TMAM is supported by NIH grant P30GM127607 and American Heart Association grant 16SDG29950012. RB is supported by P01HL78825 and UM1HL113530.

All animal procedures were in accordance with the institutional guidelines of the University of Louisville and approved by the Institutional Animal Care and Use Committee.
1. Preparation for Slicing (One Day Before Slicing)
<ol>
	<li>Preparation of the vibrating microtome
	<ol>
		<li>Place the ceramic blade into its holder by following these steps: after carefully unwrapping the blade, place the sharp edge first into the slot of the blade tool. Then, fit the blade into the holder by loosening the two screws on the arms of the holder and slide the blade under each washer and push it firmly back against the rear stops. Tighten the screws to secure the blade. 
		NOTE: The screws should not be overtightened.</li>
		<li>Calibrate the blade using the calibration protocol and tools according to the manufacturer&#39;s protocol. 
		NOTE: Briefly, in order to facilitate alignment of the blade with the axis of travel and to minimize the Z axis deflections, the vibrating microtome uses a demountable calibration device. When the calibration device is plugged into the instrument its presence will automatically be detected, and the vibrating microtome will take control of adjusting the amplitude and frequency settings of the blade to a magnitude which best allows the adjustment of the blade alignment error.
		<ol>
			<li>Press the Slice button to start the alignment process which will automatically move the blade so that the cutting edge is in optimal position relative to the calibration device for best alignment evaluation. Follow the on-screen instructions to make adjustments to the blade using the provided screwdriver to ensure that the Z-alignment is within 1 &#181;m.</li>
		</ol>
		</li>
		<li>Autoclave the inner bath and all the metal parts of the vibrating microtome.</li>
	</ol>
	</li>
	<li>Autoclave the large metal trays (for collecting the slices and soaking the electrical stimulation plate covers), any glass containers, regular rectangle blades, forceps, scissors, printer timing belt (cut them into 6 mm wide pieces), metal washers and 5 L of double distilled water (ddH2O).</li>
	<li>Sterilize one 1 L jar, one 500 mL jar and one plastic tray for the heart in situ and in vitro perfusion with the cardioplegic solution.</li>
	<li>Prepare 2 L of the slicing Tyrode&#39;s solution in a 2 L beaker.
	<ol>
		<li>For 1 L of the slicing Tyrode&#39;s solution, mix 3 g/L 2,3-butanedione monoxime (BDM), 140 mM NaCl (8.18 g), 6 mM KCl (0.447 g), 10 mM D-glucose (1.86 g), 10 mM HEPES (2.38 g), 1 mM MgCl2 (1 mL of 1 M solution), 1.8 mM CaCl2 (1.8 mL of 1 M solution), up to 1 L of ddH2O. Adjust the pH to 7.40 using NaOH. Then, filter the solution using a 1,000 mL, 0.22 &#181;m, vacuum filter/storage system and store at 4 &#176;C overnight.</li>
	</ol>
	</li>
	<li>Prepare 4 L of the cardioplegic solution.
	<ol>
		<li>For 1 L of the cardioplegic solution, mix 110 mM NaCl (6.43 g), 1.2 mM CaCl2 (1.2 mL of 1 M solution), 16 mM KCl (1.19 g), 16 mM MgCl2 (3.25 g), 10 mM NaHCO3 (0.84 g), 1 U/mL heparin, up to 1 L of ddH2O. Adjust the pH to 7.40 using NaOH. Then, filter the solution using a 1,000 mL, 0.22 &#181;m, vacuum filter/storage system and store at 4 &#176;C overnight. 
		NOTE: Add 1,000 U/L heparin in cardioplegia solution the day of slicing (see step 2.1).</li>
	</ol>
	</li>
	<li>Freshly prepare 500 mL of culture medium in the original bottle in a biosafety cabinet: mix medium 199, 1x insulin-transferrin-selenium (ITS) supplement, 10% fetal bovine serum (FBS), 5 ng/mL vascular endothelial growth factor (VEGF), 10 ng/mL FGF-basic, and 2x antibiotic-antimycotic. Filter sterilize through the 0.22 &#181;m, vacuum filter/storage system and store at 4 &#176;C overnight. 
	NOTE: Do not adjust pH, add VEGF and FGF freshly before use.</li>
	<li>Prepare 4% agar gel plates.
	<ol>
		<li>Add 200 mL of sterilized ddH2O in a 500 mL sterilized flask. Weigh 8 g of agarose, and gently pour into the flask. Boil for 2 min in a microwave, then cool for 5 min at room temperature.</li>
		<li>Pour 25 mL in each 100 mm Petri dish in a biosafety cabinet (prepare 6 dishes) and wait for 1 h to solidify. Then, seal the plates with paraffin film, wrap with clean wrap, and store at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Prepare 4 L of 70% ethanol by diluting the 200-proof absolute ethanol in autoclaved ddH2O.</li>
	<li>Prepare 2x antibiotic-antimycotic in sterilized ddH2O: mix 980 mL of sterilized ddH2O with 20 mL of antibiotic-antimycotic under the biosafety cabinet.</li>
</ol>
2. Pig Heart Perfusion
<ol>
	<li>Add 1 mL of 1,000 U/mL heparin to each 1 L of cardioplegic solution. Keep the solution on ice. 
	NOTE: At least 3 L are needed for the perfusion and preservation of the heart during the transfer to the tissue culture room.</li>
	<li>Take the following items to the pig surgery room: one large ice bucket full of ice, one sterilized metal tray (keep on ice bucket for heart perfusion), one sterilized 500 mL jar, one 1 L jar (to transfer the heart in cardioplegia solution back to the tissue culture room, on ice), and one sterilized plastic tray full of ice (to keep cardioplegic solution on ice).</li>
	<li>Prepare an in-situ heart perfusion system containing a 3-way stopcock, a 60 mL syringe, an 18 G butterfly needle catheter, and extension tubing to the cardioplegic solution reservoir.</li>
	<li>Anesthetize the Yorkshire male pig (2&#8722;3 month-old, 20&#8722;25 kg) first with an intramuscular injection of a ketamine (20 mg/kg)/xylazine (2 mg/kg) cocktail. Confirm proper anesthesia by the loss of jaw tension. Then, transfer the pig to the surgery room, intubate and mechanically ventilate the lungs as previously described14.</li>
	<li>Maintain anesthesia with (1.5&#8722;2%) isoflurane.</li>
	<li>Place the animal on the right lateral position and shave the fur from the chest area. Clean the skin with alcohol scrub, then sterilize the surgical site with 10% (w/v) povidone-iodine solution.</li>
	<li>Open the chest by left thoracotomy incision using a scalpel at the 4th intercostal space, and use a chest retractor between ribs to hold the chest open and expose the heart. 
	NOTE: Use all sterilized instruments for this procedure.</li>
	<li>Insert the butterfly needle catheter into the left atrium and fix the needle with a purse string closure. After intravenous injection of a bolus dose of heparin (100 U/kg), start to perfuse the heart in situ with 500 mL of ice-cold cardioplegic solution (to slow down the heart rate) via the left atrial catheter.</li>
	<li>Deeply anesthetize the pig with 5% isoflurane. Quickly excise the heart out of the chest with surgical scissors. Cannulate the aorta with an aortic cannula and perfuse the heart in vitro with another 1 L of ice-cold cardioplegic solution (100 mL/min) to flush out vasculature blood.</li>
	<li>Following heart perfusion, keep the heart in a 1 L jar filled with ice-cold cardioplegia solution and keep on ice during the transfer to the tissue culture room.</li>
