We thank the individuals who have donated their bodies for the advancement of education and research. We are grateful to the OneLegacy Foundation, which formed the basis for obtaining donor hearts for research. We are also grateful to Anthony A. Smithson and Arvin Roque-Verdeflor of the UCLA Translational Research Imaging Center (Department of Radiology) for their support in CT data acquisition. This project was supported by the UCLA Amara Yad Project. We are thankful to Drs. Kalyanam Shivkumar and Olujimi A. Ajijola for establishing and maintaining a human heart pipeline for research. We appreciate our Research Operations Manager, Amiksha S. Gandhi for her dedication to support our projects. This work was made possible by support from NIH grants OT2OD023848 &#38; P01 HL164311 and Leducq grant 23CVD04 to Kalyanam Shivkumar, the American Heart Association Career Development Award 23CDA1039446 to PH, and the UCLA Amara-Yad Project (https://www.uclahealth.org/medical-services/heart/arrhythmia/about-us/amara-yad-project). The GNEXT microPET/CT scanner used in this study was funded by an NIH Shared Instrumentation for Animal Research Grant (1 S10 OD026917-01A1).

This study used de-identified tissue samples collected from non-failing donor human hearts and was approved by the Institutional Review Board of the University of California, Los Angeles (UCLA). Samples were obtained from non-failing hearts that were rejected for transplantation. The hearts were pressure-perfused, fixed in 4% paraformaldehyde (PFA), and imaged before tissue processing per the following methods. Figure 1 summarizes the flow chart of the order of the study. The details of the reagents and the equipment used in the study are listed in the Table of Materials.
1. Micro-scale examination
<ol>
	<li>Tissue clearing using modified immunolabeling-enabled 3D imaging of solvent-cleared organs&#160;(iDISCO+) protocol.
	<ol>
		<li>Dissect 4% PFA-fixed tissue with a scalpel to fit within the 3 mm &#215; 16 mm &#215; 25 mm chamber for confocal microscopy.&#160;To image thicker tissues, additional chambers and/or spacers may be stacked on the slide.</li>
		<li>Dehydrate specimens using graded methanol (MeOH) series (20%, 40%, 60%, 80%, and 100% MeOH in deionized H2O [vol/vol]) for 1 h each at room temperature (RT) with agitation.</li>
		<li>Wash with&#160;100%&#160;MeOH for 1 h at RT and immerse in 66% dichloromethane/33% MeOH at RT with agitation overnight.</li>
		<li>The next day, wash twice in MeOH (100%) for 1 h at RT, chill at 4 &#176;C, and treat with 5% H2O2 in MeOH (vol/vol) overnight at 4&#160;&#176;C.</li>
		<li>Rehydrate with graded MeOH series (80%, 60%, 40%, and 20% MeOH) and wash in 0.01 mol/L PBS for 1 h each at RT with agitation.</li>
		<li>Wash the tissues&#160;twice in 0.01 mol/L PBS with 0.2% Triton X-100 for 1 h at RT.</li>
		<li>Prepare for immunolabeling by permeabilizing in 0.01 mol/L PBS, 20% dimethyl sulfoxide (DMSO), 0.2% Triton X-100, and 0.3 mol/L glycine for 2 days at 37&#160;&#176;C with agitation.</li>
		<li>Block in 0.01 mol/L PBS with 10% DMSO, 0.2% Triton X-100, and 5% normal donkey serum for another 2 days at 37 &#176;C with agitation.</li>
		<li>Label with a primary antibody that is compatible with MeOH conjugated to fluorophores diluted in 0.01 mol/L PBS with 10 mg/mL heparin (PTwH), 0.2% Tween-20, 5% DMSO, and 3% normal donkey serum for 3-4 days&#160;at 37&#160;&#176;C with agitation.</li>
		<li>Replenish the antibody solution and incubate for another 3-4 days at 37 &#176;C with agitation.</li>
		<li>After 1-week incubation in primary antibody solution, wash 4 to 5 times in PTwH overnight at RT.</li>
		<li>Incubate with secondary antibody conjugated to fluorophores diluted in PTwH, 3% normal donkey serum for 3 days at 37&#160;&#176;C with agitation.</li>
		<li>Replenish the secondary antibody solution incubate for another 3&#160;days at 37 &#176;C with agitation.</li>
		<li>After 6-day&#160;incubation in secondary antibody solution, wash in PTwH 4-5 times overnight at RT.</li>
		<li>Dehydrate with a graded MeOH series (20%, 40%, 60%, 80%, 100%, and 100% MeOH). The sample may be stored overnight at RT.</li>
		<li>Incubate in 66% dichloromethane/33% MeOH for 3 h at RT with agitation.</li>
		<li>Wash twice in 100% dichloromethane for 15 min at RT with agitation.</li>
		<li>Incubate and store specimens in benzyl ether. Fill the tube to minimize air from oxidizing the sample.</li>
	</ol>
	</li>
	<li>Imaging tissue-cleared specimen
	<ol>
		<li>Affix a chamber containing adhesive to a slide and apply nail polish around the perimeter of the chamber.&#160;For thicker tissues, additional chambers and/or spacers may be stacked on the slide.</li>
		<li>Place cleared tissue in the chamber, fill&#160;with benzyl ether, and apply a coverslip.</li>
		<li>Apply nail polish around the coverslip to create a&#160;seal.</li>
		<li>Obtain tilescan and Z stack images using an upright laser scanning confocal microscope with a 5x or 10x lens to image at a depth up to the working distance of the lens.</li>
		<li>Image at a resolution of 1024 x 1024 using laser lines appropriate for emission spectra of fluorophores used. Muscle autofluorescence is visible using the 488 nm laser line.</li>
		<li>Ensure that the z-axis step size is commensurate with Nyquist sampling based on the numerical aperture of the specified objective11.</li>
		<li>Stitch images and use software for 3D visualization.</li>
		<li>Create figures using maximum intensity projection (MIP) images of Z stacks for individual and merged channels (Figure 2).</li>
	</ol>
	</li>
	<li>Immunohistochemistry 
	NOTE: After the tissue is paraffin-embedded22, the following procedure is used to create slides for immunohistochemical study.
	<ol>
		<li>Preparation of refractive index matching solution (RIMS).
		<ol>
