A special thanks to Nat Johnston for the recording and editing the video.
Poonam Sharma was supported by University of Newcastle with UNIPRS and UNRS Central &#38; Faculty School (UNRSC5050) scholarships. Carmine Gentile was supported by a UTS Seed Funding, Catholic Archdiocese of Sydney Grant for Adult Stem Cell Research and a Sydney Medical School Foundation Cardiothoracic Surgery Research Grant.

NOTE: hiPSC-CMs used for this protocol are commercially available. Please seek institutional human ethics committee approval before commencing this work if required.
1. Human cardiac fibroblast&#160;and endothelial cell culture plating and growth
<ol>
	<li>Thaw cryovials containing HCFs and HCAECs in a water bath at 37 &#176;C for one minute.</li>
	<li>Move cryovials under a sterile laminar flow biosafety cabinet class 2.</li>
	<li>Collect 1 mL of cell suspension from the cryovials using a 1000 &#181;L pipette tip and add into a 15 mL tube containing 7 mL of Human Cardiac Fibroblast Medium for HCFs and 7 mL of Human Meso Endo Growth Medium for HCAECs. 
	NOTE: In order to collect the majority of the cells from each cryovial, rinse them twice with 1 mL of culture medium from the same 15 mL tube.</li>
	<li>Gently mix cell suspensions.</li>
	<li>Transfer cell suspensions to separate T75 culture flasks using a 10 mL serological pipette.</li>
	<li>Incubate cells at 37 &#176;C with 5% CO2.</li>
	<li>After 18 h, aspirate the medium from both culture flasks and rinse them once with sterile phosphate buffered saline (PBS) to remove freezing medium and dead cells.</li>
	<li>Replace PBS with 7 mL of appropriate culture medium to each culture flask and incubate at 37 &#176;C.</li>
	<li>Examine cellular expansion and viability regularly and replace media every other day until cells reach 80-90% confluency.</li>
</ol>
2. iCM&#160;culture plating and growth
<ol>
	<li>Pre-coat two T25 culture flasks with 2 mL of PBS containing 40 &#181;g/mL of fibronectin (FN) and incubate at 37 &#176;C, 5% CO2 for at least 4 hours.</li>
	<li>After 4 hours, collect one cryovial containing iCMs and place it in a water bath at 37 &#176;C for 4 min.</li>
	<li>Move the cryovial under a sterile laminar flow biosafety cabinet class 2.</li>
	<li>Gently transfer the iCMs from the cryovial to a sterile 50 mL centrifuge tube using a 1 mL pipette tip.</li>
	<li>Rinse the empty iCMs cryovial with 1 mL of room temperature plating medium to recover any residual cells. Transfer the 1 mL of plating medium rinse from the cryovial drop-wise over 90 sec to the 50 mL centrifuge tube containing the iCM cell suspension. 
	NOTE: Gently swirl the tube while adding the medium to mix the solution completely and to decrease the osmotic shock on the thawed cells.</li>
	<li>Slowly add 8 mL of room temperature Plating Medium to the 50 mL centrifuge tube. Add the first 1 mL dropwise over 30 - 60 s. Then, add the remaining volume over the next 30 s. Gently swirl the centrifuge tube while adding the Plating medium. Gently mix the contents of the 50 mL centrifuge tube by inverting 2 - 3 times (avoiding vigorous shaking or vortexing).</li>
	<li>Immediately perform the cell counting using a hemocytometer and determine the viable cell density (in cells/mL).</li>
	<li>Take the FN-pre-coated T25 flasks and aspirate the FN-PBS solution without letting the flasks dry. To this add seeding volume of iCMs (1.6 x 106 viable iCMs&#160;in 8 mL room temperature plating medium).</li>
	<li>Culture iCMs in the incubator for 48 h at 37 &#176;C, 5% CO2.</li>
	<li>Thaw the Maintenance Medium overnight at 4 &#176;C a day before use.</li>
	<li>Equilibrate the Maintenance Medium in a 37 &#176;C water bath and use immediately.</li>
	<li>After 2 days, move the iCMs T25 flasks under the biosafety cabinet.</li>
	<li>Gently wash off dead cells and debris by gently pipetting the Plating Medium up and down 5 times.</li>
	<li>Aspirate the Plating Medium and replace with the 8 mL of pre-warmed Maintenance Medium. Place the T25 flasks back in the incubator. Replace the Maintenance Medium every other day and examine the confluency regularly.</li>
</ol>
3. Cell isolation and counting
<ol>
	<li>Start by collecting first HCAECs and HCFs, and then iCMs by following steps 3.2-3.12.</li>
	<li>Prepare CS culture medium by mixing 10 mL of iCMs Maintenance Medium, 5 mL of Human Cardiac Fibroblast Medium and 5 mL of Meso Endo Growth Medium.</li>
	<li>Remove culture medium from each tissue flask containing HCFs and HCAECs and rinse once with 5 mL PBS for T75 flasks. Remove PBS.</li>
	<li>Add 5 mL of 0.25% trypsin EDTA solution to each T75 flask and incubate for 5 min at 37 &#176;C, 5% CO2.</li>
	<li>Once cells detach, immediately neutralize the trypsin EDTA solution with 5 mL of culture medium.</li>
	<li>Transfer cell suspensions to a 15 mL tube and centrifuge cells at 300 x g for 4 min.</li>
	<li>Remove the supernatant carefully from each tube. Add 1 mL of CS medium to each cell pellet and resuspend them. Keep the tube on ice and count cells using Trypan Blue and a hemocytometer.</li>
	<li>Remove Maintenance Medium from tissue flasks containing iCMs and rinse once with 3 mL of PBS.</li>
	<li>Add 1 mL of 0.25% trypsin EDTA solution to each T75 flask and incubate at 37 &#176;C, 5% CO2. Check cells every minute until detached.</li>
	<li>Once cells detach, immediately neutralize the trypsin EDTA solution with 4 mL of culture medium.</li>
	<li>Transfer cell suspensions to a 15 mL tube and centrifuge them at 300 x g for 5 min.</li>
	<li>Remove the supernatant carefully from each tube. Add 1 mL of culture medium to the cell pellet and resuspend it. Keep the tube on ice and count cells using Trypan Blue and a hemocytometer.</li>
</ol>
4. CS&#160;generation and growth
<ol>
	<li>Mix iCMs, HCFs and HCAECs in 2:1:1 ratio by plating 10,000 iCMs, 5,000 HCFs and 5,000 HCAECs per hanging drop culture containing 20 &#181;L of CS medium. Adjust to the final volume for the total number of CSs.</li>