</ol>
3. Pig Heart Tissue Slicing
<ol>
	<li>Set up the tissue bath on the vibratome, add ice to the tissue bath cooling jacket, then add Tyrode&#39;s solution into the tissue bath. Set up 1 L plastic jar to collect the disposal of melted ice from the tissue bath cooling jacket.</li>
	<li>Under the biosafety cabinet, transfer the pig heart to a tray containing 1 L of fresh cold cardioplegic solution.</li>
	<li>Dissect the heart to isolate the left ventricle using retractable sterile scalpels. Then, cut the left ventricle into blocks of 1&#8722;2 cm3 each using razor rectangle blades. Use one piece for slicing and keep the remaining pieces in a 50 mL tube in cold Tyrode&#39;s solution on ice for cutting later, if needed. 
	NOTE: Massage the tissue before slicing with hands, especially if this is the second block. Massaging helps to restore the correct muscle fiber configuration and prevents any stiffness within the heart block.</li>
	<li>Add 1&#8722;2 drops of tissue glue (Table of Materials) to the metal sample holder and stick a piece of the 4% agar block with a surface area of approximately 1 cm2.</li>
	<li>Add 1&#8722;2 drops of tissue glue on the agar.</li>
	<li>Stick the heart block to the agar with the cardiac epicardium side facing down on the tissue glue and make sure it is as flat as possible. Then, transfer the tissue holder with the heart block to its position in the slicing bath of the vibratome.</li>
	<li>Attach the oxygen tube to the slicing bath and the metal tray filled with Tyrode&#39;s solution for collecting the slices (oxygenated Tyrode&#39;s bath). Then, add 40 &#181;m cell strainers in the metal tray to collect the slices after cutting.</li>
	<li>Adjusting the slicing position
	<ol>
		<li>Using the vibratome operating software and the dashboard, adjust the height of the blade/sample. Select the height button and follow the on-screen instructions for adjusting the height. Ideally, have the blade close to the top of the tissue but below the papillary muscles and make sure there is liquid covering the tissue and the blade before slicing.</li>
		<li>Adjust where to begin slicing the tissue by selecting the advance button. Press the slice button and increase the speed using the knob to move the blade towards the edge of the tissue. Then, press slice again to stop, and press the advance button to inactivate the process.</li>
		<li>Adjust the cutting parameters: advance speed = 0.03 mm/s, vibration frequency = 80 Hz, and horizontal vibration amplitude = 2 mm. Then, start slicing by selecting the slice button.</li>
		<li>Let the vibratome slice until it reaches the end of the tissue but before it hits the back end of the specimen holder. At this point, press slice again to stop. Hit the Return button to go back to the start position.</li>
		<li>Select auto repeat (clicking it twice) which will allow the vibrating microtome to auto repeat the slicing process for up to 99 times.</li>
	</ol>
	</li>
	<li>Collecting slices
	<ol>
		<li>Wait until slices are full length and appear good (after getting past the papillary muscle layers), then start collecting the slices.</li>
		<li>Collect the slices using a plastic Pasteur pipette filled with cold Tyrode&#39;s solution to gently grab the tissue from the bath. If necessary, use forceps and spring scissors to dissociate the slice from the heart block if the tissue is still attached.</li>
		<li>Transfer the slice to one cell strainer in the oxygenated Tyrode&#39;s bath (see step 3.7) and use the liquid in the plastic Pasteur pipette to get the tissue to lie flat on one of the cell strainers, then add a metal washer on the top to hold the tissue down. 
		NOTE: The slices need to be incubated at least for 1 h in the Tyrode&#39;s solution before processing (this is for the BDM to relax the tissue).</li>
	</ol>
	</li>
</ol>
4. Culturing Heart Slices
<ol>
	<li>Preparing the slices for culture
	<ol>
		<li>Trim the slice from the edges to avoid any uneven edges. Then, glue it from both ends to sterilized polyurethane 6 mm wide printer timing belt with metal wires embedded using tissue glue.</li>
		<li>Transfer the supported heart slices to 6 well plates that contain 6 mL of culture medium in each well.</li>
		<li>Place the stimulation plate cover (Table of Materials) on the top of the 6 well plate and connect the cover to the cell culture electrical stimulator (Table of Materials). Adjust the stimulator to electrically stimulate the heart slices at 10 V, 1.2 Hz continuously all the time. 
		NOTE: After plugging in the stimulation, the heart slices will start to beat, and this movement will be visually obvious.</li>
		<li>Transfer the plates to an incubator and maintain at 37 &#176;C with humidified air and 5% CO2.</li>
	</ol>
	</li>
	<li>Change culture medium 3x per day and add 6 mL of culture medium per well during each media change. 
	NOTE: Culture medium is oxygenated for 5 min prior to each media change. Medium must be changed at least 1x per day; this is okay on weekends if necessary. Two days of only 1 media change per day is the maximum that is tested.</li>
	<li>Change the stimulation plate cover every day to prevent release of toxic graphite particles in the culture medium (usually during the mid-day media change).
	<ol>
		<li>Remove the stimulation plate cover from the culture dish and insert the white foam plug where the cover connects to the cable for the cell culture electrical stimulator, to prevent water damage to the electric circuit.</li>
		<li>Place the plate cover in a bath of autoclaved water with 2x antibiotic-antimycotic. Keep it in this bath (i.e., rinsing bath) overnight.</li>
		<li>The next day, move the plate cover into a 70% ethanol bath for 5&#8722;15 min to decontaminate. Shake off as much liquid from the device before switching baths.</li>
		<li>Then, transfer the plate cover to the third and final bath (i.e., clean bath), which consists of autoclaved water and 2x antibacterial-antimycotic, to rinse any remaining ethanol traces.</li>
		<li>After rinsing the plate cover in the clean bath, use a clean lint-free wipe (sprayed down with ethanol) to dry off any residual water on the plastic parts, then remove the white foam piece and carefully place the plate cover back on the culture plate. 
		NOTE: Do not touch the black graphite electrodes that go into the well, as the device will no longer be sterile.</li>
		<li>Using sterile forceps, make sure the tissue is in the center of the well so the plate cover does not touch the tissue. Ensure that the curved side of the plate cover matches up with the angled side of the plate and that the square corners line up. 
		NOTE: To minimize contamination of the stimulation plate covers, keep them in clean and empty 6 well plates after finishing the experiment.</li>
	</ol>
	</li>
	<li>Perform an MTT assay, calcium measurements and contractile function assessments on fresh pig heart slices (day 0), and after 6 days in culture according to Ou et al.13.</li>
</ol>