			<li>Prepare 0.1 mol/L phosphate buffer by adding 10.9 g of Na2HPO4 (anhydrous) and 3.1 g of NaH2PO4 (monohydrate) to deionized H2O to a total volume of 1 L (pH 7.4). Filter-sterilize the solution and store it at RT.</li>
			<li>Dilute the phosphate buffer to 0.02 mol/L.</li>
			<li>Dissolve Histodenz in 30 mL of 0.02 mol/L phosphate buffer by stirring the solution for 10 min with a magnetic stir bar in the final storage bottle that may be sealed to minimize evaporation and contamination.</li>
			<li>Add sodium azide to a total concentration of 0.01% (w/v) and adjust the pH to 7.5 with NaOH.</li>
			<li>Adjust the RI by varying the final concentration of Histodenz.</li>
			<li>Store the RIMS at RT for months. Discard if microbial contamination is noted. 
			NOTE: Do NOT autoclave any solutions containing sodium azide.</li>
		</ol>
		</li>
		<li>Preparation of slides for immunohistochemical study
		<ol>
			<li>Create sections of 5 &#181;m thickness with microtome. Apply tissue section to charged slides.</li>
			<li>Remove paraffin by incubating the slides in &#62;75% xylene for 10 min. Move slides to a second container with xylene for an additional 10 min.</li>
			<li>Remove xylene by immersing the slides in 100% EtOH for 10 min, then in 95% EtOH for 5 min and 70% EtOH for 5 min.</li>
			<li>Rinse the slides with deionized H2O for 5 min.</li>
			<li>Immerse slides in antigen retrieval buffer for 25 min at 90-95&#176;C.</li>
			<li>Allow the container to cool to RT for 1 h with agitation.</li>
			<li>Immerse the slides in soaking buffer (0.01 mol/L PBS + 0.4% Triton X-100) for 30 min at 4&#176;C.</li>
			<li>Encircle the tissue with a PAP pen. Add PBS to each slide and place it in a humidified chamber to prevent desiccation.</li>
			<li>Wash slides with PBS at RT with agitation for 5 min.</li>
			<li>Block with blocking buffer (0.01 mol/L PBS + 10% Donkey Serum + 0.1% TX-100) for 1 h with agitation.</li>
			<li>Incubate with a primary antibody diluted in blocking buffer overnight at 4&#176;C.</li>
			<li>The next day, allow slides to warm to RT for 15 min.</li>
			<li>Wash the slides 3 times with 0.01 mol/L PBS + 0.2% TritonX-100 for 5 min.</li>
			<li>Incubate with a secondary antibody diluted in blocking buffer for 1 h at RT with agitation.</li>
			<li>Wash the slides 3 times with 0.01 mol/L PBS + 0.2% TritonX-100 for 5 min.</li>
			<li>Wash the slides 3 times with 0.01 mol/L PBS for 5 min.</li>
			<li>Place 1 drop of RIMS with a dropper and apply a coverslip.</li>
			<li>Apply nail polish around the coverslip to create the seal.</li>
			<li>As a negative control, run a sample without the primary antibody to demonstrate the absence of specific staining.</li>
		</ol>
		</li>
		<li>Imaging immunostained slides
		<ol>
			<li>Image the slides with a laser scanning confocal microscope with 10x, 20x and 40x objective lenses.</li>
			<li>Image at a resolution of 1024 x 1024 using laser lines appropriate for the emission spectra of the secondary antibodies used.</li>
			<li>Create figures using maximum intensity projection (MIP) images of Z stacks for individual and merged channels (Figure 3).</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Meso-scale examination
<ol>
	<li>Stereoscopic dissection
	<ol>
		<li>Perform delicate dissections focusing on tiny or thin structures, such as atrioventricular node, atrioventricular node artery, and cardiac nerve plexus (submillimeter to millimeters in scale) with either a magnifying desk lamp with clamp, surgical telescopes, or stereomicroscope.</li>
	</ol>
	</li>
	<li>Micro-CT scanning 
	NOTE: CT imaging is performed after pressure perfusion and fixation and at any stages of dissection using a micro-positron emission tomography (PET)/CT scanner (Figure 4).
	<ol>
		<li>Warm up the CT X-ray source for 25 min before sample imaging.</li>
		<li>Place the heart sample on the scanner bed.</li>
		<li>Move the scanner bed to a horizontal position of 544 mm and a vertical position of 14 mm to center the heart in the CT field of view (FOV).</li>
		<li>Acquire CT image at 80 kVp, 150 &#181;A, with 720 projections during 1 min scan time at a spatial resolution of 200 &#181;m.</li>
		<li>Reconstruct the CT data with a 12 cm x 12 cm x 10&#160;cm field of view and a matrix of 600 x 600 x 500 voxels, and save as a DICOM file.</li>
	</ol>
	</li>
</ol>
3. Macro-scale examination
<ol>
	<li>Pressure perfusion and fixation 
	NOTE: The authors modify previously described pressure perfusion and fixation techniques and apply them to the non-failing human hearts rejected for transplantation14,19,20,21.
	<ol>
		<li>Use high-flow pumps for perfusion fixation. Use either 100% ethanol14, 4% PFA, or 10% formalin for the fixative. 
		NOTE: The heart is recovered with the ascending aorta, pulmonary trunk, and both vena cavae, and pulmonary veins&#160;resected as distally as possible and delivered in&#160;University of Wisconsin solution23.</li>
		<li>Use two 20-24 Fr surgical cannulas for right- and left-heart perfusion. For right-heart perfusion, cannulate the superior vena cava, and place a vent at the pulmonary trunk or pulmonary artery with half-cut 12-30 mL sized plastic syringes with Luer-Lock tips attached to three-way stopcocks.</li>
		<li>Occlude the inferior vena cava and the other pulmonary artery with twine after putting an appropriately sized locked half-cut plastic syringe or 1.5-5.0 mL centrifuge tube.
		<ol>
			<li>For antegrade perfusion of the left heart, cannulate one of the pulmonary veins and place a vent at the distal cut end of the aorta with the half-cut 12-30 mL sized plastic syringes with Luer-Lock tips attached to three-way stopcocks.</li>
			<li>For retrograde perfusion of the left heart, cannulate one of the aortic arch branches and place a vent at another branch of the aortic arch. Place the tips of the cannulas in both ventricles.</li>
