The authors would like to express gratitude to Thao Nguyen and Atsushi Nakano from UCLA for providing the mice sample to image. This study was supported by grants NIH HL118650 (to Tzung K. Hsiai), HL083015 (to Tzung K. Hsiai), HD069305 (to N. C. Chi and Tzung K. Hsiai.), HL111437 (to Tzung K. Hsiai and N. C. Chi), HL129727 (to Tzung K. Hsiai), and University of Texas System STARS funding (to Juhyun Lee).

All the procedures involving the use of animals have been approved by the Institutional Review Committees (IACUC) at the University of California, Los Angeles, California.
1. Imaging System Setup
Note: See Figure 1 and Figure 2.
<ol>
	<li>Retrieve a continuous wave (CW) laser with 3 wavelengths: 405 nm, 473 nm, and 532 nm. Place 2 mirrors (M1 and M2) 150 mm apart and align them with their mirror planes at 45&#176; to the beam. 
	Note: This step is performed to redirect the laser away from the dual-sided illumination setup. The resulting beam will be in the same direction as the initial beam.</li>
	<li>Pass the beam through a 25-mm diameter iris diaphragm/pinhole (PH), a 50-mm diameter neutral density filter (NDF) with an optical density range of 0 - 4.0, a beam expander (BE), a 30 mm slit (S) with the width of ~0 - 6 mm, and a mirror (M3), all positioned 150 mm from each other. Place the mirror with its mirror plane at 45&#176; to the beam.</li>
	<li>Pass the beam through a 50:50 beam splitter placed 150 mm from M3. Place a mirror (M6) 150 mm from the beam splitter and align it so that its mirror plane is at 45&#176; to the beam that is emitted in the forward direction. Use the reflected beam to form one side of the dual-illumination light sheet.</li>
	<li>Place a mirror (M4) 150 mm from the beam splitter and align it so that its mirror plane is at 45&#176; to the beam that was emitted at a 90&#176; angle from the beam splitter. Place a second mirror (M5) 150 mm from M4 and align it so that its mirror plane is at 45&#176; to the beam reflected from M6. Use the beam emitted from M5 to form the second side of the dual-illumination light sheet.</li>
	<li>Set up the dual-sided illumination system in a symmetric fashion (see Figure 1). Place one cylindrical lens (CL1) [diameter (d) = 1 in; focal length (f) = 50 mm] 150 mm in line with the beam emitted from M5 on one side and another identical cylindrical lens (CL2) 150 mm in line with the beam emitted from M6 on the other side of the dual-illumination setup. Place 2 mirrors (M7 and M8) each in line with the cylindrical lenses at distances of 50 mm to reflect the beam at 90&#176;.</li>
	<li>Form an achromatic doublet from a pair of lenses. Place the first lens, L1 (d = 1 in; f = 100 mm), 100 mm from M7, and the second lens, L2 (d = 1 in; f = 60 mm), 160 mm from L1. Repeat this on the other side of the dual-illumination with identical lenses (L3, L4) placed the same distance from M8.</li>
	<li>Place mirrors, M9 and M10, 60 mm from lenses L2 and L4, respectively, and in line with the beam. Place the illumination objectives, OL1 and OL2 (d = 2 in; f = 150 mm), 150 mm from M9 and M10 and in line with the beam. The beam emitted from the objectives forms the light sheet for imaging the samples.</li>
	<li>Use a 3-D printer to print the sample holder from acrylonitrile butadiene styrene (ABS). Embed a piece of cover glass [refractive index (RI) = 1.4745] on each side of the chamber, perpendicular to the illumination beam, to minimize refractive index mismatching. Evenly place the chamber in between the beams emitted from the objective lenses.</li>
	<li>Install a stereo microscope with a 1X magnification objective and a scientific complementary metal oxide semiconductor (sCMOS) camera perpendicular to the illumination plane (see Figure 2).</li>
</ol>
2. Imaging System Calibration
<ol>
	<li>Retrieve fluorescent polystyrene beads that are 0.53 &#956;m in diameter.</li>
	<li>Prepare the fluorescent bead sample by diluting the bead solution in a refractive index matching solution (RIMS) with 1% low-melt pointing agarose to 1:150,000. Cut a piece of borosilicate glass tubing with an inner diameter of 12 mm and an outer diameter of 18 mm to a length of 30 mm. 
	Note: The borosilicate glass tubing is used to match the refractive index (1.47).</li>
	<li>Mix the diluted bead sample with 1% agarose and pipette the bead/agarose solution into the borosilicate tubing. Allow the agarose to solidify at room temperature (23 &#176;C).</li>
	<li>Fill the ABS chamber (from step 1.7) with a 99.5% glycerol solution. Place the borosilicate glass tubing containing the beads inside the chamber. Place the ABS chamber in the imaging system so that it is in the center of the Gaussian beam created by the dual illumination system.</li>
