NIH/NINDS R01NS112706 (R.L.)

Institutional Animal Care and Use Committee of Columbia University Medical Center&#39;s approvals were obtained for all methods described. Wild-type C57Bl/6 male postnatal day 10 mice were used for the study.
1. Preparation of Langendorff apparatus
<ol>
	<li>To minimize complexity, use non-recirculating oxygenated perfusate within the Langendorff apparatus (see Table of Materials) via constant flow or constant pressure.
	<ol>
		<li>Use Krebs-Henseleit buffer (KHB), containing 120 mmol/L of NaCl, 4.7 mmol/L of KCl, 1.2 mmol/L of MgSO4, 1.2 mmol/L of KH2PO4, 1.25 mmol/L of CaCl2, 25 mmol/L of NaHCO3,&#160;and 11 mmol/L of glucose at pH 7.4 (see Table of Materials), equilibrate with 95% of O2 and 5% of CO2 within the Langendorff apparatus and maintain at 37 &#176;C.</li>
	</ol>
	</li>
	<li>For the constant flow approach, maintain a continuous flow rate at ~2.5 mL&#8729;min-1. 
	NOTE: This flow rate will approximate coronary flow of ~75-80 mL/g&#8729;min, given that the average weight of a 10 day old (P10) mouse heart is ~30 mg17,18.</li>
</ol>
2. Fabrication of aortic cannula
<ol>
	<li>Fabricate the newborn mouse aortic cannula from a 26 G stainless steel needle (see Table of Materials). Using sharp scissors, cut off the tip of the needle to blunt the end. Take care not to crimp or restrict the diameter of the needle lumen. Smooth the cut edge and remove any burs by gently scraping the blunted end on the laboratory benchtop using a to-and-fro motion. 
	NOTE: Microscopic burs and sharp edges must be removed because they can tear the newborn mouse aorta and damage the aortic valve. Alternatively, use fine-grit sandpaper.</li>
	<li>Attach the fabricated cannula to the Langendorff apparatus and assess flow and resistance. Measure flow rates through the cannula by collecting and measuring buffer quantity over a known time period. Ensure actual flow is equal to the set flow rate of 2.5 mL min-1.</li>
	<li>Quantify the pressure differential across the cannula with KHB flowing by following the steps below.
	<ol>
		<li>Measure pressure in the system with and without the fabricated cannula attached.</li>
		<li>Divide pressure differential across cannula by the flow rate to obtain cannula resistance as per Ohm&#39;s law15.</li>
		<li>Ensure that the fabricated cannula resistance comprises ~16.0 &#177; 1.9 mmHg&#8729;min&#8729;mL-1 of the total resistance6. Excessive resistance suggests a potentially compromised cannula lumen. 
		NOTE: Sample calculation: Pwith cannula - P without cannula = &#916;P. If Pwith = 48 and Pwithout = 8 then &#916;P = 40. At a flow rate (Q) of 2.5 mL min-1 and &#916;P of 40 cannula resistance equals 16 mmHg&#8729;min&#8729;mL-1 using R =&#160;&#916;P/Q&#160;= 40 / 2.5 = 16.</li>
	</ol>
	</li>
	<li>Remove the 26 G cannula and attach the high-pressure tubing (see Table of Materials) to the cannulation site on the Langendorff apparatus. Attach the aortic cannula to the distal end of the tubing. De-air the tubing and the cannula with oxygenated buffer, ensuring that all bubbles are removed. 
	NOTE: The use of high-pressure tubing in this manner permits the cannula to be extended to a more remote position. This is necessary to allow aortic cannulation with a dissecting microscope adjacent to the setup (Figure 1).</li>
</ol>
3. Organ harvesting
<ol>
	<li>Anticoagulate mice via intraperitoneal (IP) injection of heparin (10 kU/kg) (see Table of Materials) to prevent the formation of coronary microthrombi using a 26 G needle on 1 mL syringe. Allow several minutes for heparin to circulate before proceeding with the injection of any anesthetic.</li>
	<li>Anesthetize the animal with an IP injection using a 26 G needle on 1 mL syringe. 
	NOTE: It is essential to carefully monitor the animal after anesthetic injection to avoid apnea and subsequent hypoxia. Pentobarbital (70 mg/kg) is a reliable choice of anesthetic, as it allows for rapid onset of sedation without inducing apnea19,20. Other anesthetic agents can be utilized, provided that the doses used do not cause apnea21. Investigators should consider the effects of alternative sedative-hypnotics on cardiac function22,23. Cervical dislocation as a primary mode of euthanasia may prolong pre-cannulation hypoxia and ischemia.</li>
	<li>Place the mouse in the supine position and secure limbs immediately upon loss of consciousness. Use small gauge hypodermic needles to secure each limb.&#160;Begin harvesting as soon as the animal is unresponsive to toe pinch; the animal should breathe spontaneously during the initial dissection.</li>
	<li>Make a transverse subxiphoid incision across the animal&#39;s width to expose the abdominal cavity using straight dissecting scissors (see Table of Materials). 
	NOTE: Sterile technique is not necessary given that the procedure represents nonsurvival surgery.
	<ol>
		<li>Identify the diaphragm superiorly and incise the anterior portion completely. Cut the ribcage bilaterally along the mid-axillary line in a cephalad direction. Ask an assistant to grasp the xiphoid process with forceps and reflect the sternum and ribs cranially to expose the thoracic organs.</li>
	</ol>
	</li>
	<li>Identify the infra-diaphragmatic inferior vena cava (IVC) above the liver. Transect the IVC with a curved iris scissor while maintaining slight anterior and cephalad tension on the proximal segment with iris forceps (see Table of Materials).
