Research was funded by DZHK grants 81Z0600207 (JH, PS, and DM) and 81X2600253 (AD and TS).
The authors would like to thank Claudia Fahney, Mei-Ping Wu, and Matthias Semisch for their support in preparing the set-ups, as well as for the regular maintenance of the tissue cultivation.

Tissue collection for the experiments described here was approved by the Institutional Review Boards of the University of Munich and the Ruhr-University Bochum. Studies were conducted according to Declaration of Helsinki guidelines. Patients gave their written informed consent prior to tissue collection.
1. Tissue acquisition
<ol>
	<li>Obtain human tissue from patients undergoing heart transplantation or cardiac surgery.</li>
	<li>Before procuring the tissue, prepare 2 L of cardioplegic solution (further referred to as slicing buffer (Table 1)).</li>
	<li>After obtaining a 4 cm x 4 cm sized transmural left ventricular (LV) biopsy, place the tissue immediately (within 5 min after excision) in a closable plastic single-use sterile beaker containing approximately 70 mL of cold (4 &#176;C) slicing buffer. Keep at 4 &#176;C. 
	&#8203;NOTE: The slicing buffer used in this protocol allows for cold storage (4 &#176;C) of the tissue for up to 36 h. This permits cold transport of the tissue from clinics which are not close to the laboratory. However, transport times &#8804; 24 h have proven to be optimal.</li>
</ol>
2. Preparing agarose and the vibratome
<ol>
	<li>Make sure sufficient cultivation chambers are prepared and sterilized (Figure 1A).
	<ol>
		<li>Submerge the chambers and graphite electrodes in 1 L of a 10% isopropanol solution and agitate overnight. The following day, transfer the chambers to a 100% isopropanol solution for 3 min and autoclave the graphite electrodes at 120 &#176;C for 10 min.</li>
		<li>Let the chambers and graphite electrodes air-dry under a laminar flow hood.</li>
		<li>Attach a circuit board to each of the chambers according to the available positions on the rocker. Place two graphite electrodes to the circuit board as per manufacturer&#39;s instructions.</li>
		<li>Place a 35 mm Petri dish lid on top of the chamber to prevent contamination.</li>
	</ol>
	</li>
	<li>Set up the Myodish cultivation system (see Table of Materials) in an incubator at 37 &#176;C and 5% CO2 by connecting the system to a computer (Figure 1B).</li>
	<li>Prepare the 4% low-melt agarose in slicing buffer (without glucose; Table 2). The agarose can be stored at 4 &#176;C for 6 months.
	<ol>
		<li>On the day of the experiment, melt a stock volume of agarose solution using a water bath set at 80 &#176;C. Depending on the quantity, melting takes approximately 30 min.</li>
		<li>When the agarose is liquid, draw up 8 mL of liquid agarose into a 10 mL syringe, with an additional 2 mL of air. Close the syringe with a sterile cap and place it upside down in a 37 &#176;C water bath for 20 min, to prevent the agarose from solidifying, and to equilibrate its temperature to prevent hyperthermia damage of the sample.</li>
	</ol>
	</li>
	<li>If present, turn on water cooling device of the vibratome at least 30 min prior to tissue slicing to allow for sufficient cooling capacity. Set the temperature of the water circulation to 4 &#176;C. 
	NOTE: The cutting tray and cooling plate used here both contained a built-in water circulation. This is highly recommended for cooling and to lower the contamination risks. It is however also possible to use ice as a cooling technique. Also, the water-cooling device does not need to be connected to the vibratome&#39;s cutting tray and/or cooling plate at this point in the protocol.</li>
	<li>Clean the vibratome&#39;s slicing tray and sample plate by flushing all surfaces with 100% isopropanol for at least 3 min. 
	NOTE: Depending on the vibratome system used, the vibratome set-up might differ. Refer to the manual of the vibratome present in the laboratory for information about the set-up and blade calibration methods.</li>
	<li>Fill the cutting tray up to 90%-95% with slicing buffer. In the currently used set-up, this corresponds to approximately 400 mL. Connect the tubing of the water-cooling device to the valves on the vibratome&#39;s cutting tray and cooling plate.</li>
	<li>Disinfect all tools needed for the preparation by submerging them in a 100% isopropanol solution for 5 s. Remove the tools from the isopropanol solution and air-dry under the laminar flow hood.</li>
</ol>
3. Trimming and embedding the samples
<ol>
	<li>Transfer the tissue sample to a 100 mm Petri dish filled with cold slicing buffer. Keep the dish on a cooling plate at 4 &#176;C (connected to cooler or placed on ice).</li>
	<li>Remove endocardial trabeculae by holding the endocardium with tweezers and using scissors to cut away approximately 3 mm of endocardial tissue. In the same way, remove excessive adipose tissue underneath the epicardium if present.</li>
	<li>Fixate the cut tissue sample, endocardial side up, to a 2 cm x 2 cm rubber patch using four 0.9 mm x 70 mm 20 G needles that are fixed in a square position (0.9 cm x 0.9 cm; Figure 1C, D). Make sure that the diagonal edge of each needle tip is pointing inward. This enhances fixation and prevents damage to the myocardium. 