	<li>Attach a 3-D motorized translational stage to the borosilicate glass tubing to control the movement and orientation of the sample within the ABS chamber (see Supplemental Figure 1).</li>
	<li>Acquire images using the sCMOS camera at a rate of 30 frames per second (fps). Using the motor controller, move the sample 1 mm in the lateral direction and acquire images at each 1-mm increment using the sCMOS camera. Continue until the entire sample has been imaged.</li>
	<li>Stack the acquired images using a visualization software (see Table of Materials). Measure the point spread function (PSF) of the system using these bead images. Use this PSF for deconvolution during the image processing in later steps.</li>
</ol>
3. Sample Preparation
<ol>
	<li>Dissect hearts from a wild type P1 mouse and from a double heterozygous sarcolipin-Cre knockin mouse with the Rosa26-YFP gene (SlnCre/+; R26YFP reporter/+) at P7. 
	Note: Euthanasia was performed using pentobarbital and the technique described by Robbins et al.10. The dissection was performed according to the technique described by the National Heart, Lung, and Blood Institute (NHLBI)10,11.</li>
	<li>Perform a chemical clearing of the murine hearts as described by Kevin Sung et al.12. 
	Note: The following steps should be performed in a fume hood since the chemicals being used are toxic.
	<ol>
		<li>Rinse the hearts in 1x phosphate-buffered saline (PBS) 3x for 10 min. After rinsing, place the hearts in a 4% paraformaldehyde solution and incubate these samples at 4 &#176;C overnight.</li>
		<li>Place the samples in a 4% acrylamide solution containing 0.5% w/v of 2,2&#8217;-Azobis dihydrochloride. Allow the samples to incubate at 4 &#176;C overnight. After the overnight incubation, remove the samples and incubate them at 37 &#176;C for 2 - 3 h.</li>
		<li>Rinse the samples with PBS and then place them in a solution of 8% w/v sodium dodecyl sulfate and 1.25% w/v boric acid. Incubate the samples at 37 &#176;C until they are clear. This can take several hours depending on the thickness of the sample. After clearing, remove the samples and place them in 1x phosphate-buffered saline for 1 day.</li>
		<li>Prepare a refractive index matching solution with 40 g of nonionic density gradient medium (see Table of Materials) in 30 mL of 0.02 M phosphate buffer (PB), 0.1% Tween-20, and 0.01% sodium azide. Bring the solution to a pH of 7.5 with sodium hydroxide.</li>
	</ol>
	</li>
	<li>Place the cleared sample in RIMS with a refractive index of 1.46 - 1.48 and 1% agarose. Insert the sample into borosilicate glass tubing and allow the agarose to solidify at room temperature (23 &#176;C).</li>
	<li>Attach a 3-D motorized translational stage to the borosilicate glass tubing to control the movement and orientation of the sample within the 3-D printed ABS chamber (see Supplemental Figure 1).</li>
</ol>
4. Adult Heart Imaging
<ol>
	<li>Inject 8.7 x 1012 viruses of type adeno-associated virus vector 9 (AAV9) containing a cardiac-specific troponin T promoter (cTnT) and green fluorescent protein (GFP) in a 100-&#956;L volume into the tail vein of a 2-month wild type (C57BL/6) mouse. 
	Note: The AAV9 was developed according to the technique described by Yuan et al.13. This causes GFP to bind to the renal outer medullary potassium channel (ROMK) producing AAV9-cTnT-ROMK-GFP (see Table of Materials).</li>
	<li>Dissect the adult mouse hearts at 7.5 months of age according to the technique described by the NHLBI11. Prepare the samples using steps 3.1 - 3.3.</li>
	<li>Connect the motor controller to the motor actuator. Attach the actuator to the borosilicate glass tubing to control the movement and orientation of the sample within the 3-D printed ABS chamber.</li>
	<li>Position the sample so that it is in the center of the Gaussian beam created by the dual illumination system. Acquire images using the sCMOS camera at a rate of 100 fps.</li>
	<li>Using the motor controller, move the sample 1 mm in the axial direction and acquire images at each 1-mm increment with the sCMOS camera. Continue until the entire sample has been imaged. 
	Note: The system is controlled by a custom-developed instrument-control software.</li>
	<li>Stack the acquired images using a visualization software (see Table of Materials). Develop 3-D images using these image stacks and the visualization software14. Deconvolve the PSF from step 2.6 with the acquired image stack. Set a pixel threshold intensity value to observe the contours of the heart and add pseudo-color to the images based on this gray-scale intensity.</li>
</ol>