		</ol>
		</li>
		<li>Occlude other vessel orifices with twine after inserting an appropriately sized locked half-cut plastic syringe or 1.5-5.0 mL microcentrifuge tube. Use a thin gauze to cover the inserting portion of the syringes/ tubes/ cannulas to prevent leakage and slipping. Fix large leaks using suture, banding, or clumping. Small leaks are permissible.</li>
		<li>Suspend the heart in a plastic container.</li>
		<li>Connect 22-24 Fr soft plastic tubing to each cannula and insert the other end of the tube into the container filled with fixative.</li>
		<li>Circulate fixative through the right- and left-heart circuits using a high-flow pump set at approximately 100-300 mL/min for the right heart and 200-400 mL/min for the left heart to achieve approximately 20 mmHg in the right ventricle and 80 mmHg in the left ventricle, respectively.</li>
		<li>Maintain perfusion at 4 &#176;C for 24 h.</li>
		<li>Wash the heart with 0.01 mol/L PBS for 30 min with agitation four times.</li>
		<li>Store heart in 0.01 mol/L PBS/0.02% sodium azide at 4 &#176;C. 
		NOTE: The pressure-perfusion fixation is only effective for a fresh heart, not for a heart recovered from an embalmed cadaver.</li>
	</ol>
	</li>
	<li>Gross dissection
	<ol>
		<li>Perform progressive dissection with photographic recordings at each stage of dissection.</li>
		<li>To maintain clinical relevance, pay special attention to avoid distorting/deforming any structures to maintain the physiological morphology of the heart.</li>
		<li>Image target structures using clinically relevant orientation, such as right anterior oblique orientation.</li>
	</ol>
	</li>
	<li>Photography
	<ol>
		<li>Place the pressure-perfused and fixed heart on a tripod with a platform mounted with multiple prongs and the ability to rotate 360o.</li>
		<li>Photograph the heart using a digital single-lens reflex camera&#160;(Figure 5)24 while using multiple light-emitting diode light panels set on the C-Stands and wide black duvetyn background cloth.</li>
		<li>Photograph using the lens with a long focal length (200 mm) for a working distance of 4-6 ft to minimize distortion of the subject14.</li>
	</ol>
	</li>
	<li>Anaglyphs
	<ol>
		<li>To display anaglyphic images, reconstruct a pair of photographs or volume-rendered images from CT datasets with a 10&#176; difference in the rotation angle on the horizontal plane.</li>
		<li>Convert a set of these two-dimensional (2D) images, referred to as a stereogram, into anaglyphs using freeware16.</li>
		<li>To view an anaglyph, use red/cyan glasses.</li>
	</ol>
	</li>
	<li>Photogrammetry 
	NOTE: Photogrammetry is the applied science of generating a three-dimensional surface-rendered reconstruction from multiple two-dimensional photographs taken at varying angles17.
	<ol>
		<li>Suspend the sample on C-Stands or place it on the rotation table to obtain hundreds of multi-directional photographs with a smartphone.</li>
		<li>Generate the 3D model in FBX format using commercially available software.</li>
	</ol>
	</li>
	<li>CT scanning 
	NOTE: CT scanning may be performed after pressure perfusion and fixation and at any stage of dissection.
	<ol>
		<li>Suspend the heart sample from a bar placed across the top of the container. To prevent the heart from swinging during the scan, support the base of the heart with plastic prongs fixed at the bottom of the container. Thus, the air will serve as a negative contrast.</li>
		<li>Perform the CT scan using a commercially available multi-detector row CT scanner with the following parameters: tube voltage of 120 kV, tube current of 800-900 mA, and a gantry rotation of 280 ms. The dose length product is generally 500-1200 mGy.cm.</li>
		<li>Reconstruct axial image data using the following parameters: a section thickness, 0.6 mm; an incremental interval, 0.3 mm; a field of view, as small as possible (generally 100-200 mm); and a matrix, 512 &#215; 512.</li>
	</ol>
	</li>
	<li>Virtual dissection
	<ol>
		<li>Analyze the CT scan images using commercially available software to generate virtual dissection images. 
		NOTE: Virtual dissection is a modification of the volume-rendering process wherein the focus is shifted to the walls of the cardiac chambers and vessels18. In this process, manual thresholding virtually removes the enhanced chamber from the original datasets.</li>
		<li>Visualize the non-enhanced walls, septa, and valves with virtual dissection to produce images similar to gross dissection. Unlike gross dissection of the heart specimens, the cut planes during virtual dissection are practically unlimited. Almost any view can be recreated to visualize the structures of interest as needed.</li>
	</ol>
	</li>
	<li>3D printing
	<ol>
		<li>Open the compatible file of the heart sample in 3D printer software.</li>
		<li>Use the profile of 0.10 mm Fast DETAIL for the print Settings in the 3D printer and reduce the print speed to 20 mm/s. Enable Generate support material.</li>
		<li>Use the profile of TPU&#8203;&#160;filament for &#34;Filament Settings&#34; in the 3D printer.</li>
		<li>Use the profile of the Original Prusa MK4 Input Shaper 0.4 nozzle for &#34;Printer Settings&#34; in the printer.</li>
		<li>After slicing is completed, save the BGCODE file in a USB flash drive for 3D printing.</li>
		<li>Use the 1.75 mm TPU filament for 3D printing the human heart sample. Before 3D printing, dry the TPU filament for 6 h using a&#160;filament dryer.</li>
		<li>To reduce filament tension during 3D printing, place the filament spool on a spool holder with a built-in bearing to facilitate filament spool rotation. Perform 3D printing using a commercially available 3D printer with a textured powder-coated steel sheet.</li>
		<li>Carefully remove support materials when 3D printing is complete.</li>
	</ol>
	</li>
</ol>

