We express our gratitude to&#160;Dr. Eric Olson&#39;s group at UT Southwestern Medical Center for generously sharing the animal models. We appreciate all the constructive comments provided by D-incubator members at UT Dallas. This work was supported by NIH R00HL148493 (Y.D.), R01HL162635 (Y.D.), and UT Dallas STARS program (Y.D.).

Animal protocols and experiments have been approved and conducted under the oversight of the University of Texas at Dallas Institutional Animal Care and Use Committee (IACUC #21-03). C57BL6 mice, including neonates at postnatal day 1 (P1) and 8-week-old adults, were used in this study. No difference was observed between males and females. All data acquisition and image post-processing were carried out using open-source software or platforms with research or educational licenses. The resources are available from the authors upon reasonable request.
1. Sample preparation and tissue clearing (6 - 10 days)
<ol>
	<li>Prepare all chemical solutions in the Table of Materials required for the tissue clearing method before initiating the tissue clearing procedure. 
	NOTE: Perform all procedures in a fume hood while wearing proper personal protective equipment, especially when handling toxic chemicals.</li>
	<li>Euthanize animals with CO2 by placing them in a CO2 chamber at the desired time points, such as P1 and 8 weeks, for whole heart collection and immerse the freshly isolated heart at room temperature (RT) in 0.2 M KCl to arrest in diastole16.</li>
	<li>Fix the heart by immersing in 4% paraformaldehyde (PFA) in 1x phosphate buffered saline (PBS) with gentle agitation on a rocker overnight to preserve the cardiac anatomy at 4 &#176;C, see Table 1. 
	CAUTION: PFA is a toxic chemical and should be performed under a fume hood.</li>
	<li>Wash the heart with 1x PBS at RT for 30 min, 3x. 
	NOTE: Optional: Embed the heart in agarose gel to create a heart assembly for the custom-built LSM system in step 2 as described below.
	<ol>
		<li>Prepare the 1% agarose solution by dissolving agarose in PBS and heat the mixture in the microwave until complete dissolution. Pour the solution into a mold to create a bottom layer until the solution cools down to 40 &#176;C.</li>
		<li>Place the heart in the mold and fill the mold with agarose gel until it solidifies. Eliminate any bubbles in the agarose gel to minimize artifacts in the imaging. 
		NOTE: Embedding the sample reduces the risk of potential damage to the heart anatomy during mounting. 
		CAUTION: Utilize low-melting-point agarose and avoid placing the heart directly in heated agarose to mitigate the risk of exposing the sample to elevated temperatures.</li>
	</ol>
	</li>
	<li>Dehydrate the heart using gradients of methanol mixed with deionized water (Figure 1B), processing through sequential concentrations of 20%, 40%, 60%, 80%, and 100% as outlined in Table 1. Repeat with fresh 100% methanol 1x to ensure complete dehydration. Dehydrate neonatal mouse hearts for 1 h in each methanol mixture with shaking. Adjust the incubation time to 2 h for 8-week-old mouse hearts and rat hearts. 
	CAUTION: Methanol is a volatile, irritating, and flammable chemical.</li>
	<li>Replace and chill the heart with fresh 100% methanol for 10 min at 4 &#176;C.</li>
	<li>Replace the solution and bleach the heart with 5% hydrogen peroxide (H2O2) in methanol using a volume ratio of 1:5 (30% H2O2 to 100% methanol) overnight at 4 &#176;C to remove the pigments. 
	NOTE: The removal of pigments allows for minimizing photon absorption, leading to improved image quality with reduced artifacts.</li>
	<li>Replace the solution and incubate the heart in 100% methanol for 1 h at RT.</li>
	<li>Delipidate the heart with the solution containing 66% dichloromethane (DCM) and 33% methanol for 3 h with shaking. Ensure the heart sinks to the bottom of the vial by the end of this step. If not, renew the solution (66% DCM / 33% methanol) and extend the incubation time (e.g., 1 more hour) to achieve full delipidation. Transfer the heart into a fresh solution quickly to prevent desiccation, as DCM is highly volatile. 
	CAUTION: Use polypropylene or glassware for the experiment, as DCM is not compatible with&#160;polystyrene.</li>
	<li>Utilize dibenzyl ether (DBE) for refractive index matching (RI = 1.56). For neonatal mouse hearts, the process of refractive index matching takes 2 days, while for 8-week-old mouse hearts, it takes 5 days with mild shaking. Replace it with fresh DBE and extend the incubation time until the heart achieves complete transparency. 
	CAUTION: DBE is hazardous. Avoid any contact with the skin.</li>
</ol>
2. Sample mounting (1 day)
NOTE: In case a commercial LSM system is used, follow its specific protocol provided by the company to fix the heart and skip steps 2.1 - 2.9.
<ol>
	<li>Customize a chamber and a holder resistant to the refractive index-matching solution (e.g., DBE in the adapted iDISCO approach) using a 3D printer and Onyx materials (Figure 2A)16.</li>
	<li>Affix a magnet to the bottom of the chamber to facilitate quick connection and accurate localization under the LSM system (Figure 2B).</li>
	<li>Attach 25 mm x 50 mm x 0.17 mm cover glasses to the chamber using transparent silicone glue and verify the chamber&#39;s waterproof integrity by filling it with DBE and checking for leaks after 1 day.</li>