<ol><li>Richardson, D. S., Lichtman, J. W. <a target="_blank" href="https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4537058/">Clarifying tissue clearing.</a> Cell. 162 (2), 246-257 (2015).</li>
<li>Lee, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((4-Dimensional+light-sheet+microscopy+to+elucidate+shear+stress+modulation+of+cardiac+trabeculation%5BTitle%5D)+AND+%22Journal+of+Clinical+Investigation%22%5BJournal%5D)">4-Dimensional light-sheet microscopy to elucidate shear stress modulation of cardiac trabeculation.</a> Journal of Clinical Investigation. 126 (5), 1679-1690 (2016).</li>
<li>Huisken, J., Swoger, J., Del Bene, F., Wittbrodt, J., Stelzer, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Optical+sectioning+deep+inside+live+embryos+by+selective+plane+illumination+microscopy%5BTitle%5D)+AND+%22Science%22%5BJournal%5D)">Optical sectioning deep inside live embryos by selective plane illumination microscopy.</a> Science. 305 (5686), 1007-1009 (2004).</li>
<li>Huisken, J., Stainier, D. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Selective+plane+illumination+microscopy+techniques+in+developmental+biology%5BTitle%5D)+AND+%22Development%22%5BJournal%5D)">Selective plane illumination microscopy techniques in developmental biology.</a> Development. 136 (12), 1963-1975 (2009).</li>
<li>Bakkers, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Zebrafish+as+a+model+to+study+cardiac+development+and+human+cardiac+disease%5BTitle%5D)+AND+%22Cardiovascular+Research%22%5BJournal%5D)">Zebrafish as a model to study cardiac development and human cardiac disease.</a> Cardiovascular Research. 91 (2), 279-288 (2011).</li>
<li>High, F. A., Epstein, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+multifaceted+role+of+Notch+in+cardiac+development+and+disease%5BTitle%5D)+AND+%22Nature+Reviews+Genetics%22%5BJournal%5D)">The multifaceted role of Notch in cardiac development and disease.</a> Nature Reviews Genetics. 9 (1), 49-61 (2008).</li>
<li>Sachinidis, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Cardiac+specific+differentiation+of+mouse+embryonic+stem+cells%5BTitle%5D)+AND+%22Cardiovascular+Research%22%5BJournal%5D)">Cardiac specific differentiation of mouse embryonic stem cells.</a> Cardiovascular Research. 58 (2), 278-291 (2003).</li>
<li>Ding, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Light-sheet+fluorescence+imaging+to+localize+cardiac+lineage+and+protein+distribution%5BTitle%5D)+AND+%22Scientific+Reports%22%5BJournal%5D)">Light-sheet fluorescence imaging to localize cardiac lineage and protein distribution.</a> Scientific Reports. 7, 42209(2017).</li>
<li>Tomer, R., Ye, L., Hsueh, B., Deisseroth, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Advanced+CLARITY+for+rapid+and+high-resolution+imaging+of+intact+tissues%5BTitle%5D)+AND+%22Nature+Protocols%22%5BJournal%5D)">Advanced CLARITY for rapid and high-resolution imaging of intact tissues.</a> Nature Protocols. 9 (7), 1682-1697 (2014).</li>
<li>Robbins, N., Thompson, A., Mann, A., Blomkalns, A. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Isolation+and+excision+of+murine+aorta%3B+a+versatile+technique+in+the+study+of+cardiovascular+disease%5BTitle%5D)+AND+%22Journal+of+Visualized+Experiments%22%5BJournal%5D)">Isolation and excision of murine aorta; a versatile technique in the study of cardiovascular disease.</a> Journal of Visualized Experiments. (93), e52172(2014).</li>
<li>National Heart, L., Blood Institute, <a target="_blank" href="http://community.parentprojectmd.org/profiles/blogs/standard-operating-procedures-for-duchenne-animal-models">Standard Operating Procedures (SOP's) for Duchenne Animal Models</a>. , (2015).</li>
<li>Sung, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Simplified+three-dimensional+tissue+clearing+and+incorporation+of+colorimetric+phenotyping%5BTitle%5D)+AND+%22Scientific+Reports%22%5BJournal%5D)">Simplified three-dimensional tissue clearing and incorporation of colorimetric phenotyping.</a> Scientific Reports. 6, 30736(2016).</li>
<li>Yuan, Z., Qiao, C., Hu, P., Li, J., Xiao, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+versatile+adeno-associated+virus+vector+producer+cell+line+method+for+scalable+vector+production+of+different+serotypes%5BTitle%5D)+AND+%22Human+Gene+Therapy%22%5BJournal%5D)">A versatile adeno-associated virus vector producer cell line method for scalable vector production of different serotypes.</a> Human Gene Therapy. 22 (5), 613-624 (2011).</li>
<li>FEI, <a target="_blank" href="https://www.fei.com/software/amira-user-guide/">Amira User's Guide</a>. , Konrad-Zuse-Zentrum fur Informationstechnik. Berlin. 59-138 (2017).</li>
<li>Gao, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Noninvasive+imaging+of+3D+dynamics+in+thickly+fluorescent+specimens+beyond+the+diffraction+limit%5BTitle%5D)+AND+%22Cell%22%5BJournal%5D)">Noninvasive imaging of 3D dynamics in thickly fluorescent specimens beyond the diffraction limit.</a> Cell. 151 (6), 1370-1385 (2012).</li>
<li>Truong, T. V., Supatto, W., Koos, D. S., Choi, J. M., Fraser, S. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Deep+and+fast+live+imaging+with+two-photon+scanned+light-sheet+microscopy%5BTitle%5D)+AND+%22Nature+Methods%22%5BJournal%5D)">Deep and fast live imaging with two-photon scanned light-sheet microscopy.</a> Nature Methods. 8 (9), 757-760 (2011).</li>
<li>Ding, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Integrating+light-sheet+imaging+with+virtual+reality+to+recapitulate+developmental+cardiac+mechanics%5BTitle%5D)+AND+%22JCI+Insight%22%5BJournal%5D)">Integrating light-sheet imaging with virtual reality to recapitulate developmental cardiac mechanics.</a> JCI Insight. 2 (22), (2017).</li>
</ol>