Comprehensive 3D Analysis of Human Hearts from Micro to Macro Scales

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K., Izawa, Y., Nishii, T., Mori, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Virtual+dissection:+Emerging+as+the+gold+standard+of+analyzing+living+heart+anatomy.">Virtual dissection: Emerging as the gold standard of analyzing living heart anatomy.</a> J Cardiovasc Dev Dis. 7 (3), 30 (2020).</li><li>Thomas, A. C., Davies, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+demonstration+of+cardiac+pathology+using+perfusion-fixation.">The demonstration of cardiac pathology using perfusion-fixation.</a> Histopathology. 9 (1), 5-19 (1985).</li><li>Glagov, S., Eckner, F. A., Lev, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Controlled+pressure+fixation+apparatus+for+hearts.">Controlled pressure fixation apparatus for hearts.</a> Arch Pathol. 76, 640-646 (1963).</li><li>Iaizzo, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+visible+heart(r)+project+and+free-access+website+'atlas+of+human+cardiac+anatomy.">The visible heart(r) project and free-access website 'atlas of human cardiac anatomy.</a> Europace. 18, 163-172 (2016).</li><li>Yang, Y., Huang, H., Li, L., Yang, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multiplex+immunohistochemistry+staining+for+paraffin-embedded+lung+cancer+tissue.">Multiplex immunohistochemistry staining for paraffin-embedded lung cancer tissue.</a> J Vis Exp. (201), e65850 (2023).</li><li>Tripathy, S., Das, S. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Strategies+for+organ+preservation:+Current+prospective+and+challenges.">Strategies for organ preservation: Current prospective and challenges.</a> Cell Biol Int. 47 (3), 520-538 (2023).</li><li>Mori, S., Shivkumar, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Atlas+of+cardiac+anatomy+(anatomical+basis+of+cardiac+interventions.+Vol.+1).">Atlas of cardiac anatomy (anatomical basis of cardiac interventions. Vol. 1).</a> Cardiotext. , (2022).</li><li>Crosado, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phenoxyethanol-based+embalming+for+anatomy+teaching:+An+18+years'+experience+with+crosado+embalming+at+the+university+of+otago+in+new+zealand.">Phenoxyethanol-based embalming for anatomy teaching: An 18 years' experience with crosado embalming at the university of otago in new zealand.</a> Anat Sci Educ. 13 (6), 778-793 (2020).</li><li>Titmus, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+workflow+for+the+creation+of+photorealistic+3d+cadaveric+models+using+photogrammetry.">A workflow for the creation of photorealistic 3d cadaveric models using photogrammetry.</a> J Anat. 243 (2), 319-333 (2023).</li><li>Silva, J. N. A., Southworth, M., Raptis, C., Silva, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Emerging+applications+of+virtual+reality+in+cardiovascular+medicine.">Emerging applications of virtual reality in cardiovascular medicine.</a> JACC Basic Transl Sci. 3 (3), 420-430 (2018).</li><li>Maresky, H. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Virtual+reality+and+cardiac+anatomy:+Exploring+immersive+three-dimensional+cardiac+imaging,+a+pilot+study+in+undergraduate+medical+anatomy+education.">Virtual reality and cardiac anatomy: Exploring immersive three-dimensional cardiac imaging, a pilot study in undergraduate medical anatomy education.</a> Clin Anat. 32 (2), 238-243 (2019).</li><li>Mori, S., Shivkumar, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Real+three-dimensional+cardiac+imaging+using+leading-edge+holographic+display.">Real three-dimensional cardiac imaging using leading-edge holographic display.</a> Clin Anat. 34 (6), 966-968 (2021).</li></ol>

The present study demonstrates&#160;the comprehensive pipeline to analyze samples obtained from whole human hearts. Representative results show&#160;micro- to macroscale anatomical examinations carried out routinely for a single heart. As a&#160;human heart sample is extremely precious, a multi-scale approach is ideal and effective so as not to waste any parts of the sample by applying multiple&#160;protocols for various purposes, including discovery in scientific research, improvement of clinical procedures, and anatomi...

As function follows form, understanding the architecture of the heart is fundamental for appreciation of its physiology. Although numerous investigations have revealed cardiac anatomy from micro- to macroscales1,2,3, multiple questions remain unresolved, especially those related to human cardiac anatomy. This is in part because basic studies focusing on functional anatomy generally utilized animal hearts4,5,6, which are often distinct from human hearts1,7,8. Furthermore, each individual study, even those using human heart samples, tends to focus on very specific structures, which renders it difficult to apply the findings in the context of the whole heart. This is even more so if the focused structures are at micro- or mesoscales, such as the perinexus9 and ganglionated plexuses10.
In this context, systemic structural study of the human heart rejected for transplantation provides a unique and rare opportunity to obtain a comprehensive atlas of cardiac structures in focus across microscopic and macroscopic scales11. Microscopic examination protocols include tissue clearing (modified immunolabeling-enabled three-dimensional (3D) imaging of solvent-cleared organs, iDISCO+)12,13, and immunohistochemical staining. Mesoscopic examination protocols include stereoscopic dissection, macro photography, and micro-computed tomographic (CT) scanning. Macroscopic examination protocols include gross dissection14, photography (including anaglyphs and photogrammetry)15,16,17, CT, virtual dissection18, and 3D printing of the physically or virtually dissected or whole heart17. In preparation for&#160;macroscopic examination, pressure-perfusion fixation is&#160;performed to maintain the 3D architecture and physiologically relevant morphology of the heart14,19,20,21.&#160;The combined application of these techniques is unique and crucial to correlate distinct anatomic features in the context of the 3D architecture of the human heart.
As the opportunity to obtain a non-pathological human heart sample is extremely limited, a multi-scale approach described herein maximizes the use of the sample. By applying various procedures described below, representative results will illustrate to the reader how the findings can be utilized for multiple purposes, including discovery in scientific research11 (comprehensive analyses of cardiac innervation, distribution of ganglionated plexuses), improvement of clinical procedures (simulation for surgical and interventional approaches), and anatomical education (real 3D demonstration of cardiac anatomy).