	<li>Customize a clamp holder using a 3D printer and affix a magnet for mounting the heart assembly inside the chamber (Figure 2C) or insert a needle into the agarose gel for mounting the heart assembly.</li>
	<li>Carry out the development of an in-house LSM system using a cylindrical lens and continuous-wave diode-pumped solid-state (DPSS) lasers as previously reported16,22. Use the excitation wavelength of 532 nm to capture autofluorescence from the myocardium (Figure 2D).</li>
	<li>Mount the chamber on a 6-dimensional stage and locate the optimal position where the chamber is perpendicular to the direction of laser propagation.</li>
	<li>Use a magnetic clamp holder to position the sample in the middle of the chamber and connect the holder to a 4-dimensional motorized stage (X, Y, Z, and Yaw; Figure 2D).</li>
	<li>Submerge the heart assembly into the chamber filled with DBE using the motorized stage and determine the scanning range along the detection axis (Z-direction).</li>
	<li>Determine the scanning velocity (Vz) of the motor along the Z-direction based on the step size (S) and acquisition time of an image (T) using the following equation: 
	<img alt="Equation 1" src="/files/ftp_upload/66707/66707eq01.jpg" style="margin: auto;" /> 
	NOTE: The relationship between the step size and the axial resolution of the imaging system adheres to the Nyquist-Shannon sampling theorem. For instance, the step size was set at 1 &#181;m, given that the axial resolution was 2.91 &#181;m, as indicated in the previous report16.</li>
	<li>To verify the spatial resolution and minimize potential opacity issues at different depths, measure the point spread function (PSF) of the system by imaging fluorescent beads with the diameter of 0.53 &#181;m diluted in 1% low melting point agarose gel with a concentration of 1:1.5 x 105. Calculate the PSF across the entire sample by measuring the full width at half maximum (FWHM)16.</li>
</ol>
3. Image stitching (4-8 h)
<ol>
	<li>Calculate the number of tiles required to cover the entire mouse heart, considering the limited FOV. For each tile, the dimensions correspond to the FOV of the LSM system. For instance, a neonatal mouse heart with a cross-sectional measurement of 3 x 3.6 mm2 necessitates 2 x 2 tiles for coverage in the customized LSM (Figure 3A). In contrast, an 8-week-old mouse heart measuring 5 x 8 mm2 requires 3 x 4 tiles to cover the entire cardiac structure (Figure 3B). 
	NOTE: Image stitching is required if the conventional LSM has limited FOV for imaging the whole heart.</li>
	<li>Capture images of the heart starting from tile 1, i.e., the top left corner tile of the heart (Figure 3A-B).</li>
	<li>Sequentially capture images of the remaining tiles with a 10% overlap between consecutive tiles until the whole heart is covered, as illustrated in Figure 3C.</li>
	<li>Download and install Fiji23 open-source software. Download and install the Fiji plugin, BigStitcher24.</li>
	<li>Navigate to the Help menu, select Update, choose Manage update site, and opt for BigStitcher. Subsequently, click Close, followed by Apply changes. Upon completing the plugin download, restart the Fiji software.</li>
	<li>Open the BigStitcher plugin and import all necessary tiles through the image file directory. Save the dataset as an HDF5 file.</li>
	<li>Organize tiles by choosing Move Tile By Regular Grid, select the pattern that is used for moving the heart, and select a 10% overlap between each tile. Stitch the tiles using the Stitching Wizard option.</li>
	<li>Export the stitched data using Image Fusion, to generate a tiff file.</li>
</ol>
4. Multiview deconvolution (5 days)
<ol>
	<li>Utilize fluorescent beads for image registration of multiview reconstruction along the heart (Figure 3D).
	<ol>
		<li>Prepare resin25 by mixing Bisphenol-A diglycidyl ether (D.E.R. 332), Isophorone diamine, 5-amino-1,3,3-trimethylcyclohexanemethylamine (IPDA), and Polypropylene glycol diglycidyl ether (D.E.R 736) at a volume ratio of 4:1:1.</li>
		<li>Centrifuge 10 &#181;L of fluorescence beads at 161 x g for 5 min and add 20 &#181;L of methanol to replace the storage solution of the beads. Repeat it 3x.</li>
		<li>Prepare a 10 mL solution with 1:1 x 105 bead concentration by mixing the methanol-based bead solution into the resin prepared in step 4.1.1.</li>
		<li>Degas the solution in a vacuum chamber for 3 h to remove air pockets and bubbles.</li>
		<li>Pour the solution into a silicone mold or a plastic drinking straw and wait for 2 to 3 days until it solidifies. After 1 day before the resin becomes solid, put the glass tube inside the mold and wait until the tube attaches to the resin. Remove the resin from the mold with the glass tube (Figure 3D).</li>
		<li>Place the heart into the glass tube, fill it with DBE, and mount the glass tube in the chamber filled with DBE.</li>
	</ol>
	</li>
	<li>Capture sequential images of the mouse heart along with the beads from the illuminated section by scanning the heart assembly across the detection axis (Z-axis; Figure 3E-F). 
	NOTE: Implement image stitching in step 3 to generate individual stacks of images if the heart size exceeds the FOV.</li>