	<ol>
		<li>Cut posteriorly along the anterior surface of the spine using curved iris scissors while pulling the IVC up and out of the thoracic cavity. As the heart is mobilized, angle the scissors anteriorly and sever&#160;the great vessels superiorly to completely remove the heart and lungs. 
		&#8203;NOTE: This method permits rapid explantation of the heart and lungs en bloc.</li>
	</ol>
	</li>
	<li>Immediately submerge the specimen in ice-cold KHB or saline. The heart should stop beating within seconds.</li>
</ol>
4. Cannulation
<ol>
	<li>Cut a piece of paper towel and place it at the bottom of a shallow Petri dish to provide friction to stabilize the heart during cannulation. Moisten with ice-cold KHB to prevent the heart from adhering to it.
	<ol>
		<li>Place the prepared Petri dish under the dissecting microscope and adjust the focus. Place the aortic cannula attached to the high-pressure extension tubing under the dissecting microscope along with a 5-0 silk suture loosely tied around its hub (see Table of Materials). 
		NOTE: Care must be taken to limit the amount of fluid in the Petri dish because the air-filled lungs can float and cause the excised organs to move.</li>
	</ol>
	</li>
	<li>Place the excised thoracic organs in the Petri dish. Under the microscope, identify the thymus by its white sheen and two lobes and orient the specimen such that the thymus is anterior and superior24. This will ensure proper orientation of the heart.</li>
	<li>Using forceps, bluntly separate the lobes of the thymus to expose the great vessels. Identify the aorta by locating distinguishing branching features of the aortic arch. 
	NOTE: A dark purple hue often demarcates the right ventricle and the pulmonary artery. The ascending aorta is located between the main pulmonary artery and the right atrium.</li>
	<li>Transect the aorta with fine sharp scissors (see Table of Materials) just proximal to the subclavian artery takeoff. 
	NOTE: If the aorta is transected too close to the aortic valve, there will not be enough aortic tissue to enable the cannula to be secured. Alternatively, if the aorta is transected too high, perfusate can leak out of one or more aortic branches (such as the subclavian artery).</li>
	<li>Gently grasp the transected aorta using jeweler-style fine curved forceps (see Table of Materials). Carefully cannulate the aorta with a 26 G blunt needle, taking care not to damage the aortic valve. Hold in place by grasping the aorta with the fine curved forceps around the cannula. Once control of the aorta is established, initiate retrograde perfusion to limit the ischemic time. 
	NOTE: The heart should begin to beat and will become pale as blood is drained from the myocardium and KHB perfuses the coronary arteries. Failure to spontaneously beat, presence of ventricular engorgement, or lack of color change of the heart indicates a malpositioned cannula.</li>
	<li>Ask the assistant to grasp the ends of the loosely tied suture and carefully ensnare the aorta around the cannula. Cinch the suture above or below the curved fine forceps (holding the cannula in place), depending on the amount of aortic tissue and anatomical considerations. Tighten the suture and confirm the adequacy of coronary flow.</li>
	<li>Disconnect the high-pressure tubing from the Langendorff apparatus. Grasp the hub of the cannula and disconnect the blunt needle from the high-pressure extension tubing. Rapidly attach the hub of the cannula to the apparatus. 
	NOTE: Care must be taken not to dislodge the heart or entrain air into the cannula.</li>
	<li>Once the heart is hung on the Langendorff apparatus in the usual position, and adequate perfusion is confirmed, carefully trim off lung, thymus, and excess tissue. Incise the right atrium to permit coronary sinus effluent to drip freely.</li>
</ol>
5. Functional measurement
<ol>
	<li>Make a small knot at the end of a 5-0 silk suture (attached to a curved needle). Pierce a small piece of paraffin film (2-3 mm x 2-3 mm) with the needle and slide the paraffin to the knotted end. Carefully pass the needle through the apex of the ventricle and pull the suture through the heart until the paraffin film is snug against the lateral wall of the ventricle. 
	NOTE: The paraffin film helps to prevent the knot from tearing the heart and pulling through the ventricle.</li>
	<li>Pass the needle through the opening of the water-filled warming jacket of the Langendorff apparatus. The heart can now be encased and warmed.</li>
	<li>Attach the needle to the force transducer (see Table of Materials) in such a manner that avoids the coronary sinus drip. Adjust the suture to apply 1-2 g of basal tension, as indicated by the diastolic tension or nadir in tension tracing. 
	NOTE: Avoid pulling the heart off the cannula or twisting the aorta, thereby compromising coronary perfusion.</li>
	<li>Place surface electrodes on the superior and inferior poles of the heart to record the electrocardiogram. 
	NOTE: Use pediatric temporary epicardial pacing wire with the needle removed for flexible surface electrode connected to Bio Amp (see Table of Materials).</li>
	<li>Sample the coronary sinus effluent for analysis using a 24 G IV catheter (see Table of Materials).</li>
	<li>Subtract the cannula resistance from the total system resistance to obtain coronary resistance per Kirchhoff&#39;s law25.</li>
</ol>