<table><tbody><tr><td>1x Phosphate buffered saline</td><td>Sigma-Aldrich</td><td>P3813</td><td></td></tr><tr><td>3D Viewer</td><td>Microsoft</td><td></td><td></td></tr><tr><td>647 AffiniPure Donkey Anti-Rabbit IgG</td><td>Jackson ImmunoResearch Laboratories</td><td>711-605-152</td><td></td></tr><tr><td>647 AffiniPure Donkey Anti-Sheep IgG</td><td>Jackson ImmunoResearch Laboratories</td><td>713-605-147</td><td></td></tr><tr><td>AF Micro-NIKKOR 200 mm f/4D IF-ED lens</td><td>Nikon</td><td></td><td></td></tr><tr><td>Anti-Actin, &amp;alpha;-Smooth Muscle - Cy3 antibody</td><td>Sigma-Aldrich</td><td>C6198</td><td></td></tr><tr><td>Antigen Retrieval Buffer (100x EDTA Buffer, pH 8.0)</td><td>Abcam</td><td>ab93680</td><td></td></tr><tr><td>Anti-PGP9.5 (protein gene product 9.5)</td><td>Abcam</td><td>ab108986</td><td></td></tr><tr><td>Anti-TH (tyrosine hydrox ylase)</td><td>Abcam</td><td>ab1542</td><td></td></tr><tr><td>Anti-VAChT (vesicular acetylcholine transporter)</td><td>Synaptic Systems</td><td>139 103</td><td></td></tr><tr><td>Benzyl ether</td><td>Sigma-Aldrich</td><td>108014</td><td></td></tr><tr><td>Bovine serum albumin</td><td>Sigma-Aldrich</td><td>A4503-10G</td><td></td></tr><tr><td>Cheetah 3D printer filament (95A), 1.75 mm</td><td>NinjaTek</td><td></td><td></td></tr><tr><td>Coverslip, 22 mm x 30mm, No. 1.5</td><td>VWR</td><td>48393 151</td><td></td></tr><tr><td>Cy3 AffiniPure Donkey Anti-Rabbit IgG</td><td>Jackson ImmunoResearch Laboratories</td><td>711-165-152</td><td></td></tr><tr><td>Dichloromethane</td><td>Sigma-Aldrich</td><td>270997-100ML</td><td></td></tr><tr><td>Dimethyl sulfoxide</td><td>Sigma-Aldrich</td><td>D8418-500ML</td><td></td></tr><tr><td>Ethanol, 100%</td><td>Decon laboratories</td><td>2701</td><td></td></tr><tr><td>Glycine</td><td>Sigma-Aldrich</td><td>G7126-500G</td><td></td></tr><tr><td>GNEXT PET/CT</td><td>SOFIE Biosciences</td><td></td><td></td></tr><tr><td>Heparin sodium salt from porcine intestinal mucosa</td><td>Sigma-Aldrich</td><td>H3149-50KU</td><td></td></tr><tr><td>Histodenz</td><td>Sigma-Aldrich</td><td>D2158-100G</td><td></td></tr><tr><td>Hydrogen peroxide solution</td><td>Sigma-Aldrich</td><td>H1009-500ML</td><td></td></tr><tr><td>Imaging software</td><td>Zeiss</td><td>ZEN (black edition)</td><td></td></tr><tr><td>Imaging software</td><td>Oxford Instruments</td><td>Imaris 10</td><td></td></tr><tr><td>iSpacer</td><td>Sunjin Labs</td><td>iSpacer 3mm</td><td></td></tr><tr><td>KIRI Engine</td><td>KIRI Innovation</td><td></td><td></td></tr><tr><td>Laser scanning confocal microscope</td><td>Zeiss</td><td>LSM 880</td><td></td></tr><tr><td>LEAD-2 - Vertical &amp;amp; Multi-channels Peristaltic Pump</td><td>LONGER</td><td></td><td></td></tr><tr><td>Lightview XL&amp;nbsp;</td><td>Brightech</td><td></td><td></td></tr><tr><td>Methanol (Certified ACS)</td><td>Fischer Scientific</td><td>A412-4</td><td></td></tr><tr><td>Nikon D850</td><td>Nikon</td><td></td><td></td></tr><tr><td>NinjaTek NinjaFlex TPU @MK4</td><td>NinjaTek</td><td></td><td></td></tr><tr><td>Normal donkey serum</td><td>Jackson ImmunoResearch Laboratories</td><td>017-000-121</td><td></td></tr><tr><td>Original Prusa MK4 3D printer</td><td>Prusa Research</td><td></td><td></td></tr><tr><td>PAP pen</td><td>Abcam</td><td>ab2601</td><td></td></tr><tr><td>Paraformaldehyde, 32%</td><td>Electron Microscopy Sciences</td><td>15714-S</td><td></td></tr><tr><td>Polycam</td><td>Polycam</td><td></td><td></td></tr><tr><td>Primary antibody</td><td></td><td></td><td></td></tr><tr><td>PrusaSlicer 2.7.1</td><td>Prusa Research</td><td></td><td></td></tr><tr><td>SARA-Engine</td><td>pita4 mobile LLC</td><td></td><td></td></tr><tr><td>Scaniverse</td><td>Niantic</td><td></td><td></td></tr><tr><td>Secondary antibody</td><td></td><td></td><td></td></tr><tr><td>SlowFade Gold Antiface Mountant</td><td>Invitrogen</td><td>S36936</td><td></td></tr><tr><td>Sodium azide, 5% (w/v)</td><td>Ricca Chemical Company</td><td>7144.8-32</td><td></td></tr><tr><td>SOMATOM Definition AS</td><td>Siemens Healthcare</td><td></td><td></td></tr><tr><td>Standard Field Surgi-Spec Telescopes,&amp;nbsp;</td><td>Designs for Vision</td><td></td><td></td></tr><tr><td>Stereomicroscope System SZ61</td><td>OLYMPUS</td><td></td><td></td></tr><tr><td>StereoPhoto Maker</td><td>Free ware developed by Masuji Suto</td><td></td><td></td></tr><tr><td>Superfrost Plus Microscope Slides, Precleaned</td><td>Fisher Scientific</td><td>12-550-15</td><td></td></tr><tr><td>Triton X-100</td><td>Sigma-Aldrich</td><td>T8787-50ML</td><td></td></tr><tr><td>Tween-20</td><td>Sigma-Aldrich</td><td>P9416-100ML</td><td></td></tr><tr><td>Xylene</td><td>Sigma-Aldrich</td><td>534056-4L</td><td></td></tr><tr><td>Ziostation2</td><td>Ziosoft, AMIN</td><td></td><td></td></tr></tbody></table>

pipeline for multi-scale three-dimensional anatomic study of the human heart

Detailed study of non-failing human hearts rejected for transplantation provides a unique opportunity to perform structural analyses across microscopic and macroscopic scales. These techniques include tissue clearing (modified immunolabeling-enabled three-dimensional (3D) imaging of solvent-cleared organs) and immunohistochemical staining. Mesoscopic examination procedures include stereoscopic dissection and micro-computed tomographic (CT) scanning. Macroscopic examination procedures include gross dissection, photography (including anaglyphs and photogrammetry), CT, and 3D printing of the physically or virtually dissected or whole heart. Before macroscopic examination, pressure-perfusion fixation may be performed to maintain the 3D architecture and physiologically relevant morphology of the heart. The application of these techniques in combination to study the human heart is unique and crucial in understanding the relationship between distinct anatomic features such as coronary vasculature and myocardial innervation in the context of the 3D architecture of the heart. This protocol describes the methodologies in detail and includes representative results&#160;to illustrate progress in the research of human cardiac anatomy.

Microscale examinations 
Applying tissue clearing allows imaging of larger volumes of tissue in 3D using confocal microscopy. In the heart, ganglia containing cardiac neurons and the neural patterning of myocardial innervation can be visualized (Figure 2). Figure 3 shows a confocal image of the human left ventricle myocardium immunostained for nerves and smooth muscle cells. Blood vessels are noted to traverse the myocardium, and numerous n...

Author Spotlight: Advancing Human Cardiac Anatomy Through Multi-Scale Analysis of Hearts

This protocol presents a comprehensive pipeline to analyze samples obtained from human hearts that span the microscopic and macroscopic scales.

