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, the integration of advanced LSM techniques and tissue clearing methods enables us to overcome challenges between spatial resolution and penetration depth when imaging the rodent model9,44. Tissue clearing techniques render hearts optically transparent by removing lipids and other light-scattering elements, enabling deeper imaging penetration45. When coupled with LSM, we are able to image the myocardium. This combination enhances the imaging depth and allows for the observation of intricate myocardial micro-structure. While numerous clearing methods have been established18, we adapted the iDISCO20 to rapidly render the intact heart transparent in comparison with other tissue clearing methods15, making it readily available for the imaging of autofluorescence from the myocardium. For specific fluorescence labeling, the original iDISCO and Adipo-Clear46 protocols can be combined with immunostaining in this customized imaging system, despite the longer incubation. Meanwhile, methods that preserve endogenous fluorescence and require shorter processing time are preferred for the proposed study. In contrast to other methods, this platform offers the following advantages. Firstly, we utilize an adapted iDISCO protocol for tissue clearing, enabling rapid rendering of the heart transparent in less than 1 week while retaining myocardium autofluorescence. Secondly, we incorporate computational methods such as image stitching and multiview deconvolution into this established light-sheet system for cardiac imaging. Lastly, we design and synchronize the ASLM system to enhance axial resolution across a large FOV, enabling multiscale imaging and analysis of myocardial compaction and trabeculation at a near isotropic resolution.
To capture cardiac images of the rodent model, we have provided four approaches based on LSM: i) the conventional method for screening purposes19, ii) image stitching, iii) multiview deconvolution, and iv) ASLM scanning. Considering the variation in heart size across different models, we sought to address the fundamental trade-off in spatial resolution, acquisition speed, and FOV. While image stitching enables the large FOV with compromised spatial resolution, combining it with multiview deconvolution or ASLM provides near-isotropic resolution across the extended FOV to cover the entire adult mouse heart. Furthermore, the integration of these methods enhances image quality and contrast in the deep tissue, allowing for a comprehensive digital reconstruction of 3D models of micro-structures such as myocardial trabeculation and compaction inside ventricles. However, the deluge of datasets, extensive computational processing cost, the need for precise image registration, and the potential for artifacts in multiview deconvolution and imaging stitching hinder their broad applications. While ASLM is an emerging alternative, the complexity of the system synchronization, laser scanning strategies, and reliance on tissue transparency pose challenges that may affect system robustness and increase the risk of photo damage to the heart.
Collectively, this methodology including LSM imaging and tissue clearing provides a starting point to explore cardiac structure from neonates to adults. It holds the potential to localize the distribution of specialized cardiac lineages and their functions during cardiac development47. With the development of computational analysis tailored for cardiac applications6,48,49,50,51,52,53 , this framework can be extended to investigate cardiac remodeling processes, particularly in response to myocardial infarction. Ultimately, this holistic strategy holds the potential to unravel the underlying mechanism of cardiac morphogenesis and advance the development of therapeutic interventions.

