the paper discusses a standard cardioplegic arrest protocol with elements similar to a cardiac surgery using hypothermia and intermittent cardioplegia delivery.
Myocardial protection during the majority of cardiac surgeries relies on cardioplegia and cardiopulmonary bypass. Although cardioplegia (CP) solutions and strategies vary widely (blood, crystalloid, cold, warm etc.) the most common elements are 1) hyperkalemia and/or other strategies to arrest the heart in diastole, thereby limiting energy utilization resulting from myocardial contraction, and 2) hypothermia to slow metabolism and help maintain ATP and other energy reserves while arrested. Current cardioplegia solutions provide protection to the heart against ischemic insults that would otherwise prove lethal. However, cardioprotective strategies during surgical ischemic insults are not perfect, and the resultant mild ischemic injury can result in reversible cardiac contractile dysfunction despite adequate blood flow (myocardial stunning), acidosis, cardiomyocyte damage, and vascular effects including reduced coronary perfusion and vasospasm.
This protocol differs from standard isolated heart ischemia models evaluating myocardial infarction and severe ischemia in that it evaluates milder ischemic insults which can result in impaired cardiac function following brief ischemia or ischemic insults associated with cardioplegic arrest. (for review on Langendorff perfusion techniques and I/R studies see6&#8211;8). For general guidelines and a thorough analysis of experimental parameters associated with mouse isolated perfused hearts see Sutherland et al. 20039 The technique presented here details the necessary equipment, reagents, steps, strategies and tips to reliably induce stunning in mouse hearts. Minor modifications are necessary to apply the technique to rats.
Briefly isolated mouse hearts are Langendorff perfused for approximately 30 min with physiologic Krebs-Henseleit buffer (KHB), followed by cold protected cardiac arrest via delivery of a hyperkalemic hypothermic cardioplegia solution. Following arrest, cardiac functional recovery is monitored during rewarming and reperfusion of the heart with KHB. Changes in the degree of recovery of cardiac contractile function can be evaluated to assess cardioprotective agents and different cardioprotection strategies.

<table><tbody><tr><td>Cardioplegia Solution (St Thomas II)</td><td></td><td></td><td>Symbol / Concentrations (mM)</td></tr><tr><td>Sodium Chloride</td><td></td><td></td><td>NaCl; 110</td></tr><tr><td>Potassium chloride</td><td></td><td></td><td>KCl; 16</td></tr><tr><td>Calcium Chloride</td><td></td><td></td><td>CaCL2; 1.5</td></tr><tr><td>Magnesium Chloride</td><td></td><td></td><td>MgCL2; 16</td></tr><tr><td>Sodium Bicarbonate</td><td></td><td></td><td>NaHCO3; 10</td></tr><tr><td>Krebs-Heinslet Buffer</td><td></td><td></td><td></td></tr><tr><td>Sodium Chloride</td><td></td><td></td><td>NaCl; 118</td></tr><tr><td>Potassium Chloride</td><td></td><td></td><td>KCl; 4.8</td></tr><tr><td>Magnesium Sulfate</td><td></td><td></td><td>MgSO4; 1.7</td></tr><tr><td>Sodium Bicarbonate</td><td></td><td></td><td>NaHCO3; 24.9</td></tr><tr><td>Potassium Phosphate (monobasic)</td><td></td><td></td><td>KH2PO4; 1.2</td></tr><tr><td>Calcium Chloride</td><td></td><td></td><td>CaCL2; 1.4</td></tr><tr><td>Sodium Pyruvate</td><td></td><td></td><td>Na pyruvate; 2</td></tr><tr><td>Glucose</td><td></td><td></td><td>C6H12O6; 6</td></tr><tr><td>Balloon reagents</td><td></td><td></td><td></td></tr><tr><td>Corn Syrup</td><td></td><td></td><td></td></tr><tr><td>Spaghetti</td><td></td><td></td><td></td></tr><tr><td>Silicon Dispersion Gel</td><td></td><td></td><td></td></tr><tr><td>styrofoam block</td><td></td><td></td><td></td></tr><tr><td>lab oven/incubator ( 50C)</td><td></td><td></td><td></td></tr><tr><td>Langendorff Perfusion equipment</td><td></td><td></td><td></td></tr><tr><td>Isolated perfused heart sytem (IH-SR (Hugo-Sachs) or equivalent)</td><td></td><td></td><td></td></tr><tr><td>Data acquisition system (DSI, ADinstruments or equivalent)</td><td></td><td></td><td></td></tr><tr><td>Heated water circulator</td><td></td><td></td><td></td></tr><tr><td>Cooling water circulator</td><td></td><td></td><td></td></tr><tr><td>Perfusion pump capable of 2-30 ml/min</td><td></td><td></td><td></td></tr><tr><td>Inline perfusion filters - 1 um glass fiber</td><td></td><td></td><td></td></tr><tr><td>Pressure sensors and amplifiers for LVP and perfusion pressure</td><td></td><td></td><td></td></tr><tr><td>Small graduated cylinder (~10 mL)</td><td></td><td></td><td></td></tr><tr><td>Small temperature probe and thermometer (Werner or equivalent)</td><td></td><td></td><td></td></tr><tr><td>perfusion resevoir (1L)</td><td></td><td></td><td></td></tr><tr><td>cardioplegia resevoir (~200 mL)</td><td></td><td></td><td></td></tr><tr><td>gas bubbler</td><td></td><td></td><td></td></tr><tr><td>95/5 O2/CO2 mix</td><td></td><td></td><td></td></tr><tr><td>Surgical tools and reagents</td><td></td><td></td><td></td></tr><tr><td>Metzenbaum and Potz surgical scissors</td><td></td><td></td><td></td></tr><tr><td>two Dumont size 5 forceps</td><td></td><td></td><td></td></tr><tr><td>ketamine</td><td></td><td></td><td></td></tr><tr><td>xylazine</td><td></td><td></td><td></td></tr><tr><td>heparin</td><td></td><td></td><td></td></tr><tr><td>small clamp with soft sides to hold aorta (i.e. terminal clamp with taped ends)</td><td></td><td></td><td></td></tr><tr><td>Silk 2-0 and 4-0 sutures</td><td></td><td></td><td></td></tr></tbody></table>