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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=Cardiovascular+Toxic+Effects+of+Targeted+Cancer+Therapies.">Cardiovascular Toxic Effects of Targeted Cancer Therapies.</a> The New England Journal of Medicine. 375 (15), 1457-1467 (2016).</li><li>Robertson, C., Tran, D. D., George, S. 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TMAM holds equity in Tenaya Therapeutics. The other authors report no conflicts.

Here we describe the detailed video protocol for our recently published method for simplified medium throughput (processes up to 48 slices/device) method that enables culture of pig heart slices for a period sufficiently long to test acute cardiotoxicity13. The proposed conditions mimic the environment of the heart, including frequency of electrical stimulation, nutrient availability, and intermittent oxygenation. We attribute the prolonged viability of heart slices in our biomimetic stimulated cu...

Drug induced cardiotoxicity is a major cause of market withdrawal1. In the last decade of the 20th century, eight non-cardiovascular drugs were withdrawn from the market as they resulted in sudden death due to ventricular arrhythmias2. In addition, several anti-cancer therapies (while in many cases effective) can lead to several cardiotoxic effects including cardiomyopathy and arrhythmias. For example, both traditional (e.g., anthracyclines and radiation) and targeted (e.g., trastuzumab) breast cancer therapies can result in cardiovascular complications in a subset of patients3. A close collaboration between cardiologists and oncologists (via the emerging field of &#34;cardio-oncology&#34;) has helped make these complications manageable ensuring that patients can be treated effectively2. Less clear are the cardiovascular effects of newer agents, including Her2 and PI3K inhibitors, especially when therapies are used in combination. Therefore, there is a growing need for reliable preclinical screening strategies for cardiovascular toxicities associated with emerging anti-cancer therapies prior to human clinical trials. The lack of availability of culture systems for human heart tissue that is functionally and structurally viable for more than 24 h is a limiting factor for reliable cardiotoxicity testing. Therefore, there is an urgent need to develop a reliable system for culturing human heart tissue under physiologic conditions for testing drug toxicity.
The recent move towards the use of human induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) in cardiotoxicity testing has provided a partial solution to address this issue; however, the immature nature of the hiPSC-CMs and the loss of tissue integrity compared to multicellular nature of the heart tissue are major limitations of this technology4. A recent study has partially overcome this limitation through fabrication of cardiac tissues from hiPSC-CMs on hydrogels and subjecting them to gradual increase in electrical stimulation over time5. However, their electromechanical properties did not achieve the maturity seen in the adult human myocardium. Moreover, the heart tissue is structurally more complicated, being composed of various cell types including endothelial cells, neurons and various types of stromal fibroblasts linked together with a very specific mixture of extracellular matrix proteins6. This heterogeneity of the non-cardiomyocyte cell population7,8,9 in the adult mammalian heart is a major obstacle in modeling heart tissue using individual cell types. These major limitations highlight the importance of developing methods to enable culture of intact cardiac tissue for optimal studies involving physiological and pathological conditions of the heart5.
Culturing human heart slices is a promising model of intact human myocardium. This technology provides access to a complete 3D multicellular system that is similar to the human heart tissue which could reliably reflect the physiological or pathological conditions of the human myocardium. However, its use has been severely limited by the short period of viability in culture, which does not extend beyond 24 h using the most robust protocols reported until 201810,11,12. This limitation was due to multiple factors including the use of air-liquid interface to culture the slices, and the use of a simple culture medium that does not support the high energetic demands of the cardiac tissue. We have recently developed a submerged culture system which is able to provide continuous electrical stimulation and optimized the culture media components to keep cardiac tissue slices viable for up to 6 days13. This culture system has the potential to become a powerful predictive human in situ model for acute cardiotoxicity testing to close the gap between preclinical and clinical testing results. In the current article, we are detailing the protocol for slicing and culturing the heart slices using a pig heart as an example. The same process is applied to human, dog, sheep, or cat hearts. With this protocol, we are hoping to spread the technology to other laboratories in the scientific community.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1000ml, 0.22&#181;m, Vacuum Filter/Storage Systems</td>