The authors have nothing to disclose.

The LSFM system and technique described here utilizes a dual-sided illumination beam combined with an optical clearing to image the mouse heart at post-natal and adult stages of its development. A traditional single illumination beam suffers from photon scattering and absorption through thicker and larger tissue samples1,3. The dual-sided beams provide a more even illumination of the sample, thereby minimizing the effect of striping and other artifacts that are o...

Light-sheet fluorescence microscopy was a technique first developed in 1903 and is used today as a method to study gene expression and also to produce 3-D or 4-D models of tissue samples1,2,3. This imaging method uses a thin sheet of light to illuminate a single plane of a sample so that only that plane is captured by the detector. The sample can then be moved in the axial-direction to capture each layer, one section at a time, and render a 3-D model after the post-processing of the acquired images4. However, due to the absorption and scattering of photons, LSFM has been limited to samples that are either a few microns thick or are optically transparent1.
The limitations of LSFM have led to extensive studies of organisms that have tissues that are optically transparent, such as the zebrafish. Studies involving cardiac development and differentiation are often conducted on zebrafish since there are conserved genes between humans and zebrafish5,6. Although these studies have led to advances in cardiac research related to cardiomyopathies6,7, there is still a need to conduct similar research on higher-level organisms such as mammals.
Mammalian cardiac tissue presents a challenge due to the thickness and opacity of the tissue, the absorption due to hemoglobin in red blood cells, and the striping that occurs due to single-sided illumination of the sample under traditional LSFM methods1,8. To compensate for these limitations, we proposed to use dual-sided illumination and a simplified version of the CLARITY technique9 combined with a refractive index matching solution (RIMS). Therefore, this system allows for the imaging of a sample that is greater than 10 x 10 x 10 mm3 while maintaining a good quality resolution in the axial and lateral planes8.
This system was first calibrated using fluorescent beads arranged in different configurations within the glass tubing. Then, the system was used to image post-natal and adult murine hearts. First, the post-natal mouse heart was imaged at 7 days (P7) to reveal the ventricular cavity, the thickness of the ventricular wall, the valve structures, and the presence of trabeculation. Secondly, a study was conducted to identify cells that would differentiate into cardiomyocytes by using a post-natal mouse heart at 1 day (P1) with Cre-labeled cardiomyocytes and yellow fluorescent protein (YFP). Finally, adult mice at 7.5 months were imaged to observe the presence of renal outer medullary potassium (ROMK) channels after gene therapy8.