	CAUTION: The orientation of the above-mentioned square should be orthogonal to the expected predominant myofiber direction.</li>
	<li>Cut away all the excess tissue outside of the four needle square using a scalpel. If the size of the original sample allows, use two myocardial tissues samples during this preparation from the same raw sample.</li>
	<li>Using tweezers, place the trimmed sample on a sterile piece of tissue to remove any excess slicing buffer left on the sample.To prevent drying out of the sample, do not keep the sample on the tissue for longer than 10 s.</li>
	<li>Place the sample(s) in a 35 mm Petri dish, such that the blade cuts perpendicular to the cardiomyocyte alignment and the epicardium is facing downwards. If the preparation includes two samples, make sure the samples are centered and do not touch each other.</li>
	<li>Take the agarose syringe from the water bath and submerge the sample(s) in agarose. (Figure 2A). Let the agarose solidify for 5 min on a cooling plate. The sample(s) must stay in contact with the Petri dish, which will ensure that the cutting plane will be parallel to the predominant myocardial fiber direction. 
	&#8203;CAUTION: Do not fully empty the syringe, in order to prevent air bubbles. Pay attention to the amount of agarose that is left in the syringe during submerging of the samples. In the case of air bubbles in the dish with the samples, carefully pull air bubbles back into the syringe.</li>
</ol>
4. Placing the samples on cutting tray
<ol>
	<li>Remove the solidified agarose containing the sample(s) from the 35 mm Petri dish using a spatula or similar tool, by wedging it in-between the sample and the side of the Petri dish. Cut away some of the agarose using a scalpel while keeping the samples covered. 
	CAUTION: Do not remove too much of the agarose. There should be at least 5 mm of agarose left on the long and short sides in the X/Z plane. 
	NOTE: Steps 4.2-4.4 need to be performed in rapid succession (i.e., maximum of 5 s). Have all required tools within reach prior to starting. No repositioning of the sample is possible after placement. Try to limit the exposure of the glue to air and moisture. Contact with air or fluids will solidify the glue, making it unusable.</li>
	<li>With a pipette, place and distribute 60 &#181;L of glue in and around the center of the cutting platform.</li>
	<li>Place the epicardial side of the sample contained in agarose on top of the glued area using tweezers. Do not reposition. The endocardial side of the sample must be visible in the agarose. Let the glue solidify for 1 min. Gently press the agarose containing the sample from the top with a blunt tool (e.g., tweezers) while preventing cutting into or damaging the agarose.</li>
	<li>Place the sample platform in its designated position in the cutting tray of the vibratome, filled with slicing buffer.</li>
</ol>
5. Starting the vibratome
<ol>
	<li>Set vibration amplitude to 1 mm and the initial cutting speed to 0.07 mm/s. Set the thickness of the slice to 300 &#181;m to cut slices.</li>
	<li>As long as the blade only cuts the agarose, increase the cutting speed to its maximum, which in this case is 1.50 mm/s. As soon as the vibratome starts cutting the tissue, decrease the speed to 0.07 mm/s immediately. 
	&#8203;NOTE: If the tissue is not sliced smoothly, for example if there are large fibrotic areas in the sample, it may help to increase the cutting amplitude up to 1.5 mm and to reduce the cutting speed to 0.04 mm/s.</li>
</ol>
6. Medium and incubator preparation during slicing procedure
<ol>
	<li>Fill each cultivation chamber with 2.4 mL of complete cultivation medium (Table 3).</li>
	<li>Place the cultivation chambers filled with medium on the cultivation system in the incubator set at 37&#160;&#176;C, 5% CO2, 21% O2, and a humidity of 80%. Equilibrate the medium for at least 20 min.</li>
	<li>Connect the cultivation system with a computer and start the corresponding software program.</li>
	<li>Set the rocker speed to 60 rpm and preset the stimulation parameters (stimulation pulses and frequency). For human cardiac slices, set the standard stimulation to biphasic impulses with 50 mA current, each consisting of 3 ms positive current, a 1 ms pause and a 3 ms pulse of inverted current, at a pacing rate of 30 beats per min (BPM). 
	NOTE: In well preserved tissues, the typical stimulation threshold is around 15 mA. To ensure reliable stimulation and to account for any possible increase of the stimulation threshold, it is recommended to set current to a value that exceeds the stimulation threshold by two- to three-fold.</li>
	<li>Check the electrode indicators of the software to verify that the electrodes of the cultivation chambers are working correctly. 
	&#8203;NOTE: Action is needed whenever the channel indicator in the cultivation software turns red. In this case, the bipolar pulse charges are not balanced.</li>
</ol>
7. Preparing the slices