			<td>VWR</td>
			<td>28199-812</td>
			<td></td>
		</tr>
		<tr>
			<td>2,3-Butanedione monoxime (BDM)</td>
			<td>Fisher</td>
			<td>AC150375000</td>
			<td></td>
		</tr>
		<tr>
			<td>500ml, 0.22&#181;m, Vacuum Filter/Storage Systems</td>
			<td>VWR</td>
			<td>28199-788</td>
			<td></td>
		</tr>
		<tr>
			<td>6-well C-Dish Cover (electrical-stimulation-plate-cover)</td>
			<td>Ion Optix</td>
			<td>CLD6WFC</td>
			<td></td>
		</tr>
		<tr>
			<td>6-well plates</td>
			<td>Fisher</td>
			<td>08-772-1B</td>
			<td></td>
		</tr>
		<tr>
			<td>Agarose</td>
			<td>Bioline USA</td>
			<td>BIO-41025</td>
			<td></td>
		</tr>
		<tr>
			<td>Antibiotic-Antimycotic</td>
			<td>Thermo</td>
			<td>15-240-062</td>
			<td></td>
		</tr>
		<tr>
			<td>C-Pace EM (cell-culture-electrical-stimulator)</td>
			<td>Ion Optix</td>
			<td>CEP100</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium Chloride (CaCl2)</td>
			<td>Fisher</td>
			<td>C79-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Ceramic Blades for Vibrating Microtome</td>
			<td>Campden Instruments</td>
			<td>7550-1-C</td>
			<td></td>
		</tr>
		<tr>
			<td>Cooley Chest Retractor</td>
			<td>Millennium Surgical</td>
			<td>63-G5623</td>
			<td></td>
		</tr>
		<tr>
			<td>D-Glucose</td>
			<td>Fisher</td>
			<td>D16-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Disposable Scalpel #20</td>
			<td>Biologyproducts.com</td>
			<td>DS20X</td>
			<td></td>
		</tr>
		<tr>
			<td>Falcon Cell Strainers, Sterile, Corning</td>
			<td>VWR</td>
			<td>21008-952</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum</td>
			<td>Thermo</td>
			<td>A3160502</td>
			<td></td>
		</tr>
		<tr>
			<td>Graefe Forceps</td>
			<td>Fisher</td>
			<td>NC9475675</td>
			<td></td>
		</tr>
		<tr>
			<td>Heparin sodium salt</td>
			<td>Sigma-Aldrich</td>
			<td>H3149-50KU</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Fisher</td>
			<td>BP310-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Histoacryl BLUE Tissue glue</td>
			<td>Amazon</td>
			<td></td>
			<td><a href="https://www.amazon.com/HISTOACRYL-FLEXIBLE-1051260P-Aesculap-Adhesive/dp/B074WB5185/" target="_blank">https://www.amazon.com/HISTOACRYL-FLEXIBLE-1051260P-Aesculap-Adhesive/dp/B074WB5185/</a></td>
		</tr>
		<tr>
			<td>Iris Spring scissors</td>
			<td>Fisher</td>
			<td>NC9019530</td>
			<td></td>
		</tr>
		<tr>
			<td>Iris Straight Scissors</td>
			<td>Fisher</td>
			<td>731210</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane, USP</td>
			<td>Piramal</td>
			<td>NDC 66794-017-25</td>
			<td></td>
		</tr>
		<tr>
			<td>ITS Liquid Media Supplement</td>
			<td>Sigma-Aldrich</td>
			<td>I3146-5ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Ketamine HCl (500 mg/10 mL)</td>
			<td>West-Ward</td>
			<td>NDC 0143-9508</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium Chloride (MgCl2)</td>
			<td>Fisher</td>
			<td>M33-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Mayo SuperCut Surgical Scissors</td>
			<td>AROSurgical Instruments Corporation</td>
			<td>AROSuperCut&#8482; 07.164.17</td>
			<td></td>
		</tr>
		<tr>
			<td>Medium 199, Earle&#39;s Salts</td>
			<td>Thermo</td>
			<td>11-150-059</td>
			<td></td>
		</tr>
		<tr>
			<td>Oxygen regulator</td>
			<td>Praxair</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Oxygen tanks -</td>
			<td>Praxair</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Plastic Pasteur pipettes</td>
			<td>Fisher</td>
			<td>13-711-48</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium Chloride (KCl)</td>
			<td>Fisher</td>
			<td>AC193780010</td>
			<td></td>
		</tr>
		<tr>
			<td>Printer Timing Belt</td>
			<td>Amazon</td>
			<td></td>
			<td><a href="https://www.amazon.com/Uxcell-a14081200ux0042-PRINTER-Precision-Timing/dp/B00R1J3KDC/" target="_blank">https://www.amazon.com/Uxcell-a14081200ux0042-PRINTER-Precision-Timing/dp/B00R1J3KDC/</a></td>
		</tr>
		<tr>
			<td>Razor rectangle blades</td>
			<td>Fisher</td>
			<td>12-640</td>
			<td></td>
		</tr>
		<tr>
			<td>Recombinant Human FGF basic</td>
			<td>R&#38;D Systems</td>
			<td>233-FB-025/CF</td>
			<td></td>
		</tr>
		<tr>
			<td>Recombinant Human VEGF</td>
			<td>R&#38;D Systems</td>
			<td>293-VE-010/CF</td>
			<td></td>
		</tr>
		<tr>
			<td>Retractable scalpels</td>
			<td>Fisher</td>
			<td>22-079-716</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Bicarbonate (NaHCO3)</td>
			<td>Fisher</td>
			<td>AC217125000</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Chloride (NaCl)</td>
			<td>Fisher</td>
			<td>AC327300010</td>
			<td></td>
		</tr>
		<tr>
			<td>Vibrating Microtome</td>
			<td>Campden Instruments</td>
			<td>7000 SMZ-2</td>
			<td></td>
		</tr>
		<tr>
			<td>Xylazine HCl (100 mg/mL)</td>
			<td>Heartland Veterinary Supply</td>
			<td>NADA 139-236</td>
			<td></td>
		</tr>
	</tbody>
</table>