	<li>Rotate the mouse heart 60&#176; along the Y-axis to capture subsequent stacks from multiple angles, i.e., 0&#176;, 60&#176;, 120&#176;, 180&#176;, 240&#176;, and 300&#176; (Figure 3E-F).</li>
	<li>Open the BigStitcher plugin and import all images through the image file directory. Save the dataset as an HDF5 file.</li>
	<li>Choose Detect Interest Points and manually select beads of interest for registration.</li>
	<li>Select Affine transformation model for registration. Choose Point Spread Function and assign PSF to all views.</li>
	<li>Click on Multiview Deconvolution and choose the type of iteration as Efficient Bayesian. Define Number of iterations as 10 and execute it26.</li>
	<li>Click on Image Fusion to export a tiff file. 
	NOTE: More detailed information about the multiview deconvolution can be found in other reports or protocols24,27,28.</li>
</ol>
5. Axially swept light-sheet system hardware (1 day)
<ol>
	<li>Install an electric tunable lens (ETL) on the Fourier plane of the cylindrical lens (CL)27 (Figure 2D) to scan the laser beam axially14. Ensure the liquid interface in ETL is horizontal to prevent wavefront distortion caused by gravity. Align ETL precisely to minimize beam decentering and higher-order optical aberrations30.</li>
	<li>Install a DAQ card in the workstation and connect a BNC cable to the DAQ card to synchronize camera exposure and ETL scanning.</li>
	<li>Open the housing of the ETL driver and remove the cover (Figure 4).</li>
	<li>Solder the signal wire of the BNC cable to the Analog in B pin, as labeled on the bottom of the PCB board (Figure 4).</li>
	<li>Solder the ground wire of the BNC cable to the GND pin; after that connect the other end of the BNC cable to AO (Analog Output) in the DAQ card (Figure 4).</li>
	<li>Plug the ETL Driver into the USB port of the workstation, connect the driver to the ETL using a hirose cable, and install Lens Driver Controller Software.</li>
	<li>Change Operation mode in Lens Driver Controller Software to Analog to control the ETL with a trigger generated from the DAQ card.</li>
	<li>In the software of the sCMOS camera, switch Capture mode to External (Light-sheet). Ensure that the active pixel scanning direction is perpendicular to the laser scanning direction to assist the synchronization.</li>
	<li>Connect the external trigger port on the back of the sCMOS camera to the AO of the DAQ card using a BNC cable (Figure 4).</li>
</ol>
6. Axially swept light-sheet system synchronization (7 days)
<ol>
	<li>Define exposure time based on the acceptable signal-to-background ratio (SBR) which for the proposed study, is at least 10. Increase the exposure time if SBR is lower than 10. 
	NOTE: Exposure time ranging from 10 to 50 ms provides an acceptable SBR in this project.</li>
	<li>Define the acquisition time of an image (T) and the line interval (L) based on the exposure time (E) for the camera and the ETL frequency (f) using the following equations: 
	<img alt="Equation 2" src="/files/ftp_upload/66707/66707eq02.jpg" style="margin: auto;" /> 
	<img alt="Equation 3" src="/files/ftp_upload/66707/66707eq03.jpg" style="margin: auto;" /> 
	Where P indicates the number of pixel columns on the camera sensor. The representative parameters for the reference are f = 1 Hz, P = 2048, E = 10 ms, and L = 243 &#181;s, respectively.</li>
	<li>In LabVIEW program Trigger Generator.vi, generates both square and triangle triggers for synchronization (Figure 5). 
	NOTE: The customized LabVIEW program is available from the authors upon reasonable request.</li>
	<li>Mount fluorescent beads diluted in 1% low melting point agarose gel16 with a concentration of 1:1 x 103 to locate the position of the beam waist in the image (Figure 6A).</li>
	<li>In the program, identify the starting and ending points of the laser beam under the FOV by changing the voltage along the scanning direction.</li>
	<li>To synchronize the sCMOS camera and ETL, scan the laser beam along the X-axis and active pixels along the Y-axis to generate a 2D image (Figure 6B). The brighter and sharper beads along the diagonal in the image indicate the precise synchronization between the sCMOS and ETL. 
	NOTE: The synchronization of the scanning speed of ETL and sCMOS pixel activation is critical for image quality. Various common asynchronous errors are summarized in Figure 6C-F.</li>
	<li>After determining optimal synchronization parameters for synchronization, rotate the sCMOS camera 90&#176; around the Z-axis to align the active pixel direction with the laser scanning direction.</li>
	<li>Repeat step 2.10, and measure the FWHM of the PSF. Use the same method as indicated in the previous report16, with the averaged lateral and axial resolutions of ASLM as 2.18 &#181;m and 2.88 &#181;m, respectively (Figure 7). 
	NOTE: Uniform illumination and resolution across the entire FOV are feasible under ideal conditions (Figure 7A-B). Asynchronous operation of the ETL with the sCMOS camera, as well as misalignment of the illumination and detection, can result in non-uniform spatial resolution (Figure 7C-D).</li>
	<li>For cardiac imaging, after aligning the system and determining the optimal parameters for system synchronization, proceed with steps 2.6 to 2.9.</li>
</ol>