<table><tbody><tr><td>CW Laser</td><td>Laserglow Technologies</td><td>LMM-GBV1-PF3-00300-05</td><td>Excitation of fluorophores</td></tr><tr><td>Neutral density filter</td><td>Thorlabs</td><td>NDC-50C-4M</td><td>Controls amount of light entering system</td></tr><tr><td>Achromatic beam expander</td><td>Thorlabs</td><td>GBE05-A</td><td>Expands the beam of light</td></tr><tr><td>Mechanical slit</td><td>Thorlabs</td><td>VA100C</td><td>Controls width of beam</td></tr><tr><td>Beam splitter</td><td>Thorlabs</td><td>BS013</td><td>Forms dual-illumination beam</td></tr><tr><td>Stereo microscope with 1X objective lense</td><td>Olympus</td><td>MVX10</td><td>Used for observation of sample</td></tr><tr><td>ORCA-Flash4.0 LT sCMOS camera</td><td>Hamamatsu Photonics</td><td>C11440-42U</td><td>Used to capture Images</td></tr><tr><td>Acrylonitrile butadiene styrene (ABS)</td><td>Stratasys</td><td>uPrint</td><td>Material used to 3-D print a sample holder</td></tr><tr><td>Fluorescent polystyrene beads</td><td>Spherotech Inc</td><td>PP-05-10</td><td>Used for imaging system calibration</td></tr><tr><td>Borosilicate glass tubing</td><td>Corning</td><td>Pyrex 7740</td><td>Tubing for sample embedding</td></tr><tr><td>Glycerol</td><td>Fisher Scientific</td><td>BP229-4</td><td>Fill for sample chamber</td></tr><tr><td>Phosphate-buffered saline</td><td>Fisher Scientific</td><td>BP39920</td><td>Rinse solution for mouse hearts</td></tr><tr><td>Paraformaldehyde</td><td>Electron microscopy sciences</td><td>RT-15700</td><td>First incubation solution</td></tr><tr><td>Acrylamide</td><td>Wako Chemicals</td><td>AAL-107</td><td>Mixed with 2,2&#039;-Azobis dihydrochloride for second incubation solution for mouse hearts</td></tr><tr><td>2,2&#039;-Azobis dihydrochloride</td><td>Wako Chemicals</td><td>VA-044</td><td>Mixed with Acrylamide for second incubation solution for mouse hearts</td></tr><tr><td>Sodium dodecyl sulfate&amp;nbsp;</td><td>Sigma Aldrich</td><td>71725</td><td>Mixed with Boric acid for third incubtion solution for mouse hearts</td></tr><tr><td>Boric acid</td><td>Fischer Scientific</td><td>A74-1</td><td>Mixed with Sodium dodecyl sulfate for third incubtion solution for mouse hearts</td></tr><tr><td>Sigma D2158</td><td>Sigma Aldrich</td><td>D2158</td><td>Mixed with PB, Tween-20, and Sodium azide as a refractive index matching solution</td></tr><tr><td>Tween-20</td><td>Sigma Aldrich</td><td>11332465001</td><td>Mixed with Sigma D2158, PB, and Sodium azide as a refractive index matching solution</td></tr><tr><td>Sodium azide</td><td>Sigma Aldrich</td><td>S2002</td><td>Mixed with Sigma D2158, PB, and Tween-20 as a refractive index matching solution</td></tr><tr><td>Adeno-associated virus vector 9 with a cardiac-specific Troponin T promoter tagged with GFP</td><td>Vector Biolabs</td><td>VB2045</td><td>Expresses GFP when bound to ROMK</td></tr><tr><td>DC Servo Motor Actuator</td><td>Thorlabs</td><td>Z825B</td><td>Used for movement of sample in axial direction within light sheet</td></tr><tr><td>K-Cube Brushed DC Servo Motor Controller</td><td>Thorlabs</td><td>KDC101</td><td>Connects to motor actuator and controls movement of the actuator</td></tr><tr><td>Amira</td><td>FEI Software</td><td>N/A</td><td>Visualization software for producing 2-D and 3-D images</td></tr></tbody></table>

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.