NOTE: Initial subendocardial slices are commonly not suitable for tissue cultivation and need to be discarded because of uneven morphology. After the first five to 10 slices, slice texture and morphology improve. The ideal slice is at least 1 cm x 1 cm, has no or only limited fibrotic patches, is not fragmented, and has homogeneous fiber alignment (Figure 2B, D). Interstitial fibrosis, located between the myocyte fibers, is often present in failing human myocardium. Surprisingly, this is not a negative predictor of cultivation success.
<ol>
	<li>Pour an adequate amount of cold slicing buffer in a 5 cm Petri dish lid to ensure slices will not dry out. Place the slices in the Petri dish lid containing the cold slicing buffer.</li>
	<li>Separate the agarose from the tissue by using tweezers. Prevent touching the tissue. Handle the tissue carefully as any damage to the tissue will reduce the success rate of the cultivation.</li>
	<li>Determine the direction of the myocardial fibers by close inspection against a light source. This is of importance when attaching the plastic triangles to the tissue in step 7.6. 
	NOTE: Steps 7.4 and 7.5 need to be performed in quick succession, within 5 s.</li>
	<li>Attach two plastic triangles to a sample using glue in order to anchor the tissue inside the cultivation chambers.
	<ol>
		<li>Place 1 &#181;L of glue on a sterile Petri dish lid. Use a hooked tweezer to pick up one of the autoclaved plastic triangles. Quickly dip the front edge of the triangle into the glue and paste the triangle onto the sample, perpendicular to the cardiomyocyte alignment. Repeat for the other triangle.</li>
	</ol>
	</li>
	<li>Trim off tissue exceeding the triangle width with a scalpel (Figure 2C). Place the slice with the two mounted triangles back into the cutting tray&#39;s slicing buffer. 
	NOTE: Repeat steps 7.2 to 7.5 until enough slices are prepared to fill the cultivation chambers. We recommend preparing a few additional slices to allow for the replacement of slices with poor contraction.</li>
</ol>
8. Mounting the slices
NOTE: The afterload is determined by the stiffness of the spring wire in the cultivation chambers. Three different types are available, based on the thickness of the spring wire.
<ol>
	<li>Take a medium-filled cultivation chamber from the incubator. Select one of the prepared slices and insert it into the chamber by connecting one triangle to each pin.</li>
	<li>Adjust the distance between the mounting pins according to the sample size. Make sure the sample is submerged in medium. Place the chamber back into its designated socket of the cultivation system in the incubator. 
	CAUTION: Do not overstretch the tissue and do not over-bend the spring wire!</li>
	<li>Set the preload tension after placement of the dish onto the rocker.
	<ol>
		<li>Decrease the preload by turning the adjusting screw counterclockwise. Do this until the baseline of the corresponding graph on the computer screen does not change anymore.</li>
		<li>Carefully increase the preload (i.e., increase the tension) by turning the adjusting screw clockwise. For the chambers with the highest stiffness, continue until the corresponding baseline in the graph has increased by 1000-1200 units, which corresponds to 1 mN of preload. 
		&#8203;NOTE: The exact adjustment will depend on the individual spring constant of a cultivation chamber, which can be determined by calibration prior to an experiment according to the manufacturer&#39;s instructions. The recording software permits consideration of the individual chamber calibration so that forces will uniformly be displayed in &#181;N. It is recommended to apply electrical stimulation from the start of the cultivation. As such, it is well possible that the slice starts contracting during the preload adjustment. In this case, focus on the diastolic baseline to assess the preload.</li>
	</ol>
	</li>
</ol>
9. Changing the medium
<ol>
	<li>Prepare cultivation medium according to the recipe in Table 3. Shortly before use of the medium, add 50 &#181;L of &#946;-mercaptoethanol (50 mM) to a 50 mL tube. Store at 4 &#176;C.</li>
	<li>Once every 2 days, partly exchange the cultivation medium. Refresh the medium every 2 days, if long term cultivation of the myocardial slices is desired.</li>
	<li>Pre-warm the fresh medium in a water bath or hot-air incubator at 37 &#176;C for 30-45 min. Remove a cultivation chamber from the incubator and place it under a laminar flow hood. 
	CAUTION: It is essential to keep the chamber, as well as the medium, warm at 37 &#176;C (&#62; 35 &#176;C) to prevent hyper contractures due to low temperature. Less distinct damage may be present as an increase of diastolic tension within a few hours after medium exchange. This may seem to recover, but repeated stress might result in accumulating deterioration.</li>
	<li>Remove medium from the cultivation chamber, leaving approximately 0.8 mL in the chamber. Add 1.6 mL of fresh medium to the same chamber. The total volume of medium should be around 2.4 mL per chamber.</li>
	<li>Place the cover of the chamber back and place the cultivation chamber back into its respective position.</li>
</ol>