Pipeline for Multi-Scale Three-Dimensional Anatomic Study of the Human Heart

Detailed study of non-failing human hearts rejected for transplantation provides a unique opportunity to perform structural analyses across microscopic ...

pipeline-for-multi-scale-three-dimensional-anatomic-study-human

Research

JoVE Journal

Biology

379 Views.  David Geffen School of Medicine at UCLA. Het kennen van de anatomie van het hart is van fundamenteel belang voor het begrijpen van de hartfysiologie. Studies van functionele anatomie hebben zich uitstrekken van micro- tot macroschalen, maar hebben grotendeels dierenharten gebruikt. Systematische studie van menselijke donorharten die zijn afgewezen voor transplantatie biedt een unieke kans om een uitgebreide atlas te ontwikkelen die zich uitstrekt van microscopische tot macroscopische schalen....

Video: Auteur in de schijnwerpers: het bevorderen van de menselijke cardiale anatomie door middel van multischaalanalyse van harten

Anatomical Reconstructions of the Human Cardiac Venous System using Contrast-computed Tomography of Perfusion-fixed Specimens

A detailed understanding of the complexity and relative variability within the human cardiac venous system is crucial for the development of cardiac devices that require access to these vessels. For example, cardiac venous anatomy is known to be one of the key limitations for the proper delivery of cardiac resynchronization therapy (CRT)1 Therefore, the development of a database of anatomical parameters for human cardiac venous systems can aid in the design of CRT delivery devices to overcome such a limitation. In this research project, the anatomical parameters were obtained from 3D reconstructions of the venous system using contrast-computed tomography (CT) imaging and modeling software (Materialise, Leuven, Belgium). The following parameters were assessed for each vein: arc length, tortuousity, branching angle, distance to the coronary sinus ostium, and vessel diameter.
	CRT is a potential treatment for patients with electromechanical dyssynchrony. Approximately 10-20% of heart failure patients may benefit from CRT2. Electromechanical dyssynchrony implies that parts of the myocardium activate and contract earlier or later than the normal conduction pathway of the heart. In CRT, dyssynchronous areas of the myocardium are treated with electrical stimulation. CRT pacing typically involves pacing leads that stimulate the right atrium (RA), right ventricle (RV), and left ventricle (LV) to produce more resynchronized rhythms. The LV lead is typically implanted within a cardiac vein, with the aim to overlay it within the site of latest myocardial activation. 
	We believe that the models obtained and the analyses thereof will promote the anatomical education for patients, students, clinicians, and medical device designers. The methodologies employed here can also be utilized to study other anatomical features of our human heart specimens, such as the coronary arteries. To further encourage the educational value of this research, we have shared the venous models on our free access website: <a href="http://www.vhlab.umn.edu/atlas" target="_blank">www.vhlab.umn.edu/atlas</a>.

The objective of this research is to recreate and then access the anatomy of the human cardiac venous system using 3D reconstructions generated from contrast-computed tomography scans.

A detailed  understanding of the complexity and relative variability within the human  cardiac venous system is crucial for the development of cardiac ...

Bioengineering

Capturing the Cardiac Injury Response of Targeted Cell Populations via Cleared Heart Three-Dimensional Imaging

Cardiovascular disease outranks all other causes of death and is responsible for a staggering 31% of mortalities worldwide. This disease manifests in cardiac injury, primarily in the form of an acute myocardial infarction. With little resilience following injury, the once healthy cardiac tissue will be replaced by fibrous, non-contractile scar tissue and often be a prelude to heart failure. To identify novel treatment options in regenerative medicine, research has focused on vertebrates with innate regenerative capabilities. One such model organism is the neonatal mouse, which responds to cardiac injury with robust myocardial regeneration. In order to induce an injury in the neonatal mouse that is clinically relevant, we have developed a surgery to occlude the left anterior descending artery (LAD), mirroring a myocardial infarction triggered by atherosclerosis in the human heart. When matched with the technology to track changes both within cardiomyocytes and non-myocyte populations, this model provides us with a platform to identify the mechanisms that guide heart regeneration. Gaining insight into changes in cardiac cell populations following injury once relied heavily on methods such as tissue sectioning and histological examination, which are limited to two-dimensional analysis and often damage the tissue in the process. Moreover, these methods lack the ability to trace changes in cell lineages, instead providing merely a snapshot of the injury response. Here, we describe how technologically advanced methods in lineage tracing models, whole organ clearing, and three-dimensional (3D) whole-mount microscopy can be used to elucidate mechanisms of cardiac repair. With our protocol for neonatal mouse myocardial infarction surgery, tissue clearing, and 3D whole organ imaging, the complex pathways that induce cardiomyocyte proliferation can be unraveled, revealing novel therapeutic targets for cardiac regeneration.

Cardiomyocyte proliferation following injury is a dynamic process that requires a symphony of extracellular cues from non-myocyte cell populations. Utilizing lineage tracing, passive CLARITY, and three-dimensional whole-mount confocal microscopy techniques, we can analyze the influence of a variety of cell types on cardiac repair and regeneration.

Cardiovascular disease outranks all other causes of death and is responsible for a staggering 31% of mortalities worldwide. This disease manifests in ...

Developmental Biology

3D Modeling of the Lateral Ventricles and Histological Characterization of Periventricular Tissue in Humans and Mouse

The ventricular system carries and circulates cerebral spinal fluid (CSF) and facilitates clearance of solutes and toxins from the brain. The functional units of the ventricles are ciliated epithelial cells termed ependymal cells, which line the ventricles and through ciliary action are capable of generating laminar flow of CSF at the ventricle surface. This monolayer of ependymal cells also provides barrier and filtration functions that promote exchange between brain interstitial fluids (ISF) and circulating CSF. Biochemical changes in the brain are thereby reflected in the composition of the CSF and destruction of the ependyma can disrupt the delicate balance of CSF and ISF exchange. In humans there is a strong correlation between lateral ventricle expansion and aging. Age-associated ventriculomegaly can occur even in the absence of dementia or obstruction of CSF flow. The exact cause and progression of ventriculomegaly is often unknown; however, enlarged ventricles can show regional and, often, extensive loss of ependymal cell coverage with ventricle surface astrogliosis and associated periventricular edema replacing the functional ependymal cell monolayer. Using MRI scans together with postmortem human brain tissue, we describe how to prepare, image and compile 3D renderings of lateral ventricle volumes, calculate lateral ventricle volumes, and characterize periventricular tissue through immunohistochemical analysis of en face lateral ventricle wall tissue preparations. Corresponding analyses of mouse brain tissue are also presented supporting the use of mouse models as a means to evaluate changes to the lateral ventricles and periventricular tissue found in human aging and disease. Together, these protocols allow investigations into the cause and effect of ventriculomegaly and highlight techniques to study ventricular system health and its important barrier and filtration functions within the brain.