slicing and culturing pig hearts under physiological conditions

Many novel drugs fail in clinical studies due to cardiotoxic side effects as the currently available in vitro assays and in vivo animal models poorly predict human cardiac liabilities, posing a multi-billion-dollar burden on the pharmaceutical industry. Hence, there is a worldwide unmet medical need for better approaches to identify drug cardiotoxicity before undertaking costly and time consuming &#39;first in man&#39; trials. Currently, only immature cardiac cells (human induced pluripotent stem cell-derived cardiomyocytes [hiPSC-CMs]) are used to test therapeutic efficiency and drug toxicity as they are the only human cardiac cells that can be cultured for prolonged periods required to test drug efficacy and toxicity. However, a single cell type cannot replicate the phenotype of the complex 3D heart tissue which is formed of multiple cell types. Importantly, the effect of drugs needs to be tested on adult cardiomyocytes, which have different characteristics and toxicity responses compared to immature hiPSC-CMs. Culturing human heart slices is a promising model of intact human myocardium. This technology provides access to a complete multicellular system that mimics the human heart tissue and reflects the physiological or pathological conditions of the human myocardium. Recently, through optimization of the culture media components and the culture conditions to include continuous electrical stimulation at 1.2 Hz and intermittent oxygenation of the culture medium, we developed a new culture system setup that preserves viability and functionality of human and pig heart slices for 6 days in culture. In the current protocol, we are detailing the method for slicing and culturing pig heart as an example. The same protocol is used to culture slices from human, dog, sheep, or cat hearts. This culture system has the potential to become a powerful predictive human in situ model for acute cardiotoxicity testing that closes the gap between preclinical and clinical testing results.

Using a commercially available cell culture electrical stimulator that can accommodate eight 6 well plates at once, we emulated the adult cardiac milieu by inducing electrical stimulation at the physiological frequency (1.2 Hz), and screened for the fundamental medium components to prolong the duration of functional pig heart slices in culture13. Since pig and human hearts are similar in size and anatomy15, we developed a biomimetic heart sl...

Watch this Scientific Journal Video about Slicing and Culturing Pig Hearts under Physiological Conditions at JoVE.com

Slicing and Culturing Pig Hearts under Physiological Conditions

This protocol describes how to slice and culture heart tissue under physiological conditions for 6 days. This culture system could be used as a platform for testing the efficacy of novel heart failure therapeutics as well as reliable testing of acute cardiotoxicity in a 3D heart model.

Many novel drugs fail in clinical studies due to cardiotoxic side effects as the currently available in vitro assays and in vivo animal models poorly ...

slicing-and-culturing-pig-hearts-under-physiological-conditions

Universidad de Cartagena UDEC

Research

JoVE Journal

Medicine

15.0K Views.  University of Louisville. This protocol describes how to slice and culture heart tissue under physiological conditions for 6 days. This culture system could be used as a platform for testing the efficacy of novel heart failure therapeutics as well as reliable testing of acute cardiotoxicity in a 3D heart model.

Video: Slicing and Culturing Pig Hearts under Physiological Conditions

Assessment of Cardiac Function and Energetics in Isolated Mouse Hearts Using 31P NMR Spectroscopy

Bioengineered mouse models have become powerful research tools in determining causal relationships between molecular alterations and models of cardiovascular disease. Although molecular biology is necessary in identifying key changes in the signaling pathway, it is not a surrogate for functional significance. While physiology can provide answers to the question of function, combining physiology with biochemical assessment of metabolites in the intact, beating heart allows for a complete picture of cardiac function and energetics. For years, our laboratory has utilized isolated heart perfusions combined with nuclear magnetic resonance (NMR) spectroscopy to accomplish this task. Left ventricular function is assessed by Langendorff-mode isolated heart perfusions while cardiac energetics is measured by performing 31P magnetic resonance spectroscopy of the perfused hearts. With these techniques, indices of cardiac function in combination with levels of phosphocreatine and ATP can be measured simultaneously in beating hearts. Furthermore, these parameters can be monitored while physiologic or pathologic stressors are instituted. For example, ischemia/reperfusion or high workload challenge protocols can be adopted. The use of aortic banding or other models of cardiac pathology are apt as well. Regardless of the variants within the protocol, the functional and energetic significance of molecular modifications of transgenic mouse models can be adequately described, leading to new insights into the associated enzymatic and metabolic pathways. Therefore, 31P NMR spectroscopy in the isolated perfused heart is a valuable research technique in animal models of cardiovascular disease.