murine isolated heart model of myocardial stunning associated with cardioplegic arrest

The following protocol is of use to evaluate impaired cardiac function or myocardial stunning following moderate ischemic insults. The technique is useful for modeling ischemic injury associated with numerous clinically relevant phenomenon including cardiac surgery with cardioplegic arrest and cardiopulmonary bypass, off-pump CABG, transplant, angina, brief ischemia, etc. The protocol presents a general method to model hypothermic hyperkalemic cardioplegic arrest and reperfusion in rodent hearts focusing on measurement of myocardial contractile function. In brief, a mouse heart is perfused in langendorff mode, instrumented with an intraventricular balloon, and baseline cardiac functional parameters are recorded. Following stabilization, the heart is then subject to brief infusion of a cardioprotective hypothermic cardioplegia solution to initiate diastolic arrest. Cardioplegia is delivered intermittently over 2 hr. The heart is then reperfused and warmed to normothermic temperatures and recovery of myocardial function is monitored. Use of this protocol results in reliable depressed cardiac contractile function free from gross myocardial tissue damage in rodents.

Figure 1 presents typical results from one mouse experiment. LVP (red line), dP/dt (green line), and temperature (purple line) were continusouly recorded over ~3 hr. Letters indicate a - baseline measurement, b, c, d, e - delivery of cardioplegia solution, f - start of reperfusion, g, h, i, j - measurement of cornary flow during reperfusion. Note, depressed LVDP and dP/dt upon reperfusion compared to baseline. Figure 1B includes the data from A recorded over ~2 sec. Note, the decrease in LVP and dP/dt similar to A, slight reductions in HR and slight increases in LVEDP. Usually after 30 min of reperfusion there is an ~40% decrease in cardiac function as evidenced by the left ventricular developed pressure LVDP depicted in Figure 1C.
<img alt="Figure 1" src="/files/ftp_upload/52433/52433fig1.jpg" /> 
Figure 1.&#160;Representative results of a whole experiment. (A) recording left ventricular pressure (top), first derivative of LV pressure, and temperature. (B) Short interval recordings to see trace detail and (C) Quantitation of left ventricular developed pressure (LVDP) for multiple experiments (n = 6). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52433/52433fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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Murine Isolated Heart Model of Myocardial Stunning Associated with Cardioplegic Arrest

The goal of this protocol to assess myocardial stunning following ischemic cardioplegic arrest in rodents.

The following protocol is of use to evaluate impaired cardiac function or myocardial stunning following moderate ischemic insults. The technique is useful ...

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Nanyang Technological University

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Medicine

11.6K Views.  Brown University. The goal of this protocol to assess myocardial stunning following ischemic cardioplegic arrest in rodents.

Video: Murine Isolated Heart Model of Myocardial Stunning Associated with Cardioplegic Arrest

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 ...

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 ...

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.

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 ...

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 ...