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JH, PS, DM, and KL have nothing to disclose. AD and TS are shareholders of InVitroSys GmbH, which provides the Myodish cultivation system.

In the past, cardiovascular research has made great advances in the cultivation of cardiomyocytes. However, the 3D cultivation of cardiomyocytes with intact geometry is not yet well-established. Compared to previous protocols applied for ex vivo cultivation of myocardial tissue, the protocol that we described here resembles the in vivo environment of the tissue more closely. Moreover, the application of pre- and afterload allows for a more biomimetic environment. We are able to fully analyze and underst...

In the past decade, in vitro cultivation of myocardial cells has made great advances, ranging from 2D and three-dimensional (3D) techniques to the use of organoids and induced pluripotent stem cells differentiated into cardiac myocytes1,2,3. Ex vivo and primary cell cultivations have shown to be of great value, especially for genetic studies and drug development4,5,6. Using human tissues improves the translational value of the results. Long-term 3D cultivation of myocardial tissues with intact geometry, however, is not well-established. The intact geometry is a key feature to mimic in vivo conditions, as proper cardiac function, communication between different cells, as well as cell-matrix interactions are prerequisites. Myocardial tissue cultivation went through various phases of development. The success rate and stability of ex vivo myocardial tissue cultivation were initially quite low, but recent approaches have yielded promising results7,8,9,10,11.
Among those, Fischer et al. were the first to demonstrate that viability and contractile performance of human myocardial tissue can be maintained in ex vivo cell cultivation for many weeks7. Their technique was based on thin tissue slices cut from explanted human myocardium, which were mounted in newly developed cultivation chambers that provided defined biomechanical conditions and continuous electrical stimulation. This cultivation method closely resembles the in vivo function of myocardial tissue, and has been reproduced by several independent research groups2,12,13,14,15. Importantly, the chambers used by Fischer et al. also enabled continuous registration of developed forces for up to 4 months, and thus opened unprecedented opportunities for physiological and pharmacological research on intact human myocardium7.
Similar techniques were independently developed by other groups and applied to human, rat, porcine, and rabbit myocardium7,10,11. Pitoulis et al. subsequently developed a more physiological method, which reproduces the normal force-length relation during a contraction cycle, but is less suitable for high-throughput analysis16. As such, the general approach of biomimetic cultivation can be regarded as a further step into the reduction, refinement, and replacement (3R) of animal experiments.
However, exploitation of this potential requires standardized procedures, high content analyses, and a high throughput level. We present a technique that combines automated slicing of living human myocardium with in vitro maintenance in a biomimetic cultivation system that has become commercially available (see Table of Materials). With the proposed approach, the number of individual slices that can be generated from a single transmural myocardial specimen is only limited by the processing time. A specimen of sufficient size and quality (3 cm x 3 cm) often yields 20-40 tissue slices being conveniently cut with an automated vibratome. These slices can be placed in cultivation chambers belonging to the system. The chambers allow for electrical stimulation, the parameters of which can be modulated (i.e., pulse duration, polarity, rate, and current), as well as the adjustment of pre- and afterload, using spring wires inside the chambers. The contraction of each slice is registered from the movement of a small magnet attached to a spring wire and displayed as an interpretable graph. Data can be recorded at all times and analyzed using freely available software. Aside from the constant baseline pacing, scheduled protocols can be performed to functionally assess their refractory period, stimulation threshold, post-pause-potentiation, and force-frequency relation.
This long-term biomimetic cultivation of multiple myocardial slices from an individual heart paves the way for future ex vivo research in both human and animal tissue, and facilitates the screening for therapeutic and cardiotoxic drug effects in cardiovascular medicine. It has already been applied to various experimental approaches2,12,13,15. Here, we give a detailed step-by-step description of the preparation of human tissue and provide solutions for frequently encountered cultivation problems.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Chemicals</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Agarose Low melting point</td>
			<td>Roth</td>
			<td>6351.2</td>
			<td></td>
		</tr>
		<tr>
			<td>Bay-K8644</td>
			<td>Cayman Chemical</td>
			<td>19988</td>
			<td></td>
		</tr>
		<tr>
			<td>BDM (2,3-Butanedione monoxime)</td>
			<td>Sigma</td>
			<td>B0753-1kg</td>
			<td></td>
		</tr>
		<tr>
			<td>CaCl2*H2O</td>
			<td>Merck</td>
			<td>2382.1</td>
			<td></td>
		</tr>
		<tr>
			<td>Calciseptine</td>
			<td>Alomone Labs</td>
			<td>SPC-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Glucose*H2O</td>
			<td>AppliChem</td>
			<td>A3730.0500</td>
			<td></td>
		</tr>
		<tr>
			<td>H2O</td>
			<td>BBraun</td>
			<td>3703452</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>AppliChem</td>
			<td>A1069.0500</td>
			<td></td>
		</tr>
		<tr>
			<td>Histoacryl</td>
			<td>BBraun</td>
			<td>1050052</td>
			<td></td>
		</tr>
		<tr>
			<td>Isopropanol 100%</td>
			<td>SAV LP GmbH</td>
			<td>UN1219</td>
			<td></td>
		</tr>
		<tr>
			<td>ITS-X-supplement</td>
			<td>Gibco</td>
			<td>5150056</td>
			<td></td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Merck</td>
			<td>1.04933.0500</td>
			<td></td>
		</tr>
		<tr>
			<td>Medium 199</td>
			<td>Gibco</td>
			<td>31150-022</td>
			<td></td>
		</tr>
		<tr>
			<td>MgCl2*6H2O</td>
			<td>AppliChem</td>
			<td>A1036.0500</td>
			<td></td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma</td>
			<td>S5886-1KG</td>
			<td></td>
		</tr>
		<tr>
			<td>NaH2PO4*H2O</td>
			<td>Merck</td>
			<td>1.06346.0500</td>
			<td></td>
		</tr>
		<tr>
			<td>Nifedipine</td>
			<td>Sigma</td>
			<td>N7634-1G</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin / streptomycin x100</td>
			<td>Sigma</td>
			<td>P0781-100ML</td>
			<td></td>
		</tr>
		<tr>
			<td>&#946;-Mercaptoethanol</td>
			<td>AppliChem</td>
			<td>A1108.0100</td>
			<td></td>
		</tr>
		<tr>
			<td>Laboratory equipment</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Flow cabinet</td>
			<td>Thermo Scientific</td>
			<td>KS15</td>
			<td></td>
		</tr>
		<tr>
			<td>Frigomix waterpump and cooling + BBraun Thermomix BM</td>
			<td>BBraun</td>
			<td></td>
			<td>In-house made combination of cooling and heating solution.</td>
		</tr>
		<tr>
			<td>Incubator</td>
			<td>Binder</td>
			<td>CB240</td>
			<td></td>
		</tr>
		<tr>
			<td>MyoDish bioreactor system</td>
			<td>InVitroSys GmbH</td>
			<td>MyoDish 1</td>
			<td>Myodish cultute system</td>
		</tr>
		<tr>
			<td>Vibratome</td>
			<td>Leica</td>
			<td>VT1200s</td>
			<td></td>
		</tr>
		<tr>
			<td>Water bath 37 degrees</td>
			<td>Haake</td>
			<td>SWB25</td>
			<td></td>
		</tr>
		<tr>
			<td>Water bath 80 degrees</td>
			<td>Daglef Patz KG</td>
			<td>7070</td>
			<td></td>
		</tr>
		<tr>
			<td>Materials</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>100 mL plastic single-use beaker</td>
			<td>Sarstedt</td>
			<td>75.562.105</td>
			<td></td>
		</tr>
		<tr>
			<td>Filtration unit, Steritop Quick Release</td>
			<td>Millipore</td>
			<td>S2GPT05RE</td>
			<td></td>
		</tr>
		<tr>
			<td>Needles 0.9 x 70 mm 20G</td>
			<td>BBraun</td>
			<td>4665791</td>
			<td></td>
		</tr>
		<tr>
			<td>Plastic triangles</td>
			<td></td>
			<td></td>
			<td>In-house made</td>
		</tr>
		<tr>
			<td>Razor Derby premium</td>
			<td>Derby Tokai</td>
			<td>B072HJCFK6</td>
			<td></td>
		</tr>
		<tr>
			<td>Razor Gillette Silver Blue</td>
			<td>Gillette</td>
			<td>7393560010170</td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel disposable</td>
			<td>Feather</td>
			<td>02.001.30.020</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe 10 mL Luer tip BD Discardit</td>
			<td>BBraun</td>
			<td>309110</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue Culture Dish 10 cm</td>
			<td>Falcon</td>
			<td>353003</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue Culture Dish 3.5 cm</td>
			<td>Falcon</td>
			<td>353001</td>
			<td></td>
		</tr>
		<tr>
			<td>Tubes 50 mL</td>
			<td>Falcon</td>
			<td>352070</td>
			<td></td>
		</tr>
	</tbody>
</table>