	<li>Pipette 20 &#181;L of cell suspension into each well of the 384 well HDC plate either manually or by using a robotic multichannel pipette for automated liquid handling.</li>
	<li>Pipette 1.5 mL of sterile PBS in each side of the channel around the Hanging Drop Plate to prevent drying out CSs. Incubate HDC plate at 37 &#176;C.</li>
	<li>Examine formation of CSs on a daily basis until a fully formed spheroid is observed in the majority of wells. Add 7.5 &#181;L of CS medium to each well every other day until a CS is formed.</li>
</ol>
5. CS&#160;embedding in collagen gels
<ol>
	<li>Collect CSs with a 1 mL pipette tip. 
	NOTE: It is necessary to cut the tip of pipette around 0.2 cm from the edge before its use with a sterile sharp surface (either a scalpel or a scissor) to prevent any damage gel embedded spheroids during their collection.</li>
	<li>Collect the CS suspension into a 50 mL tube on ice.</li>
	<li>Centrifuge the tube at 300 x g for 5 min. 
	NOTE: the pellet obtained must be kept on ice until use.</li>
	<li>Prepare a collagen gel solution (100 &#181;L/well for 30 wells of 96 well plate) on ice using rat tail collagen and CS medium in a 3:7 ratio.</li>
	<li>Remove the supernatant from the tube containing CSs.</li>
	<li>Mix the pelleted CSs within the collagen gel solution.</li>
	<li>Add 1 &#181;L/mL of 5 mM sodium hydroxide to the CS-collagen gel suspension and mix gently.</li>
	<li>Transfer 100 &#181;L of CS-collagen gel to a clear flat bottom 96 well black polystyrene microplate and incubate at 37 &#176;C for 30 min.</li>
</ol>
6. Viability and toxicity measurements of DOX-treated CSs
<ol>
	<li>After 30 min collect the 96 well plate from incubator</li>
	<li>Prepare a 10 &#181;M DOX (based on previously established protocol for cell death in CSs14). 
	&#8203;NOTE: To potentially test other agents that may protect against HF in CSs, generate solutions containing DOX + Agent A, B, etc.</li>
	<li>Add 100 &#181;L of solutions containing DOX and/or other agents to each well. Control cultures contain media without any DOX.</li>
	<li>Incubate plate for 18 h at 37 &#176;C, 5% CO2.</li>
	<li>On the following day, collect the Live/Dead staining reagent stock solutions and allow them to thaw on ice in the dark in a biosafety cabinet.</li>
	<li>Prepare a solution containing Hoechst stain, 4 &#181;M of ethidium homodimer and 2 &#181;M calcein-AM.</li>
	<li>Add 100 &#181;L of Hoechst stain, calcein-AM/ethidium homodimer solution into each well.</li>
	<li>Measure the fluorescence into each well at 645 nm for ethidium homodimer and at 530 nm for calcein-AM, respectively, using multimode microplate reader.</li>
	<li>Transfer fluorescence measurements into Graphpad PRISM (or an equivalent software for statistical analysis).</li>
	<li>Use GraphPad Prism software for data analysis and statistics.</li>
	<li>For quality control, check under an epifluorescence microscope for the nuclei staining, together with calcein-AM and ethidium homodimer.</li>
</ol>
7. CS contractile function evaluation
<ol>
	<li>Collect the microplate as prepared in step 5.8.</li>
	<li>Turn the on the computer containing the IonOptix software for a video-based edge-detection, the Fluorescence Assistance Interface, and the MyoPacer Field Stimulator.</li>
	<li>Place a new cover slip on the tissue holder platform and assemble water bath with electrodes.</li>
	<li>Gently collect CSs from collagen gels using a 1 mL pipette tip cut 0.5 mm from the edge and transfer them to a falcon tube. Add media onto CS to prevent drying of CSs. Transfer CS (one at a time) with a few &#181;L of media on the stage of the IonOptix system.</li>
	<li>Select the CS to be analysed via setting peaks on left and right side of CS using the IonOptix software.</li>
	<li>Use the computer-based motion analyzer to track the movement of CS edges. 
	NOTE: Normally, contractility is measured in either % cell shortening or % fractional shortening. In this case, we measured % spheroid shortening.</li>
	<li>Stabilize both the peaks adjusting threshold and edge options from the computer.</li>
	<li>Expose CSs to different frequencies (0.3, 0.6, 1, 2 and 3 Hz) and voltages (1, 2, 3 and 5 V) using the Myopacer Field Stimulator.</li>
	<li>Record spheroid shortening as CS length changes of DOX-treated and untreated CSs. Analyse data using the Soft-Edge software and averaged for each CS.</li>
</ol>
8. Microscopy of CSs: fixation and immunolabeling
<ol>
	<li>Collect the 96 well plate after 30 min (as prepared in step 5.8) and fix CSs in 4% paraformaldehyde (PFA) for 1 h at room temperature.</li>
	<li>Remove PFA and rinse three times with PBS containing 0.01% sodium azide (PBSA).</li>
	<li>Remove PBSA.</li>
	<li>Add 200 &#181;L of PBSA containing 0.02% Triton-X-100 to each well for 30 min on a shaker. 
	NOTE: This step permeabilizes CSs for better antibody infiltration.</li>
	<li>Add 200 &#181;L of 3% bovine serum albumin in PBSA solution for 60 min at room temperature. 
	NOTE: This step blocks unspecific antibody binding in CSs.</li>
	<li>Prepare a solution containing 10 &#181;g/mL of primary mouse anti-human antibodies against CD31 diluted in blocking solution.</li>
	<li>Add 100 &#181;L of primary antibody solution to each well and incubate overnight at 4 &#176;C on a shaker.</li>
	<li>Rinse the plate three times with PBSA for 20 min at room temperature on a rocking plate.</li>
	<li>Prepare a solution containing Hoechst DNA stain and 10 &#181;g/mL of Cy3-conjugated secondary donkey anti-mouse antibody diluted in blocking solution.</li>
	<li>Add 100 &#181;L of secondary antibody solution containing Hoechst stain to each well and incubate overnight at 4 &#176;C on a shaker. 
	NOTE: Cover the plate with aluminium foil from this point onwards.</li>
	<li>Rinse the plate three times for 20 min with PBSA at room temperature on a rocking plate.</li>
	<li>Add 100 &#181;L of Vectashield mounting medium to each well.</li>
	<li>Image CSs under a laser scanning confocal microscope. Perform optical sectioning along the Z axis and collapse images into a single focal plane using ImageJ software.</li>
</ol>