Light-Sheet Microscopy and Tissue Clearing for Comprehensive Cardiac Imaging

The authors have no conflict of interest to disclose.

The advancement of imaging, computation, and tissue clearing methods has provided an unparalleled opportunity to extensively investigate cardiac structure and function. This holds great potential for deepening our understanding of cardiac morphogenesis and pathogenesis using an intact rodent heart model. In contrast to in vivo studies of zebrafish heart using a similar approach40,41,42,43</s...

Heart failure remains the leading cause of mortality worldwide, primarily due to the lack of regenerative capacity of mature cardiomyocytes1. The intricate architecture of the heart plays a crucial role in its function and provides insights into developmental processes. A profound understanding of cardiac structure is essential for elucidating the fundamental processes of cardiac morphogenesis and remodeling in response to myocardial infarction. Recent progress has demonstrated that neonatal mice can restore cardiac function following injury, while adult mice lack such regenerative capacity2. This establishes a foundation for investigating cues associated with structural and functional abnormalities in mouse models. Traditional imaging methods, such as confocal microscopy, have technical limitations, including restricted penetration depth, slow point-scanning scheme, and photo damage from prolonged exposure to laser light. These hinder comprehensive three-dimensional (3D) imaging of the intact heart. In this context, light-sheet microscopy (LSM) emerges as a powerful solution, offering the advantages of high-speed imaging, reduced photo damage, and exceptional optical sectioning capabilities3,4,5. The unique features of LSM position it as a promising method to overcome the limitations of conventional techniques, providing unprecedented insights into cardiac development and remodeling processes6,7,8.
In this protocol, we introduce an imaging strategy that combines advanced LSM with adapted tissue clearing approaches9, allowing for the imaging of entire mouse hearts without the need for specific labeling and mechanical sectioning. We further propose that conventional LSM imaging can be enhanced through multiview deconvolution10 or axially swept light-sheet microscopy (ASLM) techniques11,12,13,14,15 to improve axial resolution. Additionally, the integration of image stitching with either of these methods can effectively overcome the trade-off between spatial resolution and field of view (FOV), thereby advancing the imaging of adult mouse hearts. The incorporation of numerous tissue clearing approaches, including hydrophobic, hydrophilic, and hydrogel-based methods, enables deeper light penetration for capturing the morphology of the entire heart16,17,18,19.
While multiple clearing methods are compatible with current LSM systems, the goal is to minimize photon scattering and enhance light penetration in tissues, like the heart, by replacing lipids with a medium that closely matches its refractive index. iDISCO was chosen as the representative20,21 and adapted for autofluorescence imaging in this protocol due to its rapid processing and high transparency (Figure 1A). Collectively, the integration of the advanced LSM approach with tissue clearing techniques offers a promising framework to unravel intricate cardiac anatomy in rodent hearts, holding significant potential for advancing our understanding of cardiac morphogenesis and pathogenesis.