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1% Agarose</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>Chemicals for tissue clearing&#160;</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.&#8482; 332</td>
			<td>Sigma-Aldrich</td>
			<td>31185</td>
			<td></td>
		</tr>
		<tr>
			<td>D.E.R.&#8482; 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 (H2O2)</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>Software and algorithms</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&#246;rl et al.22</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fiji-ImageJ</td>
			<td>Schindelin et al.20</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>Key components of the customized light-sheet system</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>0.63 - 6.3X Zoom body</td>
			<td>Olympus</td>
			<td>MVX-ZB10&#160;</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&#160;</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&#8201;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&#160;</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 techniques6,8,38,39,40. Thus, to better understand the 3D myocardial architecture, we have established the protocol based on primary approaches including adapted iDISCO clearing, image stitching, multiview deconvolution, and ASLM. While we presented results using mouse models, it is important to note that rats and other rodent models are also suitable for the proposed methods. The adapted tissue clearing protocol enables the rendering of the P1 mouse heart transparent within 4 days and the 8-week-old mouse heart transparent within 7 days (Figure 1B). This step serves as a starting point for minimizing photon absorption and scattering in the intact heart. Multiview deconvolution enables the improvement of axial resolution by fusing images from multiple perspectives into a single model. After sequentially recording images from 6 views of the same heart, the multiview deconvolution method achieves near-isotropic resolution with a&#160;lateral of 4.68 &#181;m and axial of 5.06 &#181;m, following 15 h of computation (Figure 8A). To obviate the computational complexity, we further customized an ETL-based ASLM to image from neonatal to adult mouse hearts. This method allows us to scan the myocardium with a tightly focused laser beam, minimizing the out-of-focus background and enhancing the image contrast in comparison to the conventional LSM methods (Figure 8B). With the current ASLM design, the averaged lateral and axial resolutions of the system achieve 2.18 &#181;m and 2.88 &#181;m, respectively, enabling in-depth exploration of myocardial trabeculation in ventricles. In addition, image stitching can be utilized independently, or in combination with multiview deconvolution or ASLM to expand the FOV for larger sample sizes. Integrating stitching with ASLM allows us to cover the entire 8-week-old mouse heart with uniform resolution (Figure 8C), providing an effective solution to investigate the cross-section of the entire heart.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66707/66707fig01.jpg" /> 
Figure 1: Tissue clearing process. (A) The hydrophobic method involves tissue dehydration, lipid extraction, and refractive index matching using dibenzyl ether (DBE) as the organic solvent. (B) Submerge the heart in a mixture of 4% formaldehyde (PFA) solution and dehydrate it with a gradient of methanol and deionized (DI) water mixtures. Bleach the heart using 5% H2O2 in methanol at 4 &#176;C. Delipidate the heart with dichloromethane (DCM) and methanol solution until the sample sinks to the bottom of the tube. Lastly, incubate the heart in DBE in order to obtain the desired refractive index. Grid lines 5 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66707/66707fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66707/66707fig02.jpg" /> 
Figure 2: Schematic of axially swept light-sheet microscopy. (A) Customized 3D printed chamber, (B) chamber magnet, (C) clamp holder, and (D) ASLM system. Abbreviations: CL: cylindrical lens; ETL: electronic tunable lens; IL: illumination lens; DL: detection lens; FW: filter wheel; TL: tube lens. This figure has been modified from Ref16. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66707/66707fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66707/66707fig03.jpg" /> 
Figure 3: Image stitching and multiview deconvolution. (A-B) Image stitching is used to cover the entire mouse hearts, with each tile having a 10% FOV overlap with its adjacent tiles. (C) Movement pattern of the mouse heart for image stitching. (D) Fluorescent beads are affixed to the end of the glass tube to facilitate image registration, while the heart is positioned inside the glass tube containing DBE, enabling concurrent imaging of both the mouse heart and fluorescent beads. (E) Multiview deconvolution involves the acquisition of images from six distinct perspectives, each gathering images from unique directions. (F) Raw data of P1 mouse heart and beads at different angles of 0&#176;, 60&#176;, 120&#176;, 180&#176;, 240&#176;, and 300&#176;. Scale bars: 500 &#181;m. This figure has been modified from16. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66707/66707fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/66707/66707fig04.jpg" /> 