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Developmentally, proper vascular network formation is critical for the generation of functional tissues, including the human heart10,12,23,24,25,26. Consideration for the proper vascularization of 3D tissues allow the exchange of oxygen, growth factors, signalling molecules and nutrients, preventing the development of cell necrosis within any tissue thicker than 200 &#956;m6,10,12,17,24,25,26,27,28. Currently available in vitro 3D heart models that present a vascular network are primarily presenting capillary-sized, disorganized vascular networks and lack the hierarchical complex branched vascularization observed in vivo6,8,29. The alternative approach to develop complex cardiac endothelial cell network described in this manuscript presents improved cell viability and function compared to existing models (Figure 1)14,22. 3D in vitro CSs model the human heart by better recapitulating its in vivo microenvironment, including its molecular, cellular and extracellular components14,22. CS generation from stem cell-derived cells in the hanging drops allow their cultures in defined conditions (e.g., cell types and ratio, proper tissue formation). Co-cultures of iCMs together with HCFs and HCAECs within CSs define the molecular and cellular crosstalk that regulates the heart pathophysiology, including its contractile function and response to drugs at concentrations found in the patient&#39;s bloodstream14. Due to these unique features, CSs have been utilized to model cardiac fibrosis, a severe consequence of myocardial infarction and heart failure21. Our previous studies showed how the presence of both endothelial cells and fibroblasts is critical for the recapitulation of the vascular microenvironment in the human heart, allowing the optimal deposition of fibroblast-derived ECM proteins, such as laminin, fibronectin and collagen type IV, localized in proximity of a developing endothelial cell network14,21.
DOX is a well-known cardiotoxic drug that may develop heart failure in cancer patients even 17 years after their treatment30. Nevertheless, it remains a drug of choice for the treatment of leukemia and lymphoma in paediatric patients and breast cancer in women30. DOX treatment in CSs has then been used to model heart failure (HF) in vitro to study both the mechanisms regulating toxicity in cardiac myocytes, endothelial cells and fibroblasts14 and to model HF-induced cardiac fibrosis21. Cell viability was statistically reduced in DOX treated CSs within 24 h when exposed to the drug at the concentration found in the bloodstream of cancer patients (between 5 and 10 &#956;M)14 (Figure 2). Previous studies in our laboratory also demonstrated the toxic effects of DOX on both cardiac endothelial cells and fibroblasts via endothelial nitric oxide synthase (eNOS) using both genetic and chemical inhibitors of this signaling pathway14. The use of genetic (NOS3 shRNA) and chemical (N5-(1-iminoethyl)-L-ornithine, dihydrochloride, or L-NIO) antagonists of the eNOS signaling pathway as a downstream target of DOX prevented its toxic effects in both cardiac endothelial cells and fibroblasts14.
Contractile activity within CSs has also been measured thanks to the electrical coupling of cardiac cells when exposed to field potential stimulation. We found that CSs cultured with control media (DOX 0 &#956;M) contract spontaneously and homogenously at a beating rate that can be paced by field stimulation within 1 and 3 Hz, comparable with a healthy human heart. On the other hand, DOX-treated CSs do not follow the electrical stimulation as they cannot contract. Together with the measurements of cell viability and toxicity using calcein-AM and ethidium homodimer, this functional assay for CS contractile function allow the evaluation of the complex scenario typical of the human heart in vitro, currently not achievable with other models. Compared to contractile activity measurements of single cardiac cells using the same system, we are not able to visualize and measure the sarcomere in CSs. Therefore, we are limited to measurements of % spheroid shortening over time, an assay we had to develop within our laboratory. As we control the number of cells, we co-culture in each CS and therefore the size of each CS, we utilize CSs with similar size that indeed present homogenous contractile function. However, even in case we generated CSs of different sizes, their contractile activity did not change.
It is also important to report that the multicellular nature of CSs makes them heavy enough to localize at the bottom of the coverslip in the Ion-Optix system, even in case they are superfused. Based on the fact that CSs sit by themselves in a specific position, we do not need to make them adhere to the coverlip, on the contrary of what is commonly done with single cardiac cells in most laboratories.
The microscopic analysis of CSs stained with antibodies against cardiac troponin T, CD31/PECAM, and PECAM (as markers for iCMs, HCAECs, and HCFs, respectively) showed the formation of a endothelial cell network (Figure 1, blue). To fully exclude necrosis in the inner part of CSs, spatial evaluation of cell viability was performed in our laboratory by confocal analysis of calcein-AM/ethidium homodimer stained CSs (data not shown). However, it is important to acknowledge that future developments in the biofabrication field to better recapitulate other complex features typical of the human heart in vivo, currently not available in the existing model. These include: i) contractile function typical of adult cardiomyocytes; ii) blood flow and pressure forces; iii) paracrine signaling; iv) immune response, which will be critical to improve this and other in vitro cardiac models6. As any other model aims at recapitulating major features of either a healthy tissue or a disease state, the protocol for the generation and use of CS described in this manuscript aims at helping the researcher at addressing specific questions, that may not be exhaustive using this approach. For instance, the potential use of patient-derived cells for the generation of CSs would provide tools for personalized medicine, currently not available using commonly available high-throughput assays for cardiovascular research.
In conclusion, we demonstrated a simple way to better recapitulate the human heart microenvironment using cardiac cells. Cardiac spheroids present an endothelial cell network that better recapitulates the one present in the human heart compared to monolayer cultures of cardiac cells.Given their unique features, they represent advanced tools for in vitro testing for cardiovascular research. Future studies using patient-derived cells could provide options for personalized medicine and novel therapies to both prevent and better treat cardiovascular disease.