<table><tbody><tr><td>&lt;strong&gt;1% Agarose&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Low melting point agarose</td><td>Thermo Fisher</td><td>16520050</td><td></td></tr><tr><td>Deionized water</td><td>-</td><td>-</td><td></td></tr><tr><td>&lt;strong&gt;Chemicals for tissue clearing&amp;nbsp;&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>5-Amino-1,3,3-trimethylcyclohexanemethylamine, mixture of cis and trans</td><td>Sigma-Aldrich</td><td>118184</td><td></td></tr><tr><td>D.E.R.&amp;trade; 332</td><td>Sigma-Aldrich</td><td>31185</td><td></td></tr><tr><td>D.E.R.&amp;trade; 736</td><td>Sigma-Aldrich</td><td>31191</td><td></td></tr><tr><td>Dibenzyl ether (DBE)</td><td>Sigma-Aldrich</td><td>33630</td><td></td></tr><tr><td>Dichloromethane (DCM)</td><td>Sigma-Aldrich</td><td>270997</td><td></td></tr><tr><td>Fluorescent beads</td><td>Spherotech</td><td>FP-0556-2</td><td></td></tr><tr><td>Hydrogen peroxide (H&lt;sub&gt;2&lt;/sub&gt;O&lt;sub&gt;2&lt;/sub&gt;)</td><td>Sigma-Aldrich</td><td>216736</td><td></td></tr><tr><td>Methanol</td><td>Sigma-Aldrich</td><td>439193</td><td></td></tr><tr><td>Paraformaldehyde (PFA)</td><td>Thermo Fisher</td><td>47392</td><td></td></tr><tr><td>Phosphate Buffered Saline (PBS)</td><td>Sigma-Aldrich</td><td>79383</td><td></td></tr><tr><td>Potassium Chloride (KCl)</td><td>Sigma-Aldrich</td><td>P3911</td><td></td></tr><tr><td>&lt;strong&gt;Software and algorithms&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Amira</td><td>Thermo Fisher Scientific</td><td>2021.2</td><td></td></tr><tr><td>BigStitcher</td><td>H&amp;ouml;rl et al.&lt;sup&gt;22&lt;/sup&gt;</td><td></td><td></td></tr><tr><td>Fiji-ImageJ</td><td>Schindelin et al.&lt;sup&gt;20&lt;/sup&gt;</td><td>1.54f</td><td></td></tr><tr><td>HCImage Live</td><td>Hamamatsu Photonics</td><td>4.6.1.2</td><td></td></tr><tr><td>LabVIEW</td><td>National Instruments Corporation</td><td>2017 SP1</td><td></td></tr><tr><td>&lt;strong&gt;Key components of the customized light-sheet system&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>0.63 - 6.3X Zoom body</td><td>Olympus</td><td>MVX-ZB10&amp;nbsp;</td><td></td></tr><tr><td>10X Illumination objective</td><td>Nikon</td><td>MRH00105</td><td></td></tr><tr><td>1X detection objective</td><td>Olympus</td><td>MV PLAPO 1X/0.25&amp;nbsp;</td><td></td></tr><tr><td>473nm DPSS Laser</td><td>Laserglow Technologies</td><td>LRS-0473-PFM-00100-05</td><td></td></tr><tr><td>532nm DPSS laser</td><td>Laserglow Technologies</td><td>LRS-0532-PFM-00100-05</td><td></td></tr><tr><td>589&amp;thinsp;nm DPSS laser</td><td>Laserglow Technologies</td><td>LRS-0589-GFF-00100-05</td><td></td></tr><tr><td>BNC connector</td><td>National Instrument</td><td>BNC-2110</td><td></td></tr><tr><td>Cylindrical lens</td><td>Thorlabs</td><td>ACY254-050-A</td><td></td></tr><tr><td>DC-Motor Controller, 4 axes</td><td>Physik Instrumente</td><td>C-884.4DC</td><td></td></tr><tr><td>ETL</td><td>Optotune</td><td>EL-16-40-TC-VIS-5D-1-C</td><td></td></tr><tr><td>ETL Cable</td><td>Optotune</td><td>CAB-6-300</td><td></td></tr><tr><td>ETL Lens Driver</td><td>Optotune</td><td>EL-E-4i</td><td></td></tr><tr><td>Filter</td><td>Chroma</td><td>ET525/30</td><td></td></tr><tr><td>Filter</td><td>Chroma</td><td>ET585-40</td><td></td></tr><tr><td>Filter</td><td>Chroma</td><td>ET645-75</td><td></td></tr><tr><td>Filter wheel&amp;nbsp;</td><td>Shutter Instrument</td><td>LAMBDA 10-B</td><td></td></tr><tr><td>Motorized translation stage</td><td>Physik Instrumente</td><td>L-406.20DG10</td><td></td></tr><tr><td>Motorized translation stage</td><td>Physik Instrumente</td><td>L-406.40DG10</td><td></td></tr><tr><td>Motorized translation stage</td><td>Physik Instrumente</td><td>M-403.4PD</td><td></td></tr><tr><td>NI multifunction I/O</td><td>National Instrument</td><td>PCIe-6363</td><td></td></tr><tr><td>sCMOS camera</td><td>Hamamatsu</td><td>C13440-20CU</td><td></td></tr><tr><td>Stepper motor</td><td>Pololu</td><td>1474</td><td></td></tr><tr><td>Tube lens</td><td>Olympus</td><td>MVX-TLU</td><td></td></tr></tbody></table>