Assessment of Cardiac Function and Energetics in Isolated Mouse Hearts Using 31P NMR Spectroscopy

Langendorff-mode isolated heart perfusion, in conjunction with 31P NMR spectroscopy, combines the fields of biochemistry and physiology into one experiment. The protocol allows for the dynamic measurement of high energy phosphate content and turnover in the heart while concurrently monitoring physiologic function. When performed correctly, this is a valuable technique in the assessment of cardiac energetics.

Bioengineered mouse models have become powerful research tools in determining causal relationships between molecular alterations and models of ...

NADH Fluorescence Imaging of Isolated Biventricular Working Rabbit Hearts

Since its inception by Langendorff1, the isolated perfused heart remains a prominent tool for studying cardiac physiology2. However, it is not well-suited for studies of cardiac metabolism, which require the heart to perform work within the context of physiologic preload and afterload pressures. Neely introduced modifications to the Langendorff technique to establish appropriate left ventricular (LV) preload and afterload pressures3. The model is known as the isolated LV working heart model and has been used extensively to study LV performance and metabolism4-6. This model, however, does not provide a properly loaded right ventricle (RV). Demmy et al. first reported a biventricular model as a modification of the LV working heart model7, 8. They found that stroke volume, cardiac output, and pressure development improved in hearts converted from working LV mode to biventricular working mode8. A properly loaded RV also diminishes abnormal pressure gradients across the septum to improve septal function. Biventricular working hearts have been shown to maintain aortic output, pulmonary flow, mean aortic pressure, heart rate, and myocardial ATP levels for up to 3 hours8.
When studying the metabolic effects of myocardial injury, such as ischemia, it is often necessary to identify the location of the affected tissue. This can be done by imaging the fluorescence of NADH (the reduced form of nicotinamide adenine dinucleotide)9-11, a coenzyme found in large quantities in the mitochondria. NADH fluorescence (fNADH) displays a near linearly inverse relationship with local oxygen concentration12 and provides a measure of mitochondrial redox state13. fNADH imaging during hypoxic and ischemic conditions has been used as a dye-free method to identify hypoxic regions14, 15 and to monitor the progression of hypoxic conditions over time10.
The objective of the method is to monitor the mitochondrial redox state of biventricular working hearts during protocols that alter the rate of myocyte metabolism or induce hypoxia or create a combination of the two. Hearts from New Zealand white rabbits were connected to a biventricular working heart system (Hugo Sachs Elektronik) and perfused with modified Krebs-Henseleit solution16 at 37 &deg;C. Aortic, LV, pulmonary artery, and left &amp; right atrial pressures were recorded. Electrical activity was measured using a monophasic action potential electrode. To image fNADH, light from a mercury lamp was filtered (350&plusmn;25 nm) and used to illuminate the epicardium. Emitted light was filtered (460&plusmn;20 nm) and imaged using a CCD camera. Changes in the epicardial fNADH of biventricular working hearts during different pacing rates are presented. The combination of the heart model and fNADH imaging provides a new and valuable experimental tool for studying acute cardiac pathologies within the context of realistic physiological conditions.

The objective is to monitor the mitochondrial redox state of isolated hearts within the context of physiologic preload and afterload pressures. A biventricular working rabbit heart model is presented. High spatiotemporal resolution fluorescence imaging of NADH is used to monitor the mitochondrial redox state of epicardial tissue.

Since its inception by Langendorff1, the isolated perfused heart remains a prominent tool for studying cardiac physiology2. However, it is not well-suited ...

Reduction in Left Ventricular Wall Stress and Improvement in Function in Failing Hearts using Algisyl-LVR

Injection of Algisyl-LVR, a treatment under clinical development, is intended to treat patients with dilated cardiomyopathy. This treatment was recently used for the first time in patients who had symptomatic heart failure. In all patients, cardiac function of the left ventricle (LV) improved significantly, as manifested by consistent reduction of the LV volume and wall stress. Here we describe this novel treatment procedure and the methods used to quantify its effects on LV wall stress and function.
Algisyl-LVR is a biopolymer gel consisting of Na+-Alginate and Ca2+-Alginate. The treatment procedure was carried out by mixing these two components and then combining them into one syringe for intramyocardial injections. This mixture was injected at 10 to 19 locations mid-way between the base and apex of the LV free wall in patients.
Magnetic resonance imaging (MRI), together with mathematical modeling, was used to quantify the effects of this treatment in patients before treatment and at various time points during recovery. The epicardial and endocardial surfaces were first digitized from the MR images to reconstruct the LV geometry at end-systole and at end-diastole. Left ventricular cavity volumes were then measured from these reconstructed surfaces.
Mathematical models of the LV were created from these MRI-reconstructed surfaces to calculate regional myofiber stress. Each LV model was constructed so that 1) it deforms according to a previously validated stress-strain relationship of the myocardium, and 2) the predicted LV cavity volume from these models matches the corresponding MRI-measured volume at end-diastole and end-systole. Diastolic filling was simulated by loading the LV endocardial surface with a prescribed end-diastolic pressure. Systolic contraction was simulated by concurrently loading the endocardial surface with a prescribed end-systolic pressure and adding active contraction in the myofiber direction. Regional myofiber stress at end-diastole and end-systole was computed from the deformed LV based on the stress-strain relationship.

This article describes procedures for implanting a novel hydrogel in failing hearts and quantifying its effect on left ventricular wall stress and function. These procedures have been successfully applied in dogs and humans.

Injection of  Algisyl-LVR, a treatment under clinical development, is intended to treat  patients with dilated cardiomyopathy. This treatment was recently ...