Using MRI scans (human), 3D imaging software, and immunohistological analysis, we document changes to the brain&#8217;s lateral ventricles. Longitudinal 3D mapping of lateral ventricle volume changes and characterization of periventricular cellular changes that occur in the human brain due to aging or disease are then modeled in mice.

The ventricular system carries and circulates cerebral spinal fluid (CSF) and facilitates clearance of solutes and toxins from the brain. The functional ...

Neuroscience

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 ...

Medicine

Developing 3D Organized Human Cardiac Tissue within a Microfluidic Platform

The leading cause of death worldwide persists as cardiovascular disease (CVD). However, modeling the physiological and biological complexity of the heart muscle, the myocardium, is notoriously difficult to accomplish in vitro. Mainly, obstacles lie in the need for human cardiomyocytes (CMs) that are either adult or exhibit adult-like phenotypes and can successfully replicate the myocardium&#39;s cellular complexity and intricate 3D architecture. Unfortunately, due to ethical concerns and lack of available primary patient-derived human cardiac tissue, combined with the minimal proliferation of CMs, the sourcing of viable human CMs has been a limiting step for cardiac tissue engineering. To this end, most research has transitioned toward cardiac differentiation of human induced pluripotent stem cells (hiPSCs) as the primary source of human CMs, resulting in the wide incorporation of hiPSC-CMs within in vitro assays for cardiac tissue modeling.
Here in this work, we demonstrate a protocol for developing a 3D mature stem cell-derived human cardiac tissue within a microfluidic device. We specifically explain and visually demonstrate the production of a 3D in vitro anisotropic cardiac tissue-on-a-chip model from hiPSC-derived CMs. We primarily describe a purification protocol to select for CMs, the co-culture of cells with a defined ratio via mixing CMs with human CFs (hCFs), and suspension of this co-culture within the collagen-based hydrogel. We further demonstrate the injection of the cell-laden hydrogel within our well-defined microfluidic device, embedded with staggered elliptical microposts that serve as surface topography to induce a high degree of alignment of the surrounding cells and the hydrogel matrix, mimicking the architecture of the native myocardium. We envision that the proposed 3D anisotropic cardiac tissue-on-chip model is suitable for fundamental biology studies, disease modeling, and, through its use as a screening tool, pharmaceutical testing.

The goal of this protocol is to explain and demonstrate the development of a three-dimensional (3D) microfluidic model of highly aligned human cardiac tissue, composed of stem cell-derived cardiomyocytes co-cultured with cardiac fibroblasts (CFs) within a biomimetic, collagen-based hydrogel, for applications in cardiac tissue engineering, drug screening, and disease modeling.

The leading cause of death worldwide persists as cardiovascular disease (CVD). However, modeling the physiological and biological complexity of the heart ...

Simultaneous Brightfield, Fluorescence, and Optical Coherence Tomographic Imaging of Contracting Cardiac Trabeculae Ex Vivo

In cardiac muscle, intracellular Ca2+ transients activate contractile myofilaments, causing contraction, macroscopic shortening, and geometric&#160;deformation. Our understanding of the internal relationships between these events has been limited because we can neither &#39;see&#39; inside the muscle nor precisely track the spatio-temporal nature of excitation-contraction dynamics. To resolve these problems, we have constructed a device that combines a suite of imaging modalities. Specifically, it integrates a brightfield microscope to measure local changes of sarcomere length and tissue strain, a fluorescence microscope to visualize the Ca2+ transient, and an optical coherence tomograph to capture the tissue&#39;s geometric&#160;changes throughout the time-course of a cardiac cycle. We present here the imaging infrastructure and associated data collection framework. Data are collected from isolated rod-like tissue structures known as trabeculae carneae. In our instrument, a pair of position-controlled platinum hooks hold each end of an ex vivo muscle sample while it is continuously superfused with nutrient-rich saline solution. The hooks are under independent control, permitting real-time control of muscle length and force. Lengthwise translation enables the piecewise scanning of the sample, overcoming limitations associated with the relative size of the microscope imaging window (540 &#181;m by 540 &#181;m) and the length of a typical trabecula (&#62;2000 &#181;m). Platinum electrodes at either end of the muscle chamber stimulate the trabecula at a user-defined rate. We exploit the stimulation signal as a trigger for synchronizing the data from each imaging window to reconstruct the entire sample twitching under steady-state conditions. Applying image-processing techniques to these brightfield imaging data provides tissue displacement and sarcomere length maps. Such a collection of data, when incorporated into an experiment-modeling pipeline, will provide a deeper understanding of muscle contractile homogeneity and heterogeneity in physiology and pathophysiology.

Simultaneous Brightfield, Fluorescence, and Optical Coherence Tomographic Imaging of Contracting Cardiac Trabeculae Ex Vivo

This protocol presents a collection of sarcomere, calcium, and macroscopic geometric&#160;data from an actively contracting cardiac trabecula ex vivo. These simultaneous measurements are made possible by the integration of three imaging modalities.

In cardiac muscle, intracellular Ca2+ transients activate contractile myofilaments, causing contraction, macroscopic shortening, and ...

2D and 3D Echocardiography in the Axolotl (Ambystoma Mexicanum)

Cardiac malfunction as a result of ischemic heart disease is a major challenge, and regenerative therapies to the heart are in high demand. A few model species such as zebrafish and salamanders that are capable of intrinsic heart regeneration hold promise for future regenerative therapies for human patients. To evaluate the outcome of cardioregenerative experiments it is imperative that heart function can be monitored. The axolotl salamander (A. mexicanum) represents a well-established model species in regenerative biology attaining sizes that allows for evaluation of cardiac function. The purpose of this protocol is to establish methods to reproducibly measure cardiac function in the axolotl using echocardiography. The application of different anesthetics (benzocaine, MS-222, and propofol) is demonstrated, and the acquisition of two-dimensional (2D) echocardiographic data in both anesthetized and unanesthetized axolotls is described. 2D echocardiography of the three-dimensional (3D) heart can suffer from imprecision and subjectivity of measurements, and to alleviate this phenomenon a solid method, namely intra/inter-operator/observer analysis, to measure and minimize this bias is demonstrated. Finally, a method to acquire 3D echocardiographic data of the beating axolotl heart at a very high spatiotemporal resolution and with pronounced blood-to-tissue contrast is described. Overall, this protocol should provide the necessary methods to evaluate cardiac function and model anatomy, and flow dynamics in the axolotl using ultrasound imaging with applications in both regenerative biology and general physiological experiments.