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.

LSM has been demonstrated to foster cardiac studies31,32,33,34,35,36,37 due to the minimal risk of photo damage, high spatial resolution, and optical sectioning as opposed to other optical imaging methods such as brightfield and point-scanning techniques6,<...

Read this Scientific Article Protocol about Author Spotlight: Customized Light-Sheet Imaging for Investigating Myocardial Structures in Rodent Hearts at JoVE.com

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 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 ...

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755 Views.  The University of Texas at Dallas. 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.

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

Three-dimensional Optical-resolution Photoacoustic Microscopy

Optical microscopy, providing valuable insights at the cellular and organelle levels, has been widely recognized as an enabling biomedical technology. As the mainstays of in vivo three-dimensional (3-D) optical microscopy, single-/multi-photon fluorescence microscopy and optical coherence tomography (OCT) have demonstrated their extraordinary sensitivities to fluorescence and optical scattering contrasts, respectively. However, the optical absorption contrast of biological tissues, which encodes essential physiological/pathological information, has not yet been assessable. 
The emergence of biomedical photoacoustics has led to a new branch of optical microscopy optical-resolution photoacoustic microscopy (OR-PAM)1, where the optical irradiation is focused to the diffraction limit to achieve cellular1 or even subcellular2 level lateral resolution. As a valuable complement to existing optical microscopy technologies, OR-PAM brings in at least two novelties. First and most importantly, OR-PAM detects optical absorption contrasts with extraordinary sensitivity (i.e., 100%). Combining OR-PAM with fluorescence microscopy3 or with optical-scattering-based OCT4 (or with both) provides comprehensive optical properties of biological tissues. Second, OR-PAM encodes optical absorption into acoustic waves, in contrast to the pure optical processes in fluorescence microscopy and OCT, and provides background-free detection. The acoustic detection in OR-PAM mitigates the impacts of optical scattering on signal degradation and naturally eliminates possible interferences (i.e., crosstalks) between excitation and detection, which is a common problem in fluorescence microscopy due to the overlap between the excitation and fluorescence spectra. 
Unique for optical absorption imaging, OR-PAM has demonstrated broad biomedical applications since its invention, including, but not limited to, neurology5, 6, ophthalmology7, 8, vascular biology9, and dermatology10. In this video, we teach the system configuration and alignment of OR-PAM as well as the experimental procedures for in vivo functional microvascular imaging.

Optical-resolution photoacoustic microscopy (OR-PAM) is an emerging technology capable of imaging optical absorption contrasts in vivo with cellular resolution and sensitivity. Here, we provide a visualized instruction on the experimental protocols of OR-PAM, including system configuration, system alignment, typical in vivo experimental procedures, and functional imaging schemes.