preparation of human myocardial tissue for long-term cultivation

Cardiomyocyte cultivation has seen a vast number of developments, ranging from two-dimensional (2D) cell cultivation to iPSC derived organoids. In 2019, an ex vivo way to cultivate myocardial slices obtained from human heart samples was demonstrated, while approaching in vivo condition of myocardial contraction. These samples originate mostly from heart transplantations or left-ventricular assist device placements. Using a vibratome and a specially developed cultivation system, 300 &#181;m thick slices are placed between a fixed and a spring wire, allowing for stable and reproducible cultivation for several weeks. During cultivation, the slices are continuously stimulated according to individual settings. Contractions can be displayed and recorded in real-time, and pharmacological agents can be readily applied. User-defined stimulation protocols can be scheduled and performed to assess vital contraction parameters like post-pause-potentiation, stimulation threshold, force-frequency relation, and refractory period. Furthermore, the system enables a variable pre- and afterload setting for a more physiological cultivation.
Here, we present a step-by-step guide on how to generate a successful long-term cultivation of human left ventricular myocardial slices, using a commercial biomimetic cultivation solution.

The contraction of the myocardial slices was displayed on the computer screen after insertion of the cultivation chamber into its corresponding connector (Figure 3). Contraction of the human myocardial slices started immediately upon stimulation. The slices hypercontracted for 5-10 min. This was visible as an increase of diastolic forces, caused by a tonic contracture of damaged tissue fractions. This process was reverted to varying degrees within 1-1.5 h. After stabilizing, human LV tissue ...

Watch this Scientific Journal Video about Preparation of Human Myocardial Tissue for Long-Term Cultivation at JoVE.com

Preparation of Human Myocardial Tissue for Long-Term Cultivation

We present a protocol for ex vivo cultivation of human ventricular myocardial tissue. It allows for detailed analysis of contraction force and kinetics, as well as the application of pre- and afterload to mimic the in vivo physiological environment more closely.

Cardiomyocyte cultivation has seen a vast number of developments, ranging from two-dimensional (2D) cell cultivation to iPSC derived organoids. In 2019, ...

preparation-of-human-myocardial-tissue-for-long-term-cultivation

Research

JoVE Journal

Medicine

4.2K Views.  University Hospital, LMU Munich. We present a protocol for ex vivo cultivation of human ventricular myocardial tissue. It allows for detailed analysis of contraction force and kinetics, as well as the application of pre- and afterload to mimic the in vivo physiological environment more closely. 

Video: Preparation of Human Myocardial Tissue for Long-Term Cultivation

Corneal Donor Tissue Preparation for Endothelial Keratoplasty

Over the past ten years, corneal transplantation surgical techniques have undergone revolutionary changes1,2. Since its inception, traditional full thickness corneal transplantation has been the treatment to restore sight in those limited by corneal disease. Some disadvantages to this approach include a high degree of post-operative astigmatism, lack of predictable refractive outcome, and disturbance to the ocular surface. The development of Descemet's stripping endothelial keratoplasty (DSEK), transplanting only the posterior corneal stroma, Descemet's membrane, and endothelium, has dramatically changed treatment of corneal endothelial disease. DSEK is performed through a smaller incision; this technique avoids 'open sky' surgery with its risk of hemorrhage or expulsion, decreases the incidence of postoperative wound dehiscence, reduces unpredictable refractive outcomes, and may decrease the rate of transplant rejection3-6.
Initially, cornea donor posterior lamellar dissection for DSEK was performed manually1 resulting in variable graft thickness and damage to the delicate corneal endothelial tissue during tissue processing. Automated lamellar dissection (Descemet's stripping automated endothelial keratoplasty, DSAEK) was developed to address these issues. Automated dissection utilizes the same technology as LASIK corneal flap creation with a mechanical microkeratome blade that helps to create uniform and thin tissue grafts for DSAEK surgery with minimal corneal endothelial cell loss in tissue processing.
Eye banks have been providing full thickness corneas for surgical transplantation for many years. In 2006, eye banks began to develop methodologies for supplying precut corneal tissue for endothelial keratoplasty. With the input of corneal surgeons, eye banks have developed thorough protocols to safely and effectively prepare posterior lamellar tissue for DSAEK surgery. This can be performed preoperatively at the eye bank. Research shows no significant difference in terms of the quality of the tissue7 or patient outcomes8,9 using eye bank precut tissue versus surgeon-prepared tissue for DSAEK surgery. For most corneal surgeons, the availability of precut DSAEK corneal tissue saves time and money10, and reduces the stress of performing the donor corneal dissection in the operating room. In part because of the ability of the eye banks to provide high quality posterior lamellar corneal in a timely manner, DSAEK has become the standard of care for surgical management of corneal endothelial disease.
The procedure that we are describing is the preparation of the posterior lamellar cornea at the eye bank for transplantation in DSAEK surgery (Figure 1).