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(60), e3756 (2012).</li><li>Bednarczyk, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Incorporating+dynamic+assessment+of+fluid+responsiveness+into+goal-directed+therapy:+A+systematic+review+and+meta-analysis.">Incorporating dynamic assessment of fluid responsiveness into goal-directed therapy: A systematic review and meta-analysis.</a> Critical Care Medicine. 45 (9), 1538 (2017).</li><li>Louch, W., Sheehan, K., Wolska, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+in+cardiomyocyte+isolation,+culture,+and+gene+transfer.">Methods in cardiomyocyte isolation, culture, and gene transfer.</a> Journal of Molecular and Cellular Cardiology. 51 (3), 288-298 (2011).</li><li>Ackers-Johnson, M., Foo, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Langendorff-free+isolation+and+propagation+of+adult+mouse+cardiomyocytes.">Langendorff-free isolation and propagation of adult mouse cardiomyocytes.</a> Methods in Molecular Biology. 1940, 193-204 (2019).</li><li>Peng, Y., Buller, C., Charpie, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Impact+of+N-acetylcysteine+on+neonatal+cardiomyocyte+ischemia-reperfusion+injury.">Impact of N-acetylcysteine on neonatal cardiomyocyte ischemia-reperfusion injury.</a> Pediatric Research. 70 (1), 61-66 (2011).</li><li>Jarmakani, J., Nakazawa, M., Nagatomo, T., Langer, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effect+of+hypoxia+on+mechanical+function+in+the+neonatal+mammalian+heart.">Effect of hypoxia on mechanical function in the neonatal mammalian heart.</a> American Journal of Physiology-Heart and Circulatory Physiology. 235 (5), 469-474 (1978).</li><li>Podesser, B., Hausleithner, V., Wollenek, G., Seitelberger, R., Wolner, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Langendorff+and+ischemia+in+immature+and+neonatal+myocardia:+Two+essential+key-words+in+Today's+cardiothoracic+research.">Langendorff and ischemia in immature and neonatal myocardia: Two essential key-words in Today's cardiothoracic research.</a> Acta Chirurgica Austriaca. 25 (6), 434-437 (1993).</li><li>Popescu, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Getting+an+early+start+in+understanding+perinatal+asphyxia+impact+on+the+cardiovascular+system.">Getting an early start in understanding perinatal asphyxia impact on the cardiovascular system.</a> Frontiers in Pediatrics. 8, 68 (2020).</li></ol>

The authors have nothing to disclose.

The present work describes successful aortic cannulation and retrograde perfusion in the isolated newborn mouse heart. Importantly, it allows researchers to overcome the barriers that young murine age and small heart size previously presented8. While not complex in design, the approach does require a significant degree of technical skill. Key steps that will inevitably challenge even the most technically proficient investigators will be cannulation of the aorta and securing the cannula in place. D...