Optical microscopy, providing valuable insights at the cellular and organelle levels, has been widely recognized as an enabling biomedical technology. As ...

Shock Wave Application to Cell Cultures

Shock waves nowadays are well known for their regenerative effects. Basic research findings showed that shock waves do cause a biological stimulus to target cells or tissue without any subsequent damage. Therefore, in vitro experiments are of increasing interest. Various methods of applying shock waves onto cell cultures have been described. In general, all existing models focus on how to best apply shock waves onto cells.
However, this question remains: What happens to the waves after passing the cell culture? The difference of the acoustic impedance of the cell culture medium and the ambient air is that high, that more than 99% of shock waves get reflected! We therefore developed a model that mainly consists of a Plexiglas built container that allows the waves to propagate in water after passing the cell culture. This avoids cavitation effects as well as reflection of the waves that would otherwise disturb upcoming ones. With this model we are able to mimic in vivo conditions and thereby gain more and more knowledge about how the physical stimulus of shock waves gets translated into a biological cell signal (&#8220;mechanotransduction&#34;).

Shock waves nowadays are well known for their regenerative effects. Therefore in vitro experiments are of increasing interest. We therefore developed a model for in vitro shock wave trials (IVSWT) that enables us to mimic in vivo conditions thereby avoiding distracting physical effects.

Shock waves nowadays are well known for their regenerative effects. Basic research findings showed that shock waves do cause a biological stimulus to ...

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 ...

In Vivo Quantitative Assessment of Myocardial Structure, Function, Perfusion and Viability Using Cardiac Micro-computed Tomography

The use of Micro-Computed Tomography (MicroCT) for in vivo studies of small animals as models of human disease has risen tremendously due to the fact that MicroCT provides quantitative high-resolution three-dimensional (3D) anatomical data non-destructively and longitudinally. Most importantly, with the development of a novel preclinical iodinated contrast agent called eXIA160, functional and metabolic assessment of the heart became possible. However, prior to the advent of commercial MicroCT scanners equipped with X-ray flat-panel detector technology and easy-to-use cardio-respiratory gating, preclinical studies of cardiovascular disease (CVD) in small animals required a MicroCT technologist with advanced skills, and thus were impractical for widespread implementation. The goal of this work is to provide a practical guide to the use of the high-speed Quantum FX MicroCT system for comprehensive determination of myocardial global and regional function along with assessment of myocardial perfusion, metabolism and viability in healthy mice and in a cardiac ischemia mouse model induced by permanent occlusion of the left anterior descending coronary artery (LAD).

In Vivo Quantitative Assessment of Myocardial Structure, Function, Perfusion and Viability Using Cardiac Micro-computed Tomography

This method outlines the use of Quantum Micro-Computed Tomography (MicroCT) to assess cardiac morphology, function, perfusion, metabolism and viability with iodinated contrast agent in mice with experimentally-induced myocardial ischemia. The technique can be applied for non-destructive high-throughput longitudinal in vivo imaging of various animal models of human heart disease.

The use of Micro-Computed Tomography (MicroCT) for in vivo studies of small animals as models of human disease has risen tremendously due to the fact that ...

Using Extraordinary Optical Transmission to Quantify Cardiac Biomarkers in Human Serum

For a biosensing platform to have clinical relevance in point-of-care (POC) settings, assay sensitivity, reproducibility, and ability to reliably monitor analytes against the background of human serum are crucial.
Nanoimprinting lithography (NIL) was used to fabricate, at a low cost, sensing areas as large as 1.5 mm x 1.5 mm. The sensing surface was made of high-fidelity arrays of nanoholes, each with an area of about 140 nm2. The great reproducibility of NIL made it possible to employ a one-chip, one-measurement strategy on 12 individually manufactured surfaces, with minimal chip-to-chip variation. These nanoimprinted localized surface plasmon resonance (LSPR) chips were extensively tested on their ability to reliably measure a bioanalyte at concentrations varying from 2.5 to 75 ng/mL amidst the background of a complex biofluid-in this case, human serum. The high fidelity of NIL enables the generation of large sensing areas, which in turn eliminates the need for a microscope, as this biosensor can be easily interfaced with a commonly available laboratory light source. These biosensors can detect cardiac troponin in serum with a high sensitivity, at a limit of detection (LOD) of 0.55 ng/mL, which is clinically relevant. They also show low chip-to-chip variance (due to the high quality of the fabrication process). The results are commensurable with widely used enzyme-linked immunosorbent assay (ELISA)-based assays, but the technique retains the advantages of an LSPR-based sensing platform (i.e., amenability to miniaturization and multiplexing, making it more feasible for POC applications).