The human heart has a limited regenerative capacity while cardiovascular disease (CVD) remains the main cause of death worldwide despite the recent advances in tissue engineering and stem cell technologies1. The need for novel therapeutics including molecular and cellular approaches to either repair a damaged heart or to prevent a heart from failing is one of the major current clinical needs for patients suffering from heart disease2,3,4. The main goal of cardiac tissue engineering is to fabricate a three-dimensional (3D) heart tissue that presents molecular, cellular, and extracellular features typical of a human heart, including its vascular network and physiological contractile function4,5,6.
In order to bioengineer and fabricate a functional human cardiac tissue that mimics the human heart for in vitro and in vivo applications, several approaches have been investigated including engineered heart tissues (EHTs), cell sheets and spheroid cultures7,8. However, these tissues fail at recapitulating the optimal 3D microenvironment typical of the human heart and their potential use for CVD patients cannot directly translate from the bench to the bedside7. This is because they do not recapitulate the complex biology, morphology, and physiology of in vivo heart tissues9. One of the major challenges in cardiac tissue engineering includes the development of a hierarchical vascular network within the bioengineered cardiac tissue, as any tissue that is bigger than 200 &#956;m in diameter develops cell death in the middle2,10. A properly formed vascular network in a human heart tissue plays an major role for the supply of blood, oxygen and nutrients to cardiac cells11. During embryonic development, coronary capillaries and arteries form via vasculogenesis (de novo blood vessel formation) and angiogenesis (generation of blood vessels from pre-existing ones) from endothelial progenitor cells8,12. Cardiac fibroblasts also play a major role in proper vascular network formation by providing the optimal extracellular matrix (ECM) and growth composition13,14.
The 3D vascular network of bioengineered heart tissues controls cell survival and function by creating oxygen and nutrient gradients and paracrine signaling, such as homotypic cell interaction, heterotypic cell interaction, interaction of cells through secreted soluble proteins and cell to ECM interactions3,10,15,16,17,18. This prevents cell death in the middle of the tissue and promotes cell viability and physiological function in bioengineered heart tissues16,18,19.
Spheroid cultures from stem cells have been recently explored as in vitro models of the human heart20. To further improve the cardiac microenvironment in vitro, they have included the use of all the main cell types found in the human heart, such as cardiac myocytes, endothelial cells, and fibroblasts. Spheroid cultures present the required 3D structural support for cells to grow and function and can be used to bioengineer a vascular network14,20,21,22. In this context, our laboratory has developed human cardiac spheroids (CSs) by co-culturing cardiac myocytes, endothelial cells and fibroblasts at ratios found in the human heart14. This model is an expansion of the rat ventricular cardiac cells spheroid model, generated by co-culturing cardiac cells in hanging drop cultures, used to model cardiac fibrosis21. Human CSs can be used as toxicity assays by treating them doxorubicin (DOX, an anti-cancer agent used to treat leukemia, lymphoma and breast cancer), which is well-known to induce cardiac fibrosis and heart failure (HF) even 17 years following its somministration14.
In this manuscript, we describe how to generate human CSs by co-culturing human induced pluripotent stem cell derived cardiomyocytes (hiPSC-CMs or iCMs), human cardiac fibroblasts (HCFs) and human coronary artery endothelial cells (HCAECs) in hanging drop cultures. In order to use and image CSs for in vitro testing, they are embedded in a collagen gel. The confocal analysis of CSs stained with antibodies against CD31, a marker for endothelial cells, showed that these cells form a network similar to the one observed in vivo. To induce HF and potentially test novel agents that may treat or prevent it, CSs were treated with 10 &#181;M DOX (a concentration found in the bloodstream of cancer patients receiving the drug). When stained with calcein-AM and ethidium homodimer (staining live and dead cells, respectively), DOX-treated CSs present a significant decrease in viability in comparison to CSs that did not receive the drug. CSs also present a homogeneous contractile activity when paced using field potential stimulation between 1 and 3 Hz.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Bovine Serum Albumin (BSA)</td>
			<td>Sigma-Aldrich</td>
			<td>A1933</td>
			<td></td>
		</tr>
		<tr>
			<td>Donkey anti-mouse Secondary Antibodies</td>
			<td>Jackson Immunological Research Labs, Inc.</td>
			<td>715-165-150</td>
			<td>Cyanine Cy3-conjugated secondary antibody</td>
		</tr>
		<tr>
			<td>Doxorubicin hydrochloride</td>
			<td>Sigma-Aldrich</td>
			<td>D1515</td>
			<td></td>
		</tr>
		<tr>
			<td>Fibronectin</td>
			<td>Sigma-Aldrich</td>
			<td>F1141-1MG</td>
			<td>From Bovine Plasma</td>
		</tr>
		<tr>
			<td>Human cardiac fibroblasts (HCFs)</td>
			<td>Cell Applications, Inc., San Diego, CA, USA</td>
			<td>306AK-05a</td>
			<td>5x10^5 Cells (Adult), Medium &#38; Subculture Reagents</td>
		</tr>
		<tr>
			<td>Human coronary artery endothelial cells (HCAECs)</td>
			<td>Cell Applications, Inc., San Diego, CA, USA</td>
			<td>300K-05a</td>
			<td>5x10^5 Cells (Adult), Medium &#38; Subculture Reagents</td>
		</tr>
		<tr>
			<td>Human iPSC-derived cardiomyocytes (iCMs)</td>
			<td>Fujifilm Cellular Dynamics, Inc.</td>
			<td>R1057</td>
			<td>iCell Cardiomyocytes Kit, 01434</td>
		</tr>
		<tr>
			<td>HCF Growth medium</td>
			<td>Cell Applications, Inc., San Diego, CA, USA</td>
			<td>316-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Human MesoEndo Cell Growth Medium</td>
			<td>Cell Applications, Inc., San Diego, CA, USA</td>
			<td>212-500</td>
			<td></td>
		</tr>
		<tr>
			<td>LIVE/DEAD Viability/Cytotoxicity Kit</td>
			<td>Invitrogen, Carlsbad, CA, USA</td>
			<td>L3224</td>
			<td></td>
		</tr>
		<tr>
			<td>Maintenance Medium (iCells)</td>
			<td>Fujifilm Cellular Dynamics, Inc.</td>
			<td>R1057</td>
			<td>iCell Cardiomyocytes Kit, 01434</td>
		</tr>
		<tr>
			<td>Mouse Monoclonal anti-human CD31/PECAM</td>
			<td>BD Pharmingen, San Diego, CA, USA</td>
			<td>566177</td>
			<td></td>
		</tr>
		<tr>
			<td>NucBlue Live ReadyProbe Reagent (Hoechst 33342)</td>
			<td>Invitrogen, Carlsbad, CA, USA</td>
			<td>R37605</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Sigma-Aldrich</td>
			<td>P6148</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate-Buffered Saline</td>
			<td>Sigma-Aldrich</td>
			<td>D8537</td>
			<td></td>
		</tr>
		<tr>
			<td>Plating Medium (iCells)</td>
			<td>Fujifilm Cellular Dynamics, Inc.</td>
			<td>R1057</td>
			<td>iCell Cardiomyocytes Kit, 01434</td>
		</tr>
		<tr>
			<td>Rat Tail Collagen</td>
			<td>Sigma-Aldrich</td>
			<td>C3867</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Azide</td>
			<td>Sigma-Aldrich</td>
			<td>S2002</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin&#8211;EDTA, 0.25%</td>
			<td>Gibco, Thermofisher Scientific</td>
			<td>25200072</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypan Blue Solution, 0.4%</td>
			<td>Gibco, Thermofisher Scientific</td>
			<td>15250061</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton-X 100</td>
			<td>Sigma-Aldrich</td>
			<td>X100</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue culture flasks (T25)</td>
			<td>Thermofisher Scientific</td>
			<td>156367</td>
			<td></td>
		</tr>
		<tr>
			<td>96-well Flat Clear Bottom Black Polystyrene TC-treated Microplates</td>
			<td>Corning, New York, USA</td>
			<td>3603</td>
			<td></td>
		</tr>
		<tr>
			<td>384-Well Hanging Drop Plate</td>
			<td>3D Biomatrix, Ann Arbor, MI, USA</td>
			<td>HDP1385</td>
			<td></td>
		</tr>
	</tbody>
</table>