Ex-vivo heart preparations have been a staple of physiologic, pathophysiologic, and pharmacologic studies for over a century. Stemming from the work of Elias Cyon in the 1860s, Oskar Langendorff adapted the isolated frog model for retrograde perfusion, pressurizing the aortic root to provide coronary flow with an oxygenated perfusate1. Using his adaptation, Langendorff was able to demonstrate a correlation between coronary circulation and mechanical function2. The ex-vivo retrograde perfused heart, later eponymously dubbed the Langendorff technique, has remained a cornerstone of physiologic investigation, leveraging its simplicity to powerfully study the isolated heart in the absence of potential confounders. The Langendorff preparation has been modified further to permit the heart to eject (the so-called &#34;working heart&#34;) and allow the perfusate to recirculate3. However, the primary physiologic endpoints of interest have remained unchanged. Such endpoints include measures of contractile function, electrical conduction, cardiac metabolism, and coronary resistance4.
To evaluate cardiac function in his original frog heart preparation, Langendorff measured the tension generated by ventricular contraction in the longitudinal axis using a suture connected between the heart&#39;s apex and a force transducer.5 Isometric contraction was quantified in this manner with basal tension applied to the heart in the absence of ventricular filling. Refinement of the approach has led to fluid-filled balloons placed into the left ventricle via the left atrium to evaluate myocardial performance during isovolumic contraction6. To assess cardiac rhythm and the heart rate, surface leads can be placed on the poles of the heart to enable investigators to record the electrocardiogram. However, relative bradycardia can be expected, given the obligatory denervation. Extrinsic pacing may serve to overcome this and eliminate heart rate variability between experiments1. Another outcome measure, myocardial metabolism, can be assessed by measuring the oxygen and metabolic substrate content in the coronary perfusate and effluent and calculating the difference between them7. Lactate quantification in the coronary effluent can aid in characterizing periods of anaerobic metabolism as is seen with hypoxia, hypoperfusion, ischemia-reperfusion, or metabolic perturbations7.
Langendorff&#39;s original work enabled the study of the ex-vivo mammalian heart, using cats as the primary subject5. Evaluation of the isolated rat heart gained popularity in the mid-1900s with Howard Morgan, who detailed the &#39;working heart&#39; rat model in 19675. The use of mice began only 25 years ago due to the technical complexity, tissue fragility, and relatively small murine heart size. Despite the challenges associated with mice study, the lower costs and ease of genetic manipulation have increased the appeal and demand of such murine ex-vivo preparations. Unfortunately, the application of the technique has been limited to adult animals, with juvenile 4-week-old mice being the youngest subjects utilized for ex-vivo study until quite recently8,9. While juvenile mice are &#34;relatively immature&#34; compared with adults, their utility as subjects for developmental biology studies is limited because they have, by and large, weaned from their birth dam and will soon begin puberty10. Adolescence occurs well beyond the postnatal transition in myocardial substrate utilization from glucose and lactate to fatty acids11. Thus, most information about the metabolic changes in the neonatal heart has historically resulted from ex-vivo work in larger species such as rabbits and guinea pig11.
Indeed, alternative approaches to the Langendorff preparation exist. These include in vitro experimentation, which lacks the whole organ functional data and context, or in vivo studies. This can be technically challenging and complicated by confounding variables such as the cardiovascular and respiratory effects of a requisite anesthetic agent, the influence of neurohumoral input, the consequences of core temperature, the nutritional status of the animal, and substrate availability12,13. Because the Langendorff approach permits the study of the isolated-perfused heart in an ex-vivo manner in a more controlled manner in the absence of such confounders, it has been and continues to be considered a powerful investigational tool. Therefore, the technique presented here gives researchers an experimental approach for the ex-vivo study of the newborn murine heart and limits time to reperfusion.
Investigating the heart during periods of development is an important consideration given the wide-ranging biochemical, physiologic, and anatomical transitions that occur during myocardial maturation. Shifts from anaerobic metabolism to oxidative phosphorylation, changes in substrate utilization, and progression from cell proliferation to hypertrophy are dynamic processes that uniquely occur in the immature heart11,14. Another critical aspect of the developing heart is that stressors encountered during necessary periods may produce heightened responses in the newborn heart and alter future susceptibility to insults in adulthood15. Although prior work has utilized newborn rats, lambs, and rabbits to study the Langendorff-perfused neonatal heart, advances permitting mice use are necessary given the importance of this species to developmental biology research16. To address this need, the first murine Langendorff-perfused newborn heart model using 10-day old animals was recently established6. Presented here is a method to enable successful aortic cannulation and establish retrograde perfusion of the isolated newborn murine heart. This approach may be utilized for pharmacology, ischemia-reperfusion, or metabolism studies focusing on whole organ function or can be adapted for the isolation of cardiomyocytes.

<table><tbody><tr><td>Rodent Langendorff Apparatus</td><td>Radnoti</td><td>130102EZ</td><td></td></tr><tr><td>24 G catheter</td><td>BD</td><td>381511</td><td></td></tr><tr><td>26 G needle on 1 mL syringe combo</td><td>BD</td><td>309597</td><td></td></tr><tr><td>26 G steel needle</td><td>BD</td><td>305111</td><td></td></tr><tr><td>5-0 Silk Suture</td><td>Ethicon</td><td>S1173</td><td></td></tr><tr><td>Bio Amp</td><td>ADInstruments</td><td>FE135</td><td></td></tr><tr><td>Bio Cable</td><td>ADInstruments</td><td>MLA1515</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>C4901-100G</td><td></td></tr><tr><td>Circulating heating water Bath</td><td>Haake</td><td>DC10</td><td></td></tr><tr><td>curved iris scissor</td><td>Medline</td><td>MDS10033Z</td><td></td></tr><tr><td>dissecting microscope</td><td>Nikon</td><td>SMZ-2B</td><td></td></tr><tr><td>find spring scissors</td><td>Kent</td><td>INS600127</td><td></td></tr><tr><td>Force Transducer</td><td>ADInstruments</td><td>MLT1030/D</td><td></td></tr><tr><td>glucose</td><td>Sigma-Aldrich</td><td>G8270-100G</td><td></td></tr><tr><td>Heparin</td><td>Sagent</td><td>400-01</td><td></td></tr><tr><td>High pressure tubing</td><td>Edwards Lifesciences</td><td>50P184</td><td></td></tr><tr><td>iris dressing forceps</td><td>Kent</td><td>INS650915-4</td><td></td></tr><tr><td>Jeweler-style curved fine forceps</td><td>Miltex</td><td>17-307-MLTX</td><td></td></tr><tr><td>KCl</td><td>Sigma-Aldrich</td><td>P3911-25G</td><td></td></tr><tr><td>KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>P0662-25G</td><td></td></tr><tr><td>MgSO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>M7506-500G</td><td></td></tr><tr><td>NaCl</td><td>Sigma-Aldrich</td><td>S9888-25G</td><td></td></tr><tr><td>NaHCO&lt;sub&gt;3&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>S6014-25G</td><td></td></tr><tr><td>Roller Pump</td><td>Gilson</td><td>Minipuls 3</td><td></td></tr><tr><td>straight dissecting scissors</td><td>Kent</td><td>INS600393-G</td><td></td></tr><tr><td>Temporary cardiac pacing wire</td><td>Ethicon</td><td>TPW30</td><td></td></tr><tr><td>Wide Range Force Transducer</td><td>ADInstruments</td><td>MLT1030/A</td><td></td></tr></tbody></table>