This work describes a nanoimprinting lithography method to fabricate high-quality sensing arrays that work on the principle of extraordinary optical transmission. The biosensor is low-cost, robust, easy to use, and can detect cardiac troponin I in serum at clinically relevant concentrations (99th percentile cutoff &#8764;10-400 pg/mL, depending on the assay).

For a biosensing platform to have clinical relevance in point-of-care (POC) settings, assay sensitivity, reproducibility, and ability to reliably monitor ...

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. ...

Analysis of Zebrafish Cardiac Contraction with Virtual Reality Interaction

4D Light-sheet Imaging of Zebrafish Cardiac Contraction

Zebrafish is an intriguing model organism known for its remarkable cardiac regeneration capacity. Studying the contracting heart in vivo is essential for gaining insights into structural and functional changes in response to injuries. However, obtaining high-resolution and high-speed 4-dimensional (4D, 3D spatial + 1D temporal) images of the zebrafish heart to assess cardiac architecture and contractility remains challenging. In this context, an in-house light-sheet microscope (LSM) and customized computational analysis are&#160;used to overcome these technical limitations. This strategy, involving LSM system construction, retrospective synchronization, single cell tracking, and user-directed analysis, enables one to investigate the micro-structure and contractile function across the entire heart at the single-cell resolution in the transgenic Tg(myl7:nucGFP) zebrafish larvae. Additionally, we are able to further incorporate microinjection of small molecule compounds to induce cardiac injury in a precise and controlled manner. Overall, this framework allows one to track physiological and pathophysiological changes, as well as the regional mechanics at the single-cell level during cardiac morphogenesis and regeneration.

Author Spotlight: High-Resolution 4D Light-Sheet Imaging and Virtual Reality in Zebrafish for Single-Cell Analysis of Heart Function

This protocol utilizes light-sheet imaging to investigate cardiac contractile function in zebrafish larvae and gain insights into cardiac mechanics through cell tracking and interactive analysis.

Zebrafish is an intriguing model organism known for its remarkable cardiac regeneration capacity. Studying the contracting heart in vivo is essential for ...

Light Sheet Microscopy of Fast Cardiac Dynamics in Zebrafish Embryos

Embryonic cardiac research has greatly benefited from advances in fast in vivo light sheet fluorescence microscopy (LSFM). Combined with the rapid external development, tractable genetics, and translucency of the zebrafish, Danio rerio, LSFM has delivered insights into cardiac form and function at high spatial and temporal resolution without significant phototoxicity or photobleaching. Imaging of beating hearts challenges existing sample preparation and microscopy techniques. One needs to maintain a healthy sample in a constricted field of view and acquire ultrafast images to resolve the heartbeat. Here we describe optimized tools and solutions to study the zebrafish heart in vivo. We demonstrate the applications of bright transgenic lines for labeling the cardiac constituents and present novel gentle embedding and immobilization techniques that avoid developmental defects and changes in heart rate. We also propose a data acquisition and analysis pipeline adapted to cardiac imaging. The entire workflow presented here focuses on zebrafish embryonic heart imaging but can also be applied to various other samples and experiments.

We describe optimized tools to study the zebrafish heart in vivo with light sheet fluorescence microscopy. Specifically, we suggest bright cardiac transgenic lines and present new gentle embedding and immobilization techniques that avoid developmental and heart defects. A possible data acquisition and analysis pipeline adapted to cardiac imaging is also provided.

Embryonic cardiac research has greatly benefited from advances in fast in vivo light sheet fluorescence microscopy (LSFM). Combined with the rapid ...

Biology

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 ...

Light-Sheet Imaging to Reveal Cardiac Structure in Rodent Hearts

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Three-dimensional Optical-resolution Photoacoustic Microscopy

Shock Wave Application to Cell Cultures

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

In Vivo Quantitative Assessment of Myocardial Structure, Function, Perfusion and Viability Using Cardiac Micro-computed Tomography

Using Extraordinary Optical Transmission to Quantify Cardiac Biomarkers in Human Serum

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

4D Light-sheet Imaging of Zebrafish Cardiac Contraction

Light Sheet Microscopy of Fast Cardiac Dynamics in Zebrafish Embryos

Mesoscopic Optical Imaging of Whole Mouse Heart