cardiac spheroids as in vitro bioengineered heart tissues to study human heart pathophysiology

Despite several advances in cardiac tissue engineering, one of the major challenges to overcome remains the generation of a fully functional vascular network comprising several levels of complexity to provide oxygen and nutrients within bioengineered heart tissues. Our laboratory has developed a three-dimensional in vitro model of the human heart, known as the "cardiac spheroid" or "CS". This presents biochemical, physiological, and pharmacological features typical of the human heart and is generated by co-culturing its three major cell types, such as cardiac myocytes, endothelial cells, and fibroblasts. Human induced pluripotent stem cells-derived cardiomyocytes (hiPSC-CMs or iCMs) are co-cultured at ratios approximating the ones found in vivo with human cardiac fibroblasts (HCFs) and human coronary artery endothelial cells (HCAECs) in hanging drop culture plates for three to four days. The confocal analysis of CSs stained with antibodies against cardiac Troponin T, CD31 and vimentin (markers for cardiac myocytes, endothelial cells and fibroblasts, respectively) shows that CSs present a complex endothelial cell network, resembling the native one found in the human heart. This is confirmed by the 3D rendering analysis of these confocal images. CSs also present extracellular matrix (ECM) proteins typical of the human heart, such as collagen type IV, laminin and fibronectin. Finally, CSs present a contractile activity measured as syncytial contractility closer to the one typical of the human heart compared to CSs that contain iCMs only. When treated with a cardiotoxic anti-cancer agent, such as doxorubicin (DOX, used to treat leukemia, lymphoma and breast cancer), the viability of DOX-treated CSs is significantly reduced at 10 &#181;M genetic and chemical inhibition of endothelial nitric oxide synthase, a downstream target of DOX in HCFs and HCAECs, reduced its toxicity in CSs. Given these unique features, CSs are currently used as in vitro models to study heart biochemistry, pathophysiology, and pharmacology.

The protocol described in this manuscript represents an alternative approach to develop complex cardiac endothelial cell network within a bioengineered cardiac tissue with improved cell viability and function compared to existing models (Figure 1).&#160;The recapitulation of the 3D in vivo heart microenvironment within CSs promoted their response to DOX at the concentration found in the bloodstream of cancer patients (between 5 and 10 &#956;M, Figure 2). DOX treated CSs presented a statistically significant reduction in cell viability compared with control (no DOX) CSs within 24 h (Figure 2), a toxic effect that is observed in human cancer patients even 17 years after their treatment with the drug.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61962/61962fig01.jpg" /> 
Figure 1. CS Formation and Vascularization Analysis. (A) Protocol showing the steps for the formation of a CS from the co-culture of iCMs, HCAECs and HCFs in hanging drops. Brightfield images on the right side show the progressive spheroid formation from single cells in hanging drops. (B) Collapsed Z-stacks of confocal images of a CS stained with antibodies against cardiac Troponin T (cTNT), PECAM and vimentin, staining cardiac myocytes, endothelial cells and fibroblasts, respectively. This figure has been modified from14. <a href="https://www.jove.com/files/ftp_upload/61962/61962fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61962/61962fig02.jpg" /> 
Figure 2. Viability and Toxicity in CSs. Statistical analyses of calcein-AM (A) and ethidium homodimer (B) fluorescence of CSs treated in presence of either media only (DOX 0 &#956;M) or doxorubicin (10 &#956;M). <a href="https://www.jove.com/files/ftp_upload/61962/61962fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Cardiac Spheroids as in vitro Bioengineered Heart Tissues to Study Human Heart Pathophysiology at JoVE.com

Cardiac Spheroids as in vitro Bioengineered Heart Tissues to Study Human Heart Pathophysiology

This protocol aims to fabricate 3D cardiac spheroids (CSs) by co-culturing cells in hanging drops.&#160;Collagen-embedded CSs are treated with doxorubicin (DOX, a cardiotoxic agent) at physiological concentrations to model heart failure. In vitro testing using DOX-treated CSs may be used to identify novel therapies for heart failure patients.