Intramyocardial Cell Delivery: Observations in Murine Hearts

Previous studies showed that cell delivery promotes cardiac function amelioration by release of cytokines and factors that increase cardiac tissue revascularization and cell survival. In addition, further observations revealed that specific stem cells, such as cardiac stem cells, mesenchymal stem cells and cardiospheres have the ability to integrate within the surrounding myocardium by differentiating into cardiomyocytes, smooth muscle cells and endothelial cells.
Here, we present the materials and methods to reliably deliver noncontractile cells into the left ventricular wall of immunodepleted mice. The salient steps of this microsurgical procedure involve anesthesia and analgesia injection, intratracheal intubation, incision to open the chest and expose the heart and delivery of cells by a sterile 30-gauge needle and a precision microliter syringe.
Tissue processing consisting of heart harvesting, embedding, sectioning and histological staining showed that intramyocardial cell injection produced a small damage in the epicardial area, as well as in the ventricular wall. Noncontractile cells were retained into the myocardial wall of immunocompromised mice and were surrounded by a layer of fibrotic tissue, likely to protect from cardiac pressure and mechanical load.

Intramyocardial cell delivery in murine models of cardiovascular diseases, such as hypertension or myocardial infarction, is widely used to test the therapeutic potential of different cell types in regenerative studies. Therefore, a detailed description and a clear visualization of this surgical procedure will help to define the limits and advantages of cardiovascular cell therapeutic analyses in small rodents.

Previous studies showed that cell delivery promotes cardiac function amelioration by release of cytokines and factors that increase cardiac tissue ...

Induction and Assessment of Ischemia-reperfusion Injury in Langendorff-perfused Rat Hearts

The biochemical events surrounding ischemia reperfusion injury in the acute setting are of great importance to furthering novel treatment options for myocardial infarction and cardiac complications of thoracic surgery. The ability of certain drugs to precondition the myocardium against ischemia reperfusion injury has led to multiple clinical trials, with little success. The isolated heart model allows acute observation of the functional effects of ischemia reperfusion injury in real time, including the effects of various pharmacological interventions administered at any time-point before or within the ischemia-reperfusion injury window. Since brief periods of ischemia can precondition the heart against ischemic injury, in situ aortic cannulation is performed to allow for functional assessment of non-preconditioned myocardium. A saline filled balloon is placed into the left ventricle to allow for real-time measurement of pressure generation. Ischemic injury is simulated by the cessation of perfusion buffer flow, followed by reperfusion. The duration of both ischemia and reperfusion can be modulated to examine biochemical events at any given time-point. Although the Langendorff isolated heart model does not allow for the consideration of systemic events affecting ischemia and reperfusion, it is an excellent model for the examination of acute functional and biochemical events within the window of ischemia reperfusion injury as well as the effect of pharmacological intervention on cardiac pre- and postconditioning. The goal of this protocol is to demonstrate how to perform in situ aortic cannulation and heart excision followed by ischemia/reperfusion injury in the Langendorff model.

The isolated rat heart is an enduring model for ischemia reperfusion injury. Here, we describe the process of harvesting the beating heart from a rat via in situ aortic cannulation, Langendorff perfusion of the heart, simulated ischemia-reperfusion injury, and infarct staining to confirm the extent of ischemic insult.

The biochemical events surrounding ischemia reperfusion injury in the acute setting are of great importance to furthering novel treatment options for ...

Primary Outcome Assessment in a Pig Model of Acute Myocardial Infarction

Mortality after acute myocardial infarction remains substantial and is associated with significant morbidity, like heart failure. Novel therapeutics are therefore required to confine cardiac damage, promote survival and reduce the disease burden of heart failure. Large animal experiments are an essential part in the translational process from experimental to clinical therapies. To optimize clinical translation, robust and representative outcome measures are mandatory. The present manuscript aims to address this need by describing the assessment of three clinically relevant outcome modalities in a pig acute myocardial infarction (AMI) model: infarct size in relation to area at risk (IS/AAR) staining, 3-dimensional transesophageal echocardiography (TEE) and admittance-based pressure-volume (PV) loops. Infarct size is the main determinant driving the transition from AMI to heart failure and can be quantified by IS/AAR staining. Echocardiography is a reliable and robust tool in the assessment of global and regional cardiac function in clinical cardiology. Here, a method for three-dimensional transesophageal echocardiography (3D-TEE) in pigs is provided. Extensive insight into cardiac performance can be obtained by admittance-based pressure-volume (PV) loops, including intrinsic parameters of myocardial function that are pre- and afterload independent. Combined with a clinically feasible experimental study protocol, these outcome measures provide researchers with essential information to determine whether novel therapeutic strategies could yield promising targets for future testing in clinical studies.

Reliable and accurate outcome assessment is the key for translation of preclinical therapies into clinical treatment. The current paper describes how to assess three clinically relevant primary outcome parameters of cardiac performance and damage in a pig acute myocardial infarction model.

Mortality after acute myocardial infarction remains substantial and is associated with significant morbidity, like heart failure. Novel therapeutics are ...

A Detailed Protocol for Physiological Parameters Acquisition and Analysis in Neurosurgical Critical Patients

Intracranial pressure (ICP) monitoring is now widely used in neurosurgical critical patients. Besides mean ICP value, the ICP derived parameters such as ICP waveform, amplitude of pulse (AMP), the correlation of ICP amplitude and ICP mean (RAP), pressure reactivity index (PRx), ICP and arterial blood pressure (ABP) wave amplitude correlation (IAAC), and so on, can reflect intracranial status, predict prognosis, and can also be used as guidance of proper treatment. However, most of the clinicians focus only on the mean ICP value while ignoring these parameters because of the limitations of the current devices. We have recently developed a multimodality monitoring system to address these drawbacks. This portable, user-friendly system will use a data collecting and storing device to continuously acquire patients&#39; physiological parameters first, i.e., ABP, ICP, and oxygen saturation, and then analyze these physiological parameters. We hope that the multimodality monitoring system will be accepted as a key measure to monitor physiological parameters, to analyze the current clinical status, and to predict the prognosis of the neurosurgical critical patients.