Figure 4: Block diagram of ASLM system. Utilizing DAQ card for synchronizing ETL and sCMOS camera. The housing of the ETL driver has been opened for soldering of signal and ground wires to the PCB. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66707/66707fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/66707/66707fig05.jpg" /> 
Figure 5: LabView control panel. LabVIEW control panel for generating triggers to synchronize ETL and activated pixels of sCMOS camera. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66707/66707fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/66707/66707fig06.jpg" /> 
Figure 6: Synchronizing the focus of light-sheet with activated pixels. (A)&#160;The identification of the starting and ending points of the light-sheet scanning range is achieved by configuring the appropriate voltage for the ETL trigger. (B) The synchronous result of fluorescent beads is evident along the diagonal of the image. (C-F) The non-uniform spatial resolution across the entire FOV arises from the asynchronous operation of the ETL with the sCMOS camera. The ETL initiates scanning (C) later or (D) earlier than the activated pixel. (E-F) The focal area is not parallel to the diagonal of the image, indicating an incompatibility between scanning and activation speeds. ETL scans (E) faster or (F) slower than the sweeping of active pixels in sCMOS. Scale bar: 200 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66707/66707fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/66707/66707fig07.jpg" /> 
Figure 7: Troubleshooting of ASLM. (A) The light-sheet is placed exactly at the focal distance of the detection lens. The ETL scans the focus of the light-sheet along the propagation direction while synchronizing with the activated pixel of the sCMOS sensor. (B) XY view of fluorescent beads when the ETL is correctly aligned and synchronized. (C-D) Results of fluorescent beads on the focal plane and out-of-focus in cross-sectional images, indicating the misalignment between the ETL scanning and laser propagation. Scale bars: 200 &#181;m and 10 &#181;m in the inset. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66707/66707fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/66707/66707fig08.jpg" /> 
Figure 8: Light-sheet imaging of mouse hearts. (A) Volume-rendered image of the multiview deconvolution P1 mouse heart highlighting the trabeculation within the ventricular cavity. (B) Cross-sectional images of the 8-week-old mouse heart generated by conventional LSM (left) and ASLM (right). Images are presented as raw data. (C) Combination of Image stitching and ASLM for imaging an 8-week-old mouse heart, comprising 12 tiles arranged in 3 tiles horizontally and 4 tiles vertically. Scale bar: 500 &#181;m. Abbreviations: LV: left ventricle, LA: left atrium, RV: right ventricle, and RA: right atrium. This figure has been modified from Ref16. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66707/66707fig08large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="3">Procedure</td>
			<td>Time (P1 mouse heart)</td>
			<td>Time (8-week-old mouse heart)</td>
			<td>Temperature</td>
			<td></td>
		</tr>
		<tr>
			<td colspan="3">Fix in 4% PFA</td>
			<td>Overnight</td>
			<td>Overnight</td>
			<td>4&#176;C</td>
			<td rowspan="5">Sample preparation&#160;</td>
		</tr>
		<tr>
			<td colspan="3">Wash with 1x PBS&#160;</td>
			<td>30 min</td>
			<td>30 min</td>
			<td>RT</td>
		</tr>
		<tr>
			<td colspan="3">Wash with 1x PBS&#160;</td>
			<td>30 min</td>
			<td>30 min</td>
			<td>RT</td>
		</tr>
		<tr>
			<td colspan="3">Wash with 1x PBS&#160;</td>
			<td>30 min</td>
			<td>30 min</td>
			<td>RT</td>
		</tr>
		<tr>
			<td colspan="3">Embed with agarose gel&#160;</td>
			<td>1 h</td>
			<td>2 h</td>
			<td>RT</td>
		</tr>
		<tr>
			<td colspan="3">Incubate in 20% methanol/80% DI water</td>
			<td>1 h</td>
			<td>2 h</td>
			<td>RT</td>
			<td rowspan="6">Dehydration</td>
		</tr>
		<tr>
			<td colspan="3">Incubate in 40% methanol/60% DI water</td>
			<td>1 h</td>
			<td>2 h</td>
			<td>RT</td>
		</tr>
		<tr>
			<td colspan="3">Incubate in 60% methanol/40% DI water</td>
			<td>1 h</td>
			<td>2 h</td>
			<td>RT</td>
		</tr>
		<tr>
			<td colspan="3">Incubate in 80% methanol/20% DI water</td>
			<td>1 h</td>
			<td>2 h</td>
			<td>RT</td>
		</tr>
		<tr>
			<td colspan="3">Incubate in 100% methanol</td>
			<td>1 h</td>
			<td>2 h</td>
			<td>RT</td>
		</tr>
		<tr>
			<td colspan="3">Incubate in fresh 100% methanol</td>
			<td>1 h</td>
			<td>2 h</td>
			<td>RT</td>
		</tr>
		<tr>
			<td colspan="3">Incubate in fresh 100% methanol</td>
			<td>10 min</td>
			<td>10 min</td>
			<td>4&#176;C</td>
			<td rowspan="3">Depigmentation</td>
		</tr>
		<tr>
			<td colspan="3">Bleach with 5% H2O2/methanol</td>
			<td>Overnight</td>
			<td>Overnight</td>
			<td>4&#176;C</td>
		</tr>
		<tr>
			<td colspan="3">Incubate in 100% methanol</td>
			<td>1 h</td>
			<td>1 h</td>
			<td>RT</td>
		</tr>
		<tr>
			<td colspan="3">Delipidate with 66% DCM/33% methanol on the shaker</td>
			<td>3 h</td>
			<td>3 h</td>
			<td>RT</td>
			<td>Delipidation</td>
		</tr>
		<tr>
			<td colspan="3">Incubate in DBE on the shaker</td>
			<td>2 days</td>
			<td>5 days</td>
			<td>RT</td>
			<td>RI matching</td>
		</tr>
	</tbody>
</table>
Table 1: Customized tissue clearing steps for autofluorescence imaging.