Despite several advances in cardiac tissue engineering, one of the major challenges to overcome remains the generation of a fully functional vascular ...

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7.4K Views.  University of Newcastle. This protocol aims to fabricate 3D cardiac spheroids (CSs) by co-culturing cells in hanging drops. Collagen-embedded CSs are treated with doxorubicin (DOX, a cardiotoxic agent) at physiological concentrations to model heart failure. In vitro testing using DOX-treated CSs may be used to identify novel therapies for heart failure patients.

Video: Cardiac Spheroids as in vitro Bioengineered Heart Tissues to Study Human Heart Pathophysiology

Multiparametric Optical Mapping of the Langendorff-perfused Rabbit Heart

Optical imaging and fluorescent probes have significantly advanced research methodology in the field of cardiac electrophysiology in ways that could not have been accomplished by other approaches1. With the use of the calcium- and voltage-sensitive dyes, optical mapping allows measurement of transmembrane action potentials and calcium transients with high spatial resolution without the physical contact with the tissue. This makes measurements of the cardiac electrical activity possible under many conditions where the use of electrodes is inconvenient or impossible1. For example, optical recordings provide accurate morphological changes of membrane potential during and immediately after stimulation and defibrillation, while conventional electrode techniques suffer from stimulus-induced artifacts during and after stimuli due to electrode polarization1. 
The Langendorff-perfused rabbit heart is one of the most studied models of human heart physiology and pathophysiology. Many types of arrhythmias observed clinically could be recapitulated in the rabbit heart model. It was shown that wave patterns in the rabbit heart during ventricular arrhythmias, determined by effective size of the heart and the wavelength of reentry, are very similar to that in the human heart2. It was also shown that critical aspects of excitation-contraction (EC) coupling in rabbit myocardium, such as the relative contribution of sarcoplasmic reticulum (SR), is very similar to human EC coupling3. Here we present the basic procedures of optical mapping experiments in Langendorff-perfused rabbit hearts, including the Langendorff perfusion system setup, the optical mapping systems setup, the isolation and cannulation of the heart, perfusion and dye-staining of the heart, excitation-contraction uncoupling, and collection of optical signals. These methods could be also applied to the heart from species other than rabbit with adjustments to flow rates, optics, solutions, etc.
Two optical mapping systems are described. The panoramic mapping system is used to map the entire epicardium of the rabbit heart4-7. This system provides a global view of the evolution of reentrant circuits during arrhythmogenesis and defibrillation, and has been used to study the mechanisms of arrhythmias and antiarrhythmia therapy8,9. The dual mapping system is used to map the action potential (AP) and calcium transient (CaT) simultaneously from the same field of view10-13. This approach has enhanced our understanding of the important role of calcium in the electrical alternans and the induction of arrhythmia14-16.

This article describes the basic procedures for conducting optical mapping experiments in the Langendorff-perfused rabbit heart using the panoramic imaging system, and the dual (voltage and calcium) imaging modality.

Optical imaging and fluorescent probes have significantly advanced research methodology in the field of cardiac electrophysiology in ways that could not ...

Optical Mapping of Action Potentials and Calcium Transients in the Mouse Heart

The mouse heart is a popular model for cardiovascular studies due to the existence of low cost technology for genetic engineering in this species. Cardiovascular physiological phenotyping of the mouse heart can be easily done using fluorescence imaging employing various probes for transmembrane potential (Vm), calcium transients (CaT), and other parameters. Excitation-contraction coupling is characterized by action potential and intracellular calcium dynamics; therefore, it is critically important to map both Vm and CaT simultaneously from the same location on the heart1-4. Simultaneous optical mapping from Langendorff perfused mouse hearts has the potential to elucidate mechanisms underlying heart failure, arrhythmias, metabolic disease, and other heart diseases. Visualization of activation, conduction velocity, action potential duration, and other parameters at a myriad of sites cannot be achieved from cellular level investigation but is well solved by optical mapping1,5,6. In this paper we present the instrumentation setup and experimental conditions for simultaneous optical mapping of Vm and CaT in mouse hearts with high spatio-temporal resolution using state-of-the-art CMOS imaging technology. Consistent optical recordings obtained with this method illustrate that simultaneous optical mapping of Langendorff perfused mouse hearts is both feasible and reliable.

This paper details the dissection procedure, instrumental setup, and experimental conditions during optical mapping of transmembrane potential (Vm) and intracellular calcium transient (CaT) in intact isolated Langendorff perfused mouse hearts.

The mouse heart is a popular model for cardiovascular studies due to the existence of low cost technology for genetic engineering in this species. ...

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic valve leaflets are composed of interstitial cells suspended within an extracellular matrix (ECM) and are lined with an endothelial cell monolayer. The valve withstands a harsh, dynamic environment and is constantly exposed to shear, flexion, tension, and compression. Research has shown calcific lesions in diseased valves occur in areas of high mechanical stress as a result of endothelial disruption or interstitial matrix damage1-3. Hence, it is not surprising that epidemiological studies have shown high blood pressure to be a leading risk factor in the onset of aortic valve disease4. 
The only treatment option currently available for valve disease is surgical replacement of the diseased valve with a bioprosthetic or mechanical valve5. Improved understanding of valve biology in response to physical stresses would help elucidate the mechanisms of valve pathogenesis. In turn, this could help in the development of non-invasive therapies such as pharmaceutical intervention or prevention. Several bioreactors have been previously developed to study the mechanobiology of native or engineered heart valves6-9. Pulsatile bioreactors have also been developed to study a range of tissues including cartilage10, bone11 and bladder12. The aim of this work was to develop a cyclic pressure system that could be used to elucidate the biological response of aortic valve leaflets to increased pressure loads. 
The system consisted of an acrylic chamber in which to place samples and produce cyclic pressure, viton diaphragm solenoid valves to control the timing of the pressure cycle, and a computer to control electrical devices. The pressure was monitored using a pressure transducer, and the signal was conditioned using a load cell conditioner. A LabVIEW program regulated the pressure using an analog device to pump compressed air into the system at the appropriate rate. The system mimicked the dynamic transvalvular pressure levels associated with the aortic valve; a saw tooth wave produced a gradual increase in pressure, typical of the transvalvular pressure gradient that is present across the valve during diastole, followed by a sharp pressure drop depicting valve opening in systole. The LabVIEW program allowed users to control the magnitude and frequency of cyclic pressure. The system was able to subject tissue samples to physiological and pathological pressure conditions. This device can be used to increase our understanding of how heart valves respond to changes in the local mechanical environment.