Endothelial corneal transplantation is a surgical technique for treatment of posterior corneal diseases. Mechanical microkeratome dissection to prepare tissue results in thinner, more symmetric grafts with less endothelial cell loss and improved outcomes. Dissections can be performed at the eye bank prior to corneal transplantation surgery.

Over the past ten years, corneal transplantation surgical techniques have undergone revolutionary changes1,2. Since its inception, traditional full ...

Working with Human Tissues for Translational Cancer Research

Medical research for human benefit is greatly impeded by the necessity for human tissues and subjects. However, upon obtaining consent for human specimens, precious samples must be handled with the greatest care in order to ensure integrity of organs, tissues, and cells to the highest degree. Unfortunately, tissue processing by definition requires extraction of tissues from the host, a change which can cause great cellular stress and have major repercussions on subsequent analyses. These stresses could result in the specimen being no longer representative of the site from which it was retrieved. Therefore, a strict protocol must be adhered to while processing these specimens to ensure representativeness. The desired assay(s) must also be taken into consideration in order to ensure that an optimal technique is used for sample processing. Outlined here is a protocol for tissue retrieval, processing and various analyses which may be performed on processed tissue in order to maximize downstream production from limited tissue samples.

Translational cancer research is dependent on extraction of human tissues. Much work has gone into optimizing extraction methods for ex vivo analysis. Here, we describe tissue processing methods allowing for maximal data output from limited samples.

Medical research for human benefit is greatly impeded by the necessity for human tissues and subjects. However, upon obtaining consent for human ...

Long-Term Catheterization of the Intestinal Lymph Trunk and Collection of Lymph in Neonatal Pigs

Catheterization of the intestinal lymph trunk in neonatal pigs is a technique allowing for the long-term collection of large quantities of intestinal (central) efferent lymph. Importantly, the collection of central lymph from the intestine enables researchers to study both the mechanisms and lipid constitutes associated with lipid metabolism, intestinal inflammation and cancer metastasis, as well as cells involved in immune function and immunosurveillance. A ventral mid-line surgical approach permits excellent surgical exposure to the cranial abdomen and relatively easy access to the intestinal lymph trunk vessel that lies near the pancreas and the right ventral segment of the portal vein underneath the visceral aspect of the right liver lobe. The vessel is meticulously dissected and released from the surrounding fascia and then dilated with sutures allowing for insertion and subsequent securing of the catheter into the vessel. The catheter is exteriorized and approximately 1 L/24 hr of lymph is collected over a 7 day period. While this technique enables the collection of large quantities of central lymph over an extended period of time, the success depends on careful surgical dissection, tissue handling and close attention to proper surgical technique. This is particularly important with surgeries in young animals as the lymph vessels can easily tear, potentially leading to surgical and experimental failure. The video demonstrates an excellent surgical technique for the collection of intestinal lymph.

We present a surgical procedure to catheterize the intestinal lymph trunk in neonatal pigs to collect large quantities of lipid metabolism components from efferent lymph.

Catheterization of the intestinal lymph trunk in neonatal pigs is a technique allowing for the long-term collection of large quantities of intestinal ...

Surgical Placement of Catheters for Long-term Cardiovascular Exercise Testing in Swine

This protocol describes the surgical procedure to chronically instrument swine and the procedure to exercise swine on a motor-driven treadmill. Early cardiopulmonary dysfunction is difficult to diagnose, particularly in animal models, as cardiopulmonary function is often measured invasively, requiring anesthesia. As many anesthetic agents are cardiodepressive, subtle changes in cardiovascular function may be masked. In contrast, chronic instrumentation allows for measurement of cardiopulmonary function in the awake state, so that measurements can be obtained under quiet resting conditions, without the effects of anesthesia and acute surgical trauma. Furthermore, when animals are properly trained, measurements can also be obtained during graded treadmill exercise.
Flow probes are placed around the aorta or pulmonary artery for measurement of cardiac output and around the left anterior descending coronary artery for measurement of coronary blood flow. Fluid-filled catheters are implanted in the aorta, pulmonary artery, left atrium, left ventricle and right ventricle for pressure measurement and blood sampling. In addition, a 20 G&#160;catheter is positioned in the anterior interventricular vein to allow coronary venous blood sampling.
After a week of recovery, swine are placed on a motor-driven treadmill, the catheters are connected to pressure and flow meters, and swine are subjected to a five-stage progressive exercise protocol, with each stage lasting 3 min. Hemodynamic signals are continuously recorded and blood samples are taken during the last 30 sec of each exercise stage.
The major advantage of studying chronically instrumented animals is that it allows serial assessment of cardiopulmonary function, not only at rest but also during physical stress such as exercise. Moreover, cardiopulmonary function can be assessed repeatedly during disease development and during chronic treatment, thereby increasing statistical power and hence limiting the number of animals required for a study.