2D and 3D Echocardiography in the Axolotl Ambystoma Mexicanum

Here we present echocardiography protocols for two-dimensional and three-dimensional image acquisition of the beating heart of the axolotl salamander (Ambystoma mexicanum), a model species in heart regeneration. These methods allow for longitudinal evaluation of cardiac function at a high spatiotemporal resolution.

Cardiac malfunction as a result of ischemic heart disease is a major challenge, and regenerative therapies to the heart are in high demand. A few model ...

Light-Sheet Microscopy and Tissue Clearing for Comprehensive Cardiac Imaging

Light-Sheet Imaging to Reveal Cardiac Structure in Rodent Hearts

Light-sheet microscopy (LSM) plays a pivotal role in comprehending the intricate three-dimensional (3D) structure of the heart, providing crucial insights into fundamental cardiac physiology and pathologic responses. We hereby delve into the development and implementation of the LSM technique to elucidate the micro-architecture of the heart in mouse models. The methodology integrates a customized LSM system with tissue clearing techniques, mitigating light scattering within cardiac tissues for volumetric imaging. The combination of conventional LSM with image stitching and multiview deconvolution approaches allows for the capture of the entire heart. To address the inherent trade-off between axial resolution and field of view (FOV), we further introduce an axially swept light-sheet microscopy (ASLM) method to minimize out-of-focus light and uniformly illuminate the heart across the propagation direction. In the meanwhile, tissue clearing methods such as iDISCO enhance light penetration, facilitating the visualization of deep structures and ensuring a comprehensive examination of the myocardium throughout the entire heart. The combination of the proposed LSM and tissue clearing methods presents a promising platform for researchers in resolving cardiac structures in rodent hearts, holding great potential for the understanding of cardiac morphogenesis and remodeling.

Author Spotlight: Customized Light-Sheet Imaging for Investigating Myocardial Structures in Rodent Hearts

The protocol utilizes advanced light-sheet microscopy along with adapted tissue clearing methods to investigate intricate cardiac structures in rodent hearts, holding great potential for the understanding of cardiac morphogenesis and remodeling.

Light-sheet microscopy (LSM) plays a pivotal role in comprehending the intricate three-dimensional (3D) structure of the heart, providing crucial insights ...

Development and Evaluation of 3D-Printed Cardiovascular Phantoms for Interventional Planning and Training

Catheter-based interventions are standard treatment options for cardiovascular pathologies. Therefore, patient-specific models could help training physicians&#39; wire-skills as well as improving planning of interventional procedures. The aim of this study was to develop a manufacturing process of patient-specific 3D-printed models for cardiovascular interventions.
To create a 3D-printed elastic phantom, different 3D-printing materials were compared to porcine biological tissues (i.e., aortic tissue) in terms of mechanical characteristics. A fitting material was selected based on comparative tensile tests and specific material thicknesses were defined. Anonymized contrast-enhanced CT-datasets were collected retrospectively. Patient-specific volumetric models were extracted from these datasets and subsequently 3D-printed. A pulsatile flow loop was constructed to simulate the intraluminal blood flow during interventions. Models&#39; suitability for clinical imaging was assessed by x-ray imaging, CT, 4D-MRI and (Doppler) ultrasonography. Contrast medium was used to enhance visibility in x-ray-based imaging. Different catheterization techniques were applied to evaluate the 3D-printed phantoms in physicians&#39; training as well as for pre-interventional therapy planning.
Printed models showed a high printing resolution (~30 &#181;m) and mechanical properties of the chosen material were comparable to physiological biomechanics. Physical and digital models showed high anatomical accuracy when compared to the underlying radiological dataset. Printed models were suitable for ultrasonic imaging as well as standard x-rays. Doppler ultrasonography and 4D-MRI displayed flow patterns and landmark characteristics (i.e., turbulence, wall shear stress) matching native data. In a catheter-based laboratory setting, patient-specific phantoms were easy to catheterize. Therapy planning and training of interventional procedures on challenging anatomies (e.g., congenital heart disease (CHD)) was possible.
Flexible patient-specific cardiovascular phantoms were 3D-printed, and the application of common clinical imaging techniques was possible. This new process is ideal as a training tool for catheter-based (electrophysiological) interventions and can be used in patient-specific therapy planning.

Here we present development of a mock circulation setup for multimodal therapy evaluation, pre-interventional planning, and physician-training on cardiovascular anatomies. With the application of patient-specific tomographic scans, this setup is ideal for therapeutic approaches, training, and education in individualized medicine.

Catheter-based interventions are standard treatment options for cardiovascular pathologies. Therefore, patient-specific models could help training ...

Mesoscopic Optical Imaging of Whole Mouse Heart

Both genetic and non-genetic cardiac diseases can cause severe remodeling processes in the heart. Structural remodeling, such as collagen deposition (fibrosis) and cellular misalignment, can affect electrical conduction, introduce electromechanical dysfunctions and, eventually, lead to arrhythmia. Current predictive models of these functional alterations are based on non-integrated and low-resolution structural information. Placing this framework on a different order of magnitude is challenging due to the inefficacy of standard imaging methods in performing high-resolution imaging in massive tissue. In this work, we describe a methodological framework that allows imaging of whole mouse hearts with micrometric resolution. The achievement of this goal has required a technological effort where advances in tissue transformation and imaging methods have been combined. First, we describe an optimized CLARITY protocol capable of transforming an intact heart into a nanoporous, hydrogel-hybridized, lipid-free form that allows high transparency and deep staining. Then, a fluorescence light-sheet microscope able to rapidly acquire images of a mesoscopic field of view (mm-scale) with the micron-scale resolution is described. Following the mesoSPIM project, the conceived microscope allows the reconstruction of the whole mouse heart with micrometric resolution in a single tomographic scan. We believe that this methodological framework will allow clarifying the involvement of the cytoarchitecture disarray in the electrical dysfunctions and pave the way for a comprehensive model that considers both the functional and structural data, thus enabling a unified investigation of the structural causes that lead to electrical and mechanical alterations after the tissue remodeling.

We report a method for mesoscopic reconstruction of the whole mouse heart by combining new advancements in tissue transformation and staining with the development of an axially scanned light-sheet microscope.

Both genetic and non-genetic cardiac diseases can cause severe remodeling processes in the heart. Structural remodeling, such as collagen deposition ...

Pipeline for Multi-Scale Three-Dimensional Anatomic Study of the Human Heart

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