modified technique for the use of neonatal murine hearts in the langendorff preparation

The use of the ex-vivo retrograde perfused heart has long been a cornerstone of ischemia-reperfusion investigation since its development by Oskar Langendorff over a century ago. Although this technique has been applied to mice over the last 25 years, its use in this species has been limited to adult animals. Development of a successful method to consistently cannulate the neonatal murine aorta would allow for the systematic study of the isolated retrograde perfused heart during a critical period of cardiac development in a genetically modifiable and low-cost species. Modification of the Langendorff preparation enables cannulation and establishment of reperfusion in the neonatal murine heart while minimizing ischemic time. Optimization requires a two-person technique to permit successful cannulation of the newborn mouse aorta using a dissecting microscope and a modified commercially available needle. The use of this approach will reliably establish retrograde perfusion within 3 min. Because the fragility of the neonatal mouse heart and ventricular cavity size prevents direct measurement of intraventricular pressure generated using a balloon, use of a force transducer connected by a suture to the apex of the left ventricle to quantify longitudinal contractile tension is necessary. This method allows investigators to successfully establish an isolated constant-flow retrograde-perfused newborn murine heart preparation, permitting the study of developmental cardiac biology in an ex-vivo manner. Importantly, this model will be a powerful tool to investigate the physiological and pharmacological responses to ischemia-reperfusion in the neonatal heart.

P10 mice were used to model&#160;a timepoint in human infancy26,27. Fifteen isolated C57Bl/6 newborn mouse hearts were harvested and cannulated successfully. Hearts were perfused with a continuous flow of 2.5 mL min-1 of warmed oxygenated KHB. Metabolic parameters, including glucose extraction, oxygen consumption, lactate production, and physiological parameters such as heart rate, perfusion pressure, and coronary resistance, were measured. Surface ele...

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Modified Technique for the Use of Neonatal Murine Hearts in the Langendorff Preparation

The present protocol describes aortic cannulation and retrograde perfusion of the ex-vivo neonatal murine heart. A two-person strategy, using a dissecting microscope and a blunted small gauge needle, permits reliable cannulation. Quantification of longitudinal contractile tension is achieved using a force transducer connected to the apex of the left ventricle.

The use of the ex-vivo retrograde perfused heart has long been a cornerstone of ischemia-reperfusion investigation since its development by Oskar ...

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Research

JoVE Journal

Medicine

2.4K Views.  Vanderbilt University Medical Center. The present protocol describes aortic cannulation and retrograde perfusion of the ex-vivo neonatal murine heart. A two-person strategy, using a dissecting microscope and a blunted small gauge needle, permits reliable cannulation. Quantification of longitudinal contractile tension is achieved using a force transducer connected to the apex of the left ventricle.

Video: Modified Technique for the Use of Neonatal Murine Hearts in the Langendorff Preparation