Recently, we have developed a portable multimodality monitoring system for the monitoring of various physiological parameters in neurosurgical critical patients. Detailed protocols on how to use this multimodality monitoring system are presented here.

Intracranial pressure (ICP) monitoring is now widely used in neurosurgical critical patients. Besides mean ICP value, the ICP derived parameters such as ...

Cochlear Implantation in the Guinea Pig

Cochlear implants are highly efficient devices that can restore hearing in subjects with profound hearing loss. Due to improved speech perception outcomes, candidacy criteria have been expanded over the last few decades. This includes patients with substantial residual hearing that benefit from electrical and acoustical stimulation of the same ear, which makes hearing preservation during cochlear implantation an important issue. Electrode impedances and the related issue of energy consumption is another major research field, as progress in this area could pave the way for fully implantable auditory prostheses. To address these issues in a systematic way, adequate animal models are essential. Therefore, the goal of this protocol is to provide an animal model of cochlear implantation, which can be used to address various research questions. Due to its large tympanic bulla, which allows easy surgical access to the inner ear, as well as its hearing range which is relatively similar to the hearing range of humans, the guinea pig is a commonly used species in auditory research. Cochlear implantation in the guinea pig is performed via a retroauricular approach. Through the bullostomy a cochleostomy is drilled and the cochlear implant electrode is inserted into the scala tympani. This electrode can then be used for electrical stimulation, determination of electrode impedances and the measurement of compound action potentials of the auditory nerve. In addition to these applications, cochlear implant electrodes can also be used as drug delivery devices, if a topical delivery of pharmaceutical agents to the cells or fluids of the inner ear is intended.

The goal of this protocol is to provide an animal model of cochlear implantation, which can be used to address a multitude of research questions. Potential applications include the evaluation of pharmaceutical interventions or electrical stimulation for beneficial effects on hearing thresholds or electrode impedances.

Cochlear implants are highly efficient devices that can restore hearing in subjects with profound hearing loss. Due to improved speech perception ...

A Minimally Invasive Model to Analyze Endochondral Fracture Healing in Mice Under Standardized Biomechanical Conditions

Bone healing models are necessary to analyze the complex mechanisms of fracture healing to improve clinical fracture treatment. During the last decade, an increased use of mouse models in orthopedic research was noted, most probably because mouse models offer a large number of genetically-modified strains and special antibodies for the analysis of molecular mechanisms of fracture healing. To control the biomechanical conditions, well-characterized osteosynthesis techniques are mandatory, also in mice. Here, we report on the design and use of a closed bone healing model to stabilize femur fractures in mice. The intramedullary screw, made of medical-grade stainless steel, provides through fracture compression an axial and rotational stability compared to the mostly used simple intramedullary pins, which show a complete lack of axial and rotational stability. The stability achieved by the intramedullary screw allows the analysis of endochondral healing. A large amount of callus tissue, received after stabilization with the screw, offers ideal conditions to harvest tissue for biochemical and molecular analyses. A further advantage of the use of the screw is the fact that the screw can be inserted into the femur with a minimally invasive technique without inducing damage to the soft tissue. In conclusion, the screw is a unique implant that can ideally be used in closed fracture healing models offering standardized biomechanical conditions.

This protocol describes a minimally invasive osteosynthesis technique using an intramedullary screw for standardized stabilization of femur fractures, which can be used to analyze endochondral bone healing in mice.

Bone healing models are necessary to analyze the complex mechanisms of fracture healing to improve clinical fracture treatment. During the last decade, an ...

Functional and Physiological Methods of Evaluating Median Nerve Regeneration in the Rat

The main goal of this investigation is to show how to create and repair different types of median nerve (MN) lesions in the rat. Moreover, different methods of simulating postoperative physiotherapy are presented. Multiple standardized strategies are used to assess motor and sensory recovery using an MN model of peripheral nerve lesion and repair, thus permitting easy comparison of the results. Several options are included for providing a postoperative physiotherapy-like environment to rats that have undergone MN injuries. Finally, the paper provides a method to evaluate the recovery of the MN using several noninvasive tests (i.e., grasping test, pin prick test, ladder rung walking test, rope climbing test, and walking track analysis), and physiological measurements (infrared thermography, electroneuromyography, flexion strength evaluation, and flexor carpi radialis muscle weight determination). Hence, this model seems particularly appropriate to replicate a clinical scenario, facilitating extrapolation of results to the human species.
Although the sciatic nerve is the most studied nerve in peripheral nerve research, analysis of the rat MN presents various advantages. For example, there is a reduced incidence of joint contractures and automutilation of the affected limb in MN lesion studies. Furthermore, the MN is not covered by muscle masses, making its dissection easier than that of the sciatic nerve. In addition, MN recovery is observed sooner, because the MN is shorter than the sciatic nerve. Also, the MN has a parallel path to the ulnar nerve in the arm. Hence, the ulnar nerve can be easily used as the nerve graft for repairing MN injuries. Finally, the MN in rats is located in the forelimb, akin to the human upper limb; in humans, the upper limb is the site of most peripheral nerve lesions.

Presented is a protocol to produce different types of median nerve (MN) lesions and repair in the rat. Additionally, the protocol shows how to evaluate the functional recovery of the nerve using several noninvasive behavioral tests and physiological measurements.

The main goal of this investigation is to show how to create and repair different types of median nerve (MN) lesions in the rat. Moreover, different ...