Here we present a protocol to assess cardiopulmonary function in awake swine, at rest and during graded treadmill exercise. Chronic instrumentation allows for repeated hemodynamic measurements uninfluenced by cardiodepressive anesthetic agents.

This protocol describes the surgical procedure to chronically instrument swine and the procedure to exercise swine on a motor-driven treadmill. Early ...

Long-term Blood Pressure Measurement in Freely Moving Mice Using Telemetry

During the development of new vasoactive agents, arterial blood pressure monitoring is crucial for evaluating the efficacy of the new proposed drugs. Indeed, research focusing on the discovery of new potential therapeutic targets using genetically altered mice requires a reliable, long-term assessment of the systemic arterial pressure variation. Currently, the gold standard for obtaining long-term measurements of blood pressure in ambulatory mice uses implantable radio-transmitters, which require&#160;artery cannulation. This technique eliminates the need for tethering, restraining, or anesthetizing the animals which introduce stress and artifacts during data sampling. However, arterial blood pressure monitoring in mice via catheterization can be rather challenging due to the small size of the arteries. Here we present a step-by-step guide to illustrate the crucial key passages for a successful subcutaneous implantation of radio-transmitters and carotid artery cannulation in mice. We also include examples of long-term blood pressure activity taken from freely moving mice after a period of post-surgery recovery. Following this procedure will allow reliable direct blood pressure recordings from multiple animals simultaneously.

The goal of this protocol is to assess systemic blood pressure in conscious freely moving mice using implantable radio-telemetry devices.

During the development of new vasoactive agents, arterial blood pressure monitoring is crucial for evaluating the efficacy of the new proposed drugs. ...

3D Whole-heart Myocardial Tissue Analysis

Cardiac regenerative therapies aim to protect and repair the injured heart in patients with ischemic heart disease. By injecting stem cells or other biologicals that enhance angio- or vasculogenesis into the infarct border zone (IBZ), tissue perfusion is improved, and the myocardium can be protected from further damage. For maximum therapeutic effect, it is hypothesized that the regenerative substance is best delivered to the IBZ. This requires accurate injections and has led to the development of new injection techniques. To validate these new techniques, we have designed a validation protocol based on myocardial tissue analysis. This protocol includes whole-heart myocardial tissue processing that enables detailed two-dimensional (2D) and three-dimensional (3D) analysis of the cardiac anatomy and intramyocardial injections. In a pig, myocardial infarction was created by a 90-min occlusion of the left anterior descending coronary artery. Four weeks later, a mixture of a hydrogel with superparamagnetic iron oxide particles (SPIOs) and fluorescent beads was injected in the IBZ using a minimally-invasive endocardial approach. 1 h after the injection procedure, the pig was euthanized, and the heart was excised and embedded in agarose (agar). After the solidification of the agar, magnetic resonance imaging (MRI), slicing of the heart, and fluorescence imaging were performed. After image post-processing, 3D analysis was performed to assess the IBZ targeting accuracy. This protocol provides a structured and reproducible method for the assessment of the targeting accuracy of intramyocardial injections into the IBZ. The protocol can be easily used when the processing of scar tissue and/or validation of the injection accuracy of the whole heart is desired.

This protocol describes a novel method for the 3D comparison of whole-heart myocardial tissue with MRI. This is designed for the accurate assessment of intramyocardial injections in the infarct border zone of a chronic porcine model of myocardial infarction.

Cardiac regenerative therapies aim to protect and repair the injured heart in patients with ischemic heart disease. By injecting stem cells or other ...

Improvement of a Closed Chest Porcine Myocardial Infarction Model by Standardization of Tissue and Blood Sampling Procedures

Myocardial ischemia reperfusion (I/R) injury contributes to almost half of the necrotic area after myocardial infarction. To date there is no approved drug to prevent or reduce myocardial I/R injury. The study and understanding of the pathophysiological mechanisms of myocardial I/R injury is essential to develop successful treatments. Large animal experiments are an important step in translational methods. The porcine model of acute myocardial infarction has been established and described by ourselves and others. We aimed to further improve the value of the model by focusing in detail on the sampling techniques for use in future experiments. Furthermore, we emphasize small but important steps that can affect the quality of the final results. To mimic the clinical situation of myocardial I/R injury, a percutaneous coronary intervention (PCI) catheter was inserted into the left anterior descending coronary artery (LAD) of an anesthetized pig. &#176;&#176;&#176;This model mimics acute myocardial infarction and PCI treatment in humans with the possibility of accurately determining the area at risk as well as the necrotic- and viable ischemic tissue. Here the model was used to investigate the effect of a bicyclic peptide inhibitor of FXIIa. The model can also be modified to allow longer reperfusion times to study later effects of myocardial infarction.

Here we demonstrate a protocol to standardize sampling procedures of an established porcine model of acute myocardial infarction in order to increase its translational value in the understanding of the pathophysiology of myocardial ischemia/reperfusion injury and to test novel drug candidates.

Myocardial ischemia reperfusion (I/R) injury contributes to almost half of the necrotic area after myocardial infarction. To date there is no approved ...