Optical Mapping of Langendorff-perfused Rat Hearts

Optical mapping of the cardiac surface with voltage-sensitive fluorescent dyes has become an important tool to investigate electrical excitation in experimental models that range in scale from cell cultures to whole-organs[1, 2]. Using state-of-the-art optical imaging systems, generation and propagation of action potentials during normal cardiac rhythm or throughout initiation and maintenance of arrhythmias can be visualized almost instantly[1]. The latest commercially-available systems can provide information at exceedingly high spatiotemporal resolutions and were based on custom-built equipment initially developed to overcome the obstacles imposed by more conventional electrophysiological methods[1]. Advancements in high-resolution and high-speed complementary metal-oxide-semiconductor (CMOS) cameras and intensely-bright, light-emitting diodes (LEDs) as well as voltage-sensitive dyes, optics, and filters have begun to make electrical signal acquisition practical for cardiovascular cell biologists who are more accustomed to working with microscopes. Although the newest generation of CMOS cameras can acquire 10,000 frames per second on a 16,384 pixel array, depending on the type of sample preparation, long-established fluorescence acquisition technologies such as photodiode arrays, laser scanning systems, and cooled charged-coupled device (CCD) cameras still have some distinct advantages with respect to dynamic range, signal-to-noise ratio, and quantum efficiency[1, 3]. In the present study, Lewis rat hearts were perfused ex vivo with a crystalloid perfusate (Krebs-Henseleit solution) at 37&deg;C on a modified Langendorff apparatus. After a 20 minute stabilization period, the hearts were intermittently perfused with 11 mMol/L 2,3-butanedione monoxime to eliminate contraction-associated motion during image acquisition. For optical mapping, we loaded hearts with the fast-response potentiometric probe di-8-ANEPPS[4] (5 &mu;Mol/L) and briefly illuminated the preparation with 475&plusmn;15 nm excitation light. During a typical 2 second period of illumination, &gt;605 nm light emitted from the cardiac preparation was imaged with a high-speed CMOS camera connected to a horizontal macroscope. For this demonstration, hearts were paced at 300 beats per minute with a coaxial electrode connected to an isolated electrical stimulation unit. Simultaneous bipolar electrographic recordings were acquired and analyzed along with the voltage signals using readily-available software. In this manner, action potentials on the surface of Langendorff-perfused rat hearts can be visualized and registered with electrographic signals.

This article describes a high temporal and spatial resolution technique to optically image action potential movement on the surface of Langendorff-perfused rat hearts using a potentiometric dye (di-8-ANEPPS).

Optical mapping of the cardiac surface with voltage-sensitive fluorescent dyes has become an important tool to investigate electrical excitation in ...

Biology

Neonatal Cardiac Scaffolds: Novel Matrices for Regenerative Studies

The only definitive therapy for end stage heart failure is orthotopic heart transplantation. Each year, it is estimated that more than 100,000 donor hearts are needed for cardiac transplantation procedures in the United States1-2. Due to the limited numbers of donors, only approximately 2,400 transplants are performed each year in the U.S.2. Numerous approaches, from cell therapy studies to implantation of mechanical assist devices, have been undertaken, either alone or in combination, in an attempt to coax the heart to repair itself or to rest the failing heart3. In spite of these efforts, ventricular assist devices are still largely used for the purpose of bridging to transplantation and the utility of cell therapies, while they hold some curative promise, is still limited to clinical trials. Additionally, direct xenotransplantation has been attempted but success has been limited due to immune rejection. Clearly, another strategy is required to produce additional organs for transplantation and, ideally, these organs would be autologous so as to avoid the complications associated with rejection and lifetime immunosuppression. Decellularization is a process of removing resident cells from tissues to expose the native extracellular matrix (ECM) or scaffold. Perfusion decellularization offers complete preservation of the three dimensional structure of the tissue, while leaving the bulk of the mechanical properties of the tissue intact4. These scaffolds can be utilized for repopulation with healthy cells to generate research models and, possibly, much needed organs for transplantation. We have exposed the scaffolds from neonatal mice (P3), known to retain remarkable cardiac regenerative capabilities,5-8 to detergent mediated decellularization and we repopulated these scaffolds with murine cardiac cells. These studies support the feasibility of engineering a neonatal heart construct. They further allow for the investigation as to whether the ECM of early postnatal hearts may harbor cues that will result in improved recellularization strategies.

In these studies, we provide methodology for novel, neonatal, murine cardiac scaffolds for use in regenerative studies.

The only definitive therapy for end stage heart failure is orthotopic heart transplantation.  Each year, it is estimated that more than 100,000 donor ...

Bioengineering

Clonal Analysis of the Neonatal Mouse Heart using Nearest Neighbor Modeling

By replacing lost or dysfunctional myocardium, tissue regeneration is a promising approach to treat heart failure. However, the challenge of detecting bona fide heart regeneration limits the validation of potential regenerative factors. One method to detect new cardiomyocytes is multicolor lineage tracing with clonal analysis. Clonal analysis experiments can be difficult to undertake, because labeling conditions that are too sparse lack sensitivity for rare events such as cardiomyocyte proliferation, and diffuse labeling limits the ability to resolve clones. Presented here is a protocol to undertake clonal analysis of the neonatal mouse heart by using statistical modeling of nearest neighbor distributions to resolve cardiomyocyte clones. This approach enables resolution of clones over a range of labeling conditions and provides a robust analytical approach for quantifying cardiomyocyte proliferation and regeneration. This protocol can be adapted to other tissues and can be broadly used to study tissue regeneration.

Presented here is a protocol for using multicolor lineage tracing and nearest-neighbor modeling to identify clonally derived cardiomyocytes during growth and regeneration in mice. This approach is objective, works across different labeling conditions, and can be adapted to incorporate a variety of image analysis pipelines.

By replacing lost or dysfunctional myocardium, tissue regeneration is a promising approach to treat heart failure. However, the challenge of detecting ...