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 ...

light-sheet-imaging-to-reveal-cardiac-structure-in-rodent-hearts

Nanyang Technological University

Research

JoVE Journal

Bioengineering

742 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 ...

Protocol for MicroRNA Transfer into Adult Bone Marrow-derived Hematopoietic Stem Cells to Enable Cell Engineering Combined with Magnetic Targeting

While CD133+ hematopoietic stem cells (SCs) have been proven to provide high potential in the field of regenerative medicine, their low retention rates after injection into injured tissues as well as the observed massive cell death rates lead to very restricted therapeutic effects. To overcome these limitations, we sought to establish a non-viral based protocol for suitable cell engineering prior to their administration. The modification of human CD133+ expressing SCs using microRNA (miR) loaded magnetic polyplexes was addressed with respect to uptake efficiency and safety as well as the targeting potential of the cells. Relying on our protocol, we can achieve high miR uptake rates of 80&#8211;90% while the CD133+ stem cell properties remain unaffected. Moreover, these modified cells offer the option of magnetic targeting. We describe here a safe and highly efficient procedure for the modification of CD133+ SCs. We expect this approach to provide a standard technology for optimization of therapeutic stem cell effects and for monitoring of the administered cell product via magnetic resonance imaging (MRI).

This protocol illustrates a safe and efficient procedure to modify CD133+ hematopoietic stem cells. The presented non-viral, magnetic polyplex-based approach may provide a basis for the optimization of therapeutic stem cell effects as well as for monitoring the administered cell product via magnetic resonance imaging.

While CD133+ hematopoietic stem cells (SCs) have been proven to provide high potential in the field of regenerative medicine, their low retention rates ...

Light-sheet Fluorescence Microscopy to Capture 4-Dimensional Images of the Effects of Modulating Shear Stress on the Developing Zebrafish Heart

The hemodynamic forces experienced by the heart influence cardiac development, especially trabeculation, which forms a network of branching outgrowths from the myocardium. Genetic program defects in the Notch signaling cascade are involved in ventricular defects such as Left Ventricular Non-Compaction Cardiomyopathy or Hypoplastic Left Heart Syndrome. Using this protocol, it can be determined that shear stress driven trabeculation and Notch signaling are related to one another. Using Light-sheet Fluorescence Microscopy, visualization of the developing zebrafish heart was possible. In this manuscript, it was assessed whether hemodynamic forces modulate the initiation of trabeculation via Notch signaling and thus, influence contractile function occurs. For qualitative and quantitative shear stress analysis, 4-D (3-D+time) images were acquired during zebrafish cardiac morphogenesis, and integrated light-sheet fluorescence microscopy with 4-D synchronization captured the ventricular motion. Blood viscosity was reduced via gata1a-morpholino oligonucleotides (MO) micro-injection to decrease shear stress, thereby, down-regulating Notch signaling and attenuating trabeculation. Co-injection of Nrg1 mRNA with gata1a MO rescued Notch-related genes to restore trabeculation. To confirm shear stress driven Notch signaling influences trabeculation, cardiomyocyte contraction was further arrested via tnnt2a-MO to reduce hemodynamic forces, thereby, down-regulating Notch target genes to develop a non-trabeculated myocardium. Finally, corroboration of the expression patterns of shear stress-responsive Notch genes was conducted by subjecting endothelial cells to pulsatile flow. Thus, the 4-D light-sheet microscopy uncovered hemodynamic forces underlying Notch signaling and trabeculation with clinical relevance to non-compaction cardiomyopathy.

Here, we present a protocol to visualize developing hearts in zebrafish in 4-Dimensions (4-D). 4-D imaging, via light-sheet fluorescence microscopy (LSFM), takes 3-Dimensional (3-D) images over time, to reconstruct developing hearts. We show qualitatively and quantitatively that shear stress activates endocardial Notch signaling during chamber development, which promotes cardiac trabeculation.