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

A cyclic pressure bioreactor capable of subjecting heart valve tissue to physiological and pathological pressure conditions has been designed. A LabVIEW program allows users to control pressure magnitude, amplitude and frequency. This device can be used to study the mechanobiology of heart valve tissue or isolated cells.

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic ...

Analysis of Tubular Membrane Networks in Cardiac Myocytes from Atria and Ventricles

In cardiac myocytes a complex network of membrane tubules - the transverse-axial tubule system (TATS) - controls deep intracellular signaling functions. While the outer surface membrane and associated TATS membrane components appear to be continuous, there are substantial differences in lipid and protein content. In ventricular myocytes (VMs), certain TATS components are highly abundant contributing to rectilinear tubule networks and regular branching 3D architectures. It is thought that peripheral TATS components propagate action potentials from the cell surface to thousands of remote intracellular sarcoendoplasmic reticulum (SER) membrane contact domains, thereby activating intracellular Ca2+ release units (CRUs). In contrast to VMs, the organization and functional role of TATS membranes in atrial myocytes (AMs) is significantly different and much less understood. Taken together, quantitative structural characterization of TATS membrane networks in healthy and diseased myocytes is an essential prerequisite towards better understanding of functional plasticity and pathophysiological reorganization. Here, we present a strategic combination of protocols for direct quantitative analysis of TATS membrane networks in living VMs and AMs. For this, we accompany primary cell isolations of mouse VMs and/or AMs with critical quality control steps and direct membrane staining protocols for fluorescence imaging of TATS membranes. Using an optimized workflow for confocal or superresolution TATS image processing, binarized and skeletonized data are generated for quantitative analysis of the TATS network and its components. Unlike previously published indirect regional aggregate image analysis strategies, our protocols enable direct characterization of specific components and derive complex physiological properties of TATS membrane networks in living myocytes with high throughput and open access software tools. In summary, the combined protocol strategy can be readily applied for quantitative TATS network studies during physiological myocyte adaptation or disease changes, comparison of different cardiac or skeletal muscle cell types, phenotyping of transgenic models, and pharmacological or therapeutic interventions.

In cardiac myocytes, tubular membrane structures form intracellular networks. We describe optimized protocols for i) isolation of myocytes from mouse heart including quality control, ii) live cell staining for state-of-the-art fluorescence microscopy, and iii) direct image analysis to quantify the component complexity and the plasticity of intracellular membrane networks.

In cardiac myocytes a complex network of membrane tubules - the transverse-axial tubule system (TATS) - controls deep intracellular signaling functions. ...

Eye Irritation Test (EIT) for Hazard Identification of Eye Irritating Chemicals using Reconstructed Human Cornea-like Epithelial (RhCE) Tissue Model

To comply with the Seventh Amendment to the EU Cosmetics Directive and EU REACH legislation, validated non-animal alternative methods for reliable and accurate assessment of ocular toxicity in man are needed. To address this need, we have developed an eye irritation test (EIT) which utilizes a three dimensional reconstructed human cornea-like epithelial (RhCE) tissue model that is based on normal human cells. The EIT is able to separate ocular&#160;irritants and corrosives (GHS Categories 1 and 2 combined) and those that do not require labeling (GHS No Category). The test utilizes two separate protocols, one designed for liquid chemicals and a second, similar protocol for solid test articles. The EIT prediction model uses a single exposure period (30 min for liquids, 6 hr for solids) and a single tissue viability cut-off (60.0% as determined by the MTT assay). Based on the results for 83 chemicals (44 liquids and 39 solids) EIT achieved 95.5/68.2/ and 81.8% sensitivity/specificity and accuracy (SS&#38;A) for liquids, 100.0/68.4/ and 84.6% SS&#38;A for solids, and 97.6/68.3/ and 83.1% for overall SS&#38;A. The EIT will contribute significantly to classifying the ocular irritation potential of a wide range of liquid and solid chemicals without the use of animals to meet regulatory testing requirements. The EpiOcular EIT method was implemented in 2015 into the OECD Test Guidelines as TG 492.

Eye Irritation Test EIT for Hazard Identification of Eye Irritating Chemicals using Reconstructed Human Cornea-like Epithelial RhCE Tissue Model

We have developed an eye irritation test which utilizes a three dimensional reconstructed human cornea-like epithelial (RhCE) tissue model. The test is able to discriminate between ocular irritant and corrosive materials (GHS Categories 1 and 2 combined) and those that do not require labeling (GHS No Category).

To comply with the Seventh Amendment to the EU Cosmetics Directive and EU REACH legislation, validated non-animal alternative methods for reliable and ...