Developmental Biology

Myocardial Infarction in Neonatal Mice, A Model of Cardiac Regeneration

Myocardial infarction induced by coronary artery ligation has been used in many animal models as a tool to study the mechanisms of cardiac repair and regeneration, and to define new targets for therapeutics. For decades, models of complete heart regeneration existed in amphibians and fish, but a mammalian counterpart was not available. The recent discovery of a postnatal window during which mice possess regenerative capabilities has led to the establishment of a mammalian model of cardiac regeneration. A surgical model of mammalian cardiac regeneration in the neonatal mouse is presented herein. Briefly, postnatal day 1 (P1) mice are anesthetized by isoflurane and placed on an ice pad to induce hypothermia. After the chest is opened, and the left anterior descending coronary artery (LAD) is visualized, a suture is placed around the LAD to inflict myocardial ischemia in the left ventricle. The surgical procedure takes 10-15 min. Visualizing the coronary artery is crucial for accurate suture placement and reproducibility. Myocardial infarction and cardiac dysfunction are confirmed by triphenyl-tetrazolium chloride (TTC) staining and echocardiography, respectively. Complete regeneration 21 days post myocardial infarction is verified by histology. This protocol can be used to as a tool to elucidate mechanisms of mammalian cardiac regeneration after myocardial infarction.

This protocol describes a highly reproducible model of cardiac regeneration by surgical induction of myocardial infarction in the left ventricle of postnatal day 1 mice. The method involves induction of hypothermic anesthesia and ligation of the left anterior descending coronary artery.

Myocardial infarction induced by coronary artery ligation has been used in many animal models as a tool to study the mechanisms of cardiac repair and ...

Isolating Primary Melanocyte-like Cells from the Mouse Heart

We identified a novel population of melanocyte-like cells (also known as cardiac melanocytes) in the hearts of mice and humans that contribute to atrial arrhythmia triggers in mice. To investigate the electrical and biological properties of cardiac melanocytes we developed a procedure to isolate them from mouse hearts that we derived from those designed to isolate neonatal murine cardiomyocytes. In order to obtain healthier cardiac melanocytes suitable for more extensive patch clamp or biochemical studies, we developed a refined procedure for isolating and plating cardiac melanocytes based on those originally designed to isolate cutaneous melanocytes. The refined procedure is demonstrated in this review and produces larger numbers of healthy melanocyte-like cells that can be plated as a pure population or with cardiomyocytes.

In this protocol, we identified a novel population of melanocyte-like cells (also known as cardiac melanocytes) in the hearts of mice and humans that contribute to atrial arrhythmia triggers in mice.&#160;

We identified a novel population of melanocyte-like cells (also known as cardiac melanocytes) in the hearts of mice and humans that contribute to atrial ...

Isolation and Culture of Neonatal Mouse Cardiomyocytes

Cultured neonatal cardiomyocytes have long been used to study myofibrillogenesis and myofibrillar functions. Cultured cardiomyocytes allow for easy investigation and manipulation of biochemical pathways, and their effect on the biomechanical properties of spontaneously beating cardiomyocytes.	
The following 2-day protocol describes the isolation and culture of neonatal mouse cardiomyocytes. We show how to easily dissect hearts from neonates, dissociate the cardiac tissue and enrich cardiomyocytes from the cardiac cell-population. We discuss the usage of different enzyme mixes for cell-dissociation, and their effects on cell-viability. The isolated cardiomyocytes can be subsequently used for a variety of morphological, electrophysiological, biochemical, cell-biological or biomechanical assays. We optimized the protocol for robustness and reproducibility, by using only commercially available solutions and enzyme mixes that show little lot-to-lot variability. We also address common problems associated with the isolation and culture of cardiomyocytes, and offer a variety of options for the optimization of isolation and culture conditions.

Primary mouse cardiomyocyte cultures are one of the pivotal tools for the investigation of myofibrillar organization and function. The following protocol describes the isolation and culture of primary cardiomyocytes from neonatal mouse hearts. The resulting cardiomyocyte cultures may be subsequently used for a variety of biomechanical, biochemical and cell-biological assays.

Cultured neonatal cardiomyocytes have long been used to study  myofibrillogenesis and myofibrillar functions. Cultured cardiomyocytes allow  for easy ...

Modifications of the Langendorff Method for Simultaneous Isolation of Atrial and Ventricular Myocytes from Adult Mice

A single cardiomyocyte is a vital tool in the cellular and subcellular level studies of cardiac biology and diseases as a fundamental unit of contraction and electrical activity. Hence, isolating viable, high-quality cardiomyocytes from the heart is the initial and most crucial experimental step. Comparing the various protocols for isolating the cardiomyocytes of adult mice, the Langendorff retrograde perfusion is the most successful and reproducible method reported in the literature, especially for isolating ventricular myocytes. However, isolating quality atrial myocytes from the perfused heart remains challenging, and few successful isolation reports are available. Solving this complicated problem is extremely important because apart from ventricular disease, atrial disease accounts for a large part of heart diseases. Therefore, further investigations on the cellular level to reveal the mechanisms are warranted. In this paper, a protocol based on the Langendorff retrograde perfusion method is introduced and some modifications in the depth of aorta cannulation and the steps that may affect the digestion process to isolate atrial and ventricular myocytes were simultaneously made. Moreover, the isolated cardiomyocytes are confirmed to be amenable to patch clamp investigation.