Analyzing Long-Term Electrocardiography Recordings to Detect Arrhythmias in Mice

Arrhythmias are common, affecting millions of patients worldwide. Current treatment strategies are associated with significant side effects and remain ineffective in many patients. To improve patient care, novel and innovative therapeutic concepts causally targeting arrhythmia mechanisms are needed. To study the complex pathophysiology of arrhythmias, suitable animal models are necessary, and mice have been proven to be ideal model species to evaluate the genetic impact on arrhythmias, to investigate fundamental molecular and cellular mechanisms, and to identify potential therapeutic targets.
Implantable telemetry devices are among the most powerful tools available to study electrophysiology in mice, allowing continuous ECG recording over a period of several months in freely moving, awake mice. However, due to the huge number of data points (&gt;1 million QRS complexes per day), analysis of telemetry data remains challenging. This article describes a step-by-step approach to analyze ECGs and to detect arrhythmias in long-term telemetry recordings using the software, Ponemah, with its analysis modules, ECG Pro and Data Insights, developed by Data Sciences International (DSI). To analyze basic ECG parameters, such as heart rate, P wave duration, PR interval, QRS interval, or QT duration, an automated attribute analysis was performed using Ponemah to identify P, Q, and T waves within individually adjusted windows around detected R waves. 
Results were then manually reviewed, allowing adjustment of individual annotations. The output from the attribute-based analysis and the pattern recognition analysis was then used by the Data Insights module to detect arrhythmias. This module allows an automatic screening for individually defined arrhythmias within the recording, followed by a manual review of suspected arrhythmia episodes. The article briefly discusses challenges in recording and detecting ECG signals, suggests strategies to improve data quality, and provides representative recordings of arrhythmias detected in mice using the approach described above.

Here we present a step-by-step protocol for a semiautomated approach to analyze murine long-term electrocardiography (ECG) data for basic ECG parameters and common arrhythmias. Data are obtained by implantable telemetry transmitters in living and awake mice and analyzed using Ponemah and its analysis modules.

Arrhythmias are common, affecting millions of patients worldwide. Current treatment strategies are associated with significant side effects and remain ...

Technique for Obtaining Mesenchymal Stem Cell from Adipose Tissue and Stromal Vascular Fraction Characterization in Long-Term Cryopreservation

Human mesenchymal stem cells derived from adipose tissue have become increasingly attractive as they show appropriate features and are an accessible source for regenerative clinical applications. Different protocols have been used to obtain adipose-derived stem cells. This article describes different steps of an improved time-saving protocol to obtain a more significant amount of ADSC, showing how to cryopreserve and thaw ADSC to obtain viable cells for culture expansion. One hundred milliliters of lipoaspirate were collected, using a 26 cm three-hole and 3 mm caliber syringe liposuction, from the abdominal area of nine patients who subsequently underwent elective abdominoplasty. The stem cells isolation was carried out with a series of washes with Dulbecco&#39;s Phosphate Buffered Saline (DPBS) solution supplemented with calcium and the use of collagenase. Stromal Vascular Fraction (SVF) cells were cryopreserved, and their viability was checked by immunophenotyping. The SVF cellular yield was 15.7 x 105 cells/mL, ranging between 6.1-26.2 cells/mL. Adherent SVF cells reached confluence after an average of 7.5 (&#177;4.5) days, with an average cellular yield of 12.3 (&#177; 5.7) x 105 cells/mL. The viability of thawed SVF after 8 months, 1 year, and 2 years ranged between 23.06%-72.34% with an average of 47.7% (&#177;24.64) with the lowest viability correlating with cases of two-year freezing. The use of DPBS solution supplemented with calcium and bag resting times for fat precipitation with a shorter time of collagenase digestion resulted in an increased stem cell final cellular yield. The detailed procedure for obtaining high yields of viable stem cells was more efficient regarding time and cellular yield than the techniques from previous studies. Even after a long period of cryopreservation, viable ADSC cells were found in the SVF.

The present protocol describes an improved methodology for ADSC isolation resulting in a tremendous cellular yield with time gain compared to the literature. This study also provides a straightforward method for obtaining a relatively large number of viable cells after long-term cryopreservation.

Human mesenchymal stem cells derived from adipose tissue have become increasingly attractive as they show appropriate features and are an accessible ...

Long-Term Continuous Measurement of Renal Blood Flow in Conscious Rats

The kidneys play a crucial role in maintaining the homeostasis of body fluids. The regulation of renal blood flow (RBF) is essential to the vital functions of filtration and metabolism in kidney function. Many acute studies have been carried out in anesthetized animals to measure RBF under various conditions to determine mechanisms responsible for the regulation of kidney perfusion. However, for technical reasons, it has not been possible to measure RBF continuously (24 h/day) in unrestrained unanesthetized rats over prolonged periods. These methods allow the continuous determination of RBF over many weeks while also simultaneously recording blood pressure (BP) with implanted catheters (fluid-filled or by telemetry). RBF monitoring is carried out with rats placed in a circular servo-controlled rat cage that enables the unrestrained movement of the rat throughout the study. At the same time, the tangling of cables from the flow probe and arterial catheters is prevented. Rats are first instrumented with an ultrasonic flow probe placement on the left renal artery and an arterial catheter implanted in the right femoral artery. These are routed subcutaneously to the nape of the neck, and connected to the flowmeter and pressure transducer, respectively, to measure RBF and BP. Following surgical implantation, rats are immediately placed in the cage to recover for at least one week and stabilize the ultrasonic probe recordings. Urine collection is also feasible in this system. The surgical and post-surgical procedures for continuous monitoring are demonstrated in this protocol.

The present protocol describes a long-term continuous measurement of renal blood flow in conscious rats and simultaneously recording blood pressure with implanted catheters (fluid-filled or by telemetry).