Construction of Defined Human Engineered Cardiac Tissues to Study Mechanisms of Cardiac Cell Therapy

Human cardiac tissue engineering can fundamentally impact therapeutic discovery through the development of new species-specific screening systems that replicate the biofidelity of three-dimensional native human myocardium, while also enabling a controlled level of biological complexity, and allowing non-destructive longitudinal monitoring of tissue contractile function. Initially, human engineered cardiac tissues (hECT) were created using the entire cell population obtained from directed differentiation of human pluripotent stem cells, which typically yielded less than 50% cardiomyocytes. However, to create reliable predictive models of human myocardium, and to elucidate mechanisms of heterocellular interaction, it is essential to accurately control the biological composition in engineered tissues. 
To address this limitation, we utilize live cell sorting for the cardiac surface marker SIRP&#945; and the fibroblast marker CD90 to create tissues containing a 3:1 ratio of these cell types, respectively, that are then mixed together and added to a collagen-based matrix solution. Resulting hECTs are, thus, completely defined in both their cellular and extracellular matrix composition.
Here we describe the construction of defined hECTs as a model system to understand mechanisms of cell-cell interactions in cell therapies, using an example of human bone marrow-derived mesenchymal stem cells (hMSC) that are currently being used in human clinical trials. The defined tissue composition is imperative to understand how the hMSCs may be interacting with the endogenous cardiac cell types to enhance tissue function. A bioreactor system is also described that simultaneously cultures six hECTs in parallel, permitting more efficient use of the cells after sorting.

This manuscript describes the creation of defined engineered cardiac tissues using surface marker expression and cell sorting. The defined tissues can then be used in a multi-tissue bioreactor to investigate mechanisms of cardiac cell therapy in order to provide a functional, yet controlled, model system of the human heart.

Human cardiac tissue engineering can fundamentally impact therapeutic discovery through the development of new species-specific screening systems that ...

Neonatal Cardiac Scaffolds: Novel Matrices for Regenerative Studies

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

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

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

Automated Contraction Analysis of Human Engineered Heart Tissue for Cardiac Drug Safety Screening

Cardiac tissue engineering describes techniques to constitute three dimensional force-generating engineered tissues. For the implementation of these procedures in basic research and preclinical drug development, it is important to develop protocols for automated generation and analysis under standardized conditions. Here, we present a technique to generate engineered heart tissue (EHT) from cardiomyocytes of different species (rat, mouse, human). The technique relies on the assembly of a fibrin-gel containing dissociated cardiomyocytes between elastic polydimethylsiloxane (PDMS) posts in a 24-well format. Three-dimensional, force-generating EHTs constitute within two weeks after casting. This procedure allows for the generation of several hundred EHTs per week and is technically limited only by the availability of cardiomyocytes (0.4-1.0 x 106/EHT). Evaluation of auxotonic muscle contractions is performed in a modified incubation chamber with a mechanical interlock for 24-well plates and a camera placed on top of this chamber. A software controls a camera moved on an XYZ axis system to each EHT. EHT contractions are detected by an automated figure recognition algorithm, and force is calculated based on shortening of the EHT and the elastic propensity and geometry of the PDMS posts. This procedure allows for automated analysis of high numbers of EHT under standardized and sterile conditions. The reliable detection of drug effects on cardiomyocyte contraction is crucial for cardiac drug development and safety pharmacology. We demonstrate, with the example of the hERG channel inhibitor E-4031, that the human EHT system replicates drug responses on contraction kinetics of the human heart, indicating it to be a promising tool for cardiac drug safety screening.

Here, we show the generation of human engineered heart tissue from induced pluripotent stem cells (hiPSC)-derived cardiomyocytes. We present a method to analyze contraction force and exemplary alteration of contraction pattern by the hERG channel inhibitor E-4031. This method shows high level of robustness and suitability for cardiac drug screening.

Cardiac tissue engineering describes techniques to constitute three dimensional force-generating engineered tissues. For the implementation of these ...

Light-sheet Fluorescence Microscopy for the Study of the Murine Heart

Light-sheet fluorescence microscopy has been widely used for rapid image acquisition with a high axial resolution from micrometer to millimeter scale. Traditional light-sheet techniques involve the use of a single illumination beam directed orthogonally at sample tissue. Images of large samples that are produced using a single illumination beam contain stripes or artifacts and suffer from a reduced resolution due to the scattering and absorption of light by the tissue. This study uses a dual-sided illumination beam and a simplified CLARITY optical clearing technique for the murine heart. These techniques allow for deeper imaging by removing lipids from the heart and produce a large field of imaging, greater than 10 x 10 x 10 mm3. As a result, this strategy enables us to quantify the ventricular dimensions, track the cardiac lineage, and localize the spatial distribution of cardiac-specific proteins and ion-channels from the post-natal to adult mouse hearts with sufficient contrast and resolution.

This study uses a dual-sided illumination light-sheet fluorescence microscopy (LSFM) technique combined with optical clearing to study the murine heart.

Light-sheet fluorescence microscopy has been widely used for rapid image acquisition with a high axial resolution from micrometer to millimeter scale. ...

Decellularization of Whole Human Heart Inside a Pressurized Pouch in an Inverted Orientation

The ultimate solution for patients with end-stage heart failure is organ transplant. But donor hearts are limited, immunosuppression is required, and ultimately rejection can occur. Creating a functional, autologous bio-artificial heart could solve these challenges. Biofabrication of a heart comprised of scaffold and cells is one option. A natural scaffold with tissue-specific composition as well as micro- and macro-architecture can be obtained by decellularizing hearts from humans or large animals such as pigs. Decellularization involves washing out cellular debris while preserving 3D extracellular matrix and vasculature and allowing &#34;cellularization&#34; at a later timepoint. Capitalizing on our novel finding that perfusion decellularization of complex organs is possible, we developed a more &#34;physiological&#34; method to decellularize non-transplantable human hearts by placing them inside a pressurized pouch, in an inverted orientation, under controlled pressure. The purpose of using a pressurized pouch is to create pressure gradients across the aortic valve to keep it closed and improve myocardial perfusion. Simultaneous assessment of flow dynamics and cellular debris removal during decellularization allowed us to monitor both fluid inflow and debris outflow, thereby generating a scaffold that can be used either for simple cardiac repair (e.g. as a patch or valve scaffold) or as a whole-organ scaffold.

This method enables decellularization of a complex solid organ using a simple protocol based on osmotic shock and perfusion of ionic detergent with minimal organ matrix disruption. It comprises a novel decellularization technique for human hearts inside a pressurized pouch with real-time monitoring of flow dynamics and cellular debris outflow.

The ultimate solution for patients with end-stage heart failure is organ transplant. But donor hearts are limited, immunosuppression is required, and ...