This protocol describes modifications of the Langendorff method including the depth of aorta cannulation for simultaneous Isolation of atrial and ventricular myocytes from adult mice.

A single cardiomyocyte is a vital tool in the cellular and subcellular level studies of cardiac biology and diseases as a fundamental unit of contraction ...

A Simple and Effective Method to Consistently Isolate Mouse Cardiomyocytes

The need for reproducible yet technically simple methods yielding high-quality cardiomyocytes is essential for research in cardiac biology. Cellular and molecular functional experiments (e.g., contraction, electrophysiology, calcium cycling, etc.) on cardiomyocytes are the gold standard for establishing mechanism(s) of disease. The mouse is the species of choice for functional experiments and the described technique is specifically for the isolation of mouse cardiomyocytes. Previous methods requiring a Langendorff apparatus require high levels of training and precision for aortic cannulation, often resulting in ischemia. The field is shifting toward Langendorff-free isolation methods that are simple, are reproducible, and yield viable myocytes for physiological data acquisition and culture. These methods greatly diminish ischemia time compared to aortic cannulation and result in reliably obtained cardiomyocytes. Our adaptation to the Langendorff-free method includes an initial perfusion with ice-cold clearing solution, use of a stabilizing platform that ensures a steady needle during perfusion, and additional digestion steps to ensure reliably obtained cardiomyocytes for use in functional measurements and culture. This method is simple and quick to perform and requires little technical skill.

The gold standard in cardiology for cellular and molecular functional experiments are cardiomyocytes. This article describes adaptations to the non-Langendorff technique to isolate mouse cardiomyocytes.

The need for reproducible yet technically simple methods yielding high-quality cardiomyocytes is essential for research in cardiac biology. Cellular and ...

An Antegrade Perfusion Method for Cardiomyocyte Isolation from Mice

In basic research using mouse heart, isolating viable individual cardiomyocytes is a crucial technical step to overcome. Traditionally, isolating cardiomyocytes from rabbits, guinea pigs or rats has been performed via retrograde perfusion of the heart with enzymes using a Langendorff apparatus. However, a high degree of skill is required when this method is used with a small mouse heart. An antegrade perfusion method that does not use a Langendorff apparatus was recently reported for the isolation of mouse cardiomyocytes. We herein report a complete protocol for the improved antegrade perfusion of the excised heart to isolate individual heart cells from adult mice (8 - 108 weeks old). Antegrade perfusion is performed by injecting perfusate near the apex of the left ventricle of the excised heart, the aorta of which was clamped, using an infusion pump. All procedures are carried out on a pre-warmed heater mat under a microscope, which allows for the injection and perfusion processes to be monitored. The results suggest that ventricular and atrial myocytes, and fibroblasts can be well isolated from a single adult mouse simultaneously.

We developed a simple method for isolating high quality individual mouse heart cells by the antegrade perfusion technique. This method is Langendorff-free and useful for isolating ventricular and atrial myocytes or interstitial cells, such as cardiac fibroblasts or progenitors.

In basic research using mouse heart, isolating viable individual cardiomyocytes is a crucial technical step to overcome. Traditionally, isolating ...

Langendorff Retrograde Heart Perfusion: An Ex Vivo Technique to Perfuse Isolated Mouse Heart

After euthanizing an 8- to 10-week-old male C57BL/6 mouse, use tissue forceps to lift the skin of the xiphoid. Then, using tissue scissors, make a minor lateral incision through the skin. Perform a blunt dissection between the skin and fascia. Extend the skin incision toward the axillae in a V-shape on both sides and continue the incision through the rib cage.
Then, using tissue forceps, clamp the sternum and deflect the rib cage upward to fully expose the heart and lungs. Using curved forceps, peel off the pericardium. If the thymus gland covers the great vessels, use two curved forceps to tear the thymus gland toward both sides. Then, gently pull the base of the heart toward the tail until the aorta and its branch arteries are visible as a Y-shaped blood vessel.
Transect the aorta at the left common carotid artery. Then, cut the brachiocephalic artery. Excise the heart and immediately immerse it in a Petri dish containing Tyrode&#39;s solution to wash and pump out the residual blood. Then, transfer the heart to another Petri dish containing solution 1.
Under a stereomicroscope, using fine iris scissors, trim any surplus tissue. Expel air bubbles from the syringe. Then, with the assistance of two straight tying forceps, perform retrograde aortic cannulation, taking care that the whole cannulation process is performed under the liquid surface. Adjust the cannulation depth such that the cannula tip is in the ascending aorta, taking care not to penetrate the aortic valves.
Then, with a pre-knot 3-0 suture, ligate the aorta to the cannula notch. Gently inject solution 1 from the syringe to flush out the residual blood. Then, connect the cannulated heart to the Langendorff apparatus, taking care not to introduce any air bubbles into the heart. After connecting the cannulated heart to the Langendorff apparatus, perfuse the heart with solution 1 for approximately 2 minutes.