The hemodynamic forces experienced by the heart influence cardiac development, especially trabeculation, which forms a network of branching outgrowths ...

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. ...

Intra-cardiac Side-Firing Light Catheter for Monitoring Cellular Metabolism using Transmural Absorbance Spectroscopy of Perfused Mammalian Hearts

Absorbance spectroscopy of cardiac muscle provides non-destructive assessment of cytosolic and mitochondrial oxygenation via myoglobin and cytochrome absorbance respectively. In addition, numerous aspects of the mitochondrial metabolic status such as membrane potential and substrate entry can also be estimated. To perform cardiac wall transmission optical spectroscopy, a commercially available side-firing optical fiber catheter is placed in the left ventricle of the isolated perfused heart as a light source. Light passing through the heart wall is collected with an external optical fiber to perform optical spectroscopy of the heart in near real- time. The transmission approach avoids numerous surface scattering interference occurring in widely used reflection approaches. Changes in transmural absorbance spectra were deconvolved using a library of chromophore reference spectra, providing quantitative measures of all the known cardiac chromophores simultaneously. This spectral deconvolution approach eliminated intrinsic errors that may result from using common dual wavelength methods applied to overlapping absorbance spectra, as well as provided a quantitative evaluation of the goodness of fit. A custom program was designed for data acquisition and analysis, which permitted the investigator to monitor the metabolic state of the preparation during the experiment. These relatively simple additions to the standard heart perfusion system provide a unique insight into the metabolic state of the heart wall in addition to conventional measures of contraction, perfusion, and substrate/oxygen extraction.

Here we introduce a method for using an intra-ventricle optical catheter in perfused hearts to perform absorbance spectroscopy across the heart wall. The data obtained provides robust information on tissue oxygen tension as well as substrate utilization and membrane potential simultaneously with cardiac performance measures in this ubiquitous preparation.

Absorbance spectroscopy of cardiac muscle provides non-destructive assessment of cytosolic and mitochondrial oxygenation via myoglobin and cytochrome ...

Creation of Patient-Specific Silicone Cardiac Models with Applications in Pre-surgical Plans and Hands-on Training

Three dimensional&#160;models can be a valuable tool for surgeons as they develop surgical plans and medical fellows as they learn about complex cases. In particular, 3D models can play an important role in the field of cardiology, where complex congenital heart diseases occur. While many 3D printers can provide anatomically correct and detailed models, existing 3D printing materials fail to replicate myocardial tissue properties and can be extremely costly. This protocol aims to develop a process for the creation of patient-specific models of complex congenital heart defects using a low-cost silicone that more closely matches cardiac muscle properties. With improved model fidelity, actual surgical procedural training could occur in advance of the procedure. Successful creation of cardiac models begins with the segmentation of radiologic images to generate a virtual blood pool (blood that fills the chambers of the heart) and myocardial tissue mold. The blood pool and myocardial mold are 3D printed in acrylonitrile butadiene styrene (ABS), a plastic dissolvable in acetone. The mold is assembled around the blood pool, creating a negative space simulating the myocardium. Silicone with a shore hardness of 2A is poured into the negative space and allowed to cure. The myocardial mold is removed, and the remaining silicone/blood pool model is submerged in acetone. The described process results in a physical model in which all cardiac features, including intra-cardiac defects, are represented with more realistic tissue properties and are more closely approximated than a direct 3D printing approach. The successful surgical correction of a model with a ventricular septal defect (VSD) using a GORE-TEX patch (standard surgical intervention for defect) demonstrates the utility of the method.

Patient-specific models improve surgeon and fellow confidence when developing or learning surgical plans. Three-dimensional (3D) printers generate adequate detail for surgical preparation, but fail to replicate tissue haptic fidelity. A protocol is presented detailing the creation of patient-specific, silicone cardiac models, combining 3D printing precision with simulated silicone tissue.

Three dimensional&#160;models can be a valuable tool for surgeons as they develop surgical plans and medical fellows as they learn about complex cases. In ...

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Creation of Patient-Specific Silicone Cardiac Models with Applications in Pre-surgical Plans and Hands-on Training