An Intact Pericardium Ischemic Rodent Model

This protocol has shown that the pericardium and its contents play an essential anti-fibrotic role in the ischemic rodent model (coronary ligation to induce myocardial injury). The majority of pre-clinical myocardial infarction models require the disruption of pericardial integrity with loss of the homeostatic cellular milieu. However, recently a methodology has been developed by us to induce myocardial infarction, which minimizes pericardial damage and retains the heart&#39;s resident immune cell population. An improved cardiac functional recovery in mice with an intact pericardial space following coronary ligation has been observed. This method provides an opportunity to study inflammatory responses in the pericardial space following myocardial infarction. Further development of the labeling techniques can be combined with this model to understand the fate and function of pericardial immune cells in regulating the inflammatory mechanisms that drive remodeling in the heart, including fibrosis.

This protocol outlines the steps for inducing myocardial infarction in mice while preserving the pericardium and its contents.

This protocol has shown that the pericardium and its contents play an essential anti-fibrotic role in the ischemic rodent model (coronary ligation to ...

A Mouse Model to Study Brain Physiology During Cardiac Arrest

Mouse Cardiac Arrest Model for Brain Imaging and Brain Physiology Monitoring During Ischemia and Resuscitation

Most cardiac arrest (CA) survivors experience varying degrees of neurologic deficits. To understand the mechanisms that underpin CA-induced brain injury and, subsequently, develop effective treatments, experimental CA research is essential. To this end, a few mouse CA models have been established. In most of these models, the mice are placed in the supine position in order to perform chest compression for cardiopulmonary resuscitation (CPR). However, this resuscitation procedure makes the real-time imaging/monitoring of brain physiology during CA and resuscitation challenging. To obtain such critical knowledge, the present protocol presents a mouse asphyxia CA model that does not require the chest compression CPR step. This model allows for the study of dynamic changes in blood flow, vascular structure, electrical potentials, and brain tissue oxygen from the pre-CA baseline to early post-CA reperfusion. Importantly, this model applies to aged mice. Thus, this mouse CA model is expected to be a critical tool for deciphering the impact of CA on brain physiology.

Author Spotlight: A Unique Mouse Model of Asphyxia-Induced Cardiac Arrest 

This protocol demonstrates a unique mouse model of asphyxia cardiac arrest that does not require chest compression for resuscitation. This model is useful for monitoring and imaging the dynamics of brain physiology during cardiac arrest and resuscitation.

Most cardiac arrest (CA) survivors experience varying degrees of neurologic deficits. To understand the mechanisms that underpin CA-induced brain injury ...

Neuroscience

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 ...

Rodent Working Heart Model for the Study of Myocardial Performance and Oxygen Consumption

Isolated working heart models have been used to understand the effects of loading conditions, heart rate and medications on myocardial performance in ways that cannot be accomplished in vivo. For example, inotropic medications commonly also affect preload and afterload, precluding load-independent assessments of their myocardial effects in vivo. Additionally, this model allows for sampling of coronary sinus effluent without contamination from systemic venous return, permitting assessment of myocardial oxygen consumption. Further, the advent of miniaturized pressure-volume catheters has allowed for the precise quantification of markers of both systolic and diastolic performance. We describe a model in which the left ventricle can be studied while performing both volume and pressure work under controlled conditions. 
In this technique, the heart and lungs of a Sprague-Dawley rat (weight 300-500 g) are removed en bloc under general anesthesia. The aorta is dissected free and cannulated for retrograde perfusion with oxygenated Krebs buffer. The pulmonary arteries and veins are ligated and the lungs removed from the preparation. The left atrium is then incised and cannulated using a separate venous cannula, attached to a preload block. Once this is determined to be leak-free, the left heart is loaded and retrograde perfusion stopped, creating the working heart model. The pulmonary artery is incised and cannulated for collection of coronary effluent and determination of myocardial oxygen consumption. A pressure-volume catheter is placed into the left ventricle either retrograde or through apical puncture. If desired, atrial pacing wires can be placed for more precise control of heart rate. This model allows for precise control of preload (using a left atrial pressure block), afterload (using an afterload block), heart rate (using pacing wires) and oxygen tension (using oxygen mixtures within the perfusate).

Isolated working heart models can be used to measure the effect of loading conditions, heart rate, and medications on myocardial performance and oxygen consumption. We describe methods for preparation of a rodent left heart working model that permits study of systolic and diastolic performance and oxygen consumption under various conditions.

Isolated working heart models have been used to understand the effects of loading conditions, heart rate and medications on myocardial performance in ways ...

Biology

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 ...

Murine Isolated Heart Model of Myocardial Stunning Associated with Cardioplegic Arrest

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