The technique described here used a dual-sided illumination beam combined with an optical clearing of mouse heart tissue samples to achieve a deeper imaging depth and a larger imaging volume with sufficient imaging resolution (Figure 1). To calibrate the system, fluorescent beads were placed inside the glass tubing and within the imaging system. The beads were then imaged and visualized in the x-y plane, y-z plane, and in the x-z plane, to obtain the point sp...

Watch this Scientific Journal Video about Light-sheet Fluorescence Microscopy for the Study of the Murine Heart at JoVE.com

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

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. ...

light-sheet-fluorescence-microscopy-for-the-study-of-the-murine-heart

Research

JoVE Journal

Bioengineering

9.3K Views.  University of California Los Angeles. This study uses a dual-sided illumination light-sheet fluorescence microscopy (LSFM) technique combined with optical clearing to study the murine heart.

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

Analysis of Cardiac Chamber Development During Mouse Embryogenesis Using Whole Mount Epifluorescence

The goal of this protocol is to describe a method for the dissection of mouse embryos and visualization of embryonic mouse ventricular chambers during heart development using ventricular specific fluorescent reporter knock-in mice (MLC-2v-tdTomato mice). Heart development involves a linear heart tube formation, the heart tube looping, and four chamber septation. These complex processes are highly conserved in all vertebrates. The mouse embryonic heart has been widely used for heart developmental studies. However, due to their extremely small size, dissecting mouse embryonic hearts is technically challenging. In addition, visualization of cardiac chamber formation often needs in situ hybridization, beta-galactosidase staining using LacZ reporter mice, or immunostaining of sectioned embryonic hearts. Here, we describe how to dissect mouse embryonic hearts and directly visualize ventricular chamber formation of MLC-2v-tdTomato mice using whole mount epifluorescent microscopy. With this method, it is possible to directly examine heart tube formation and looping, and four chamber formation without further experimental manipulation of mouse embryos. Although the MLC-2v-tdTomato reporter knock-in mouse line is used in this protocol as an example, this protocol can be applied to other heart-specific fluorescent reporter transgenic mouse lines.

We present the protocols to examine mouse heart development using whole mount epifluorescent microscopy on mouse embryos dissected from ventricular specific MLC-2v-tdTomato reporter knock-in mice. This method allows us to directly visualize each stage of the ventricular formation during mouse heart development without labor-intensive histochemical methods.

The goal of this protocol is to describe a method for the dissection of mouse embryos and visualization of embryonic mouse ventricular chambers during ...

Developmental Biology

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 ...

Quantitative Visualization of Leukocyte Infiltrate in a Murine Model of Fulminant Myocarditis by Light Sheet Microscopy

Light-sheet fluorescence microscopy (LSFM), in combination with chemical clearing protocols, has become the gold standard for analyzing fluorescently labelled structures in large biological specimens, and is down to cellular resolution. Meanwhile, the constant refinement of underlying protocols and the enhanced availability of specialized commercial systems enable us to investigate the microstructure of whole mouse organs and even allow for the characterization of cellular behavior in various live-cell imaging approaches. Here, we describe a protocol for the spatial whole-mount visualization and quantification of the CD45+ leukocyte population in inflamed mouse hearts. The method employs a transgenic mouse strain (CD11c.DTR)that has recently been shown to serve as a robust, inducible model for the study of the development of fulminant fatal myocarditis, characterized by lethal cardiac arrhythmias. This protocol includes myocarditis induction, intravital antibody-mediated cell staining, organ preparation, and LSFM with subsequent computer-assisted image post-processing. Although presented as a highly-adapted method for our particular scientific question, the protocol represents the blueprint of an easily adjustable system that can also target completely different fluorescent structures in other organs and even in other species.

Here, we describe a light-sheet microscopy approach to visualize the cardiac CD45+ leukocyte infiltrate in a murine model of aseptic fulminant myocarditis, which is induced by the intratracheal diphtheria toxin treatment of CD11c.DTR mice.

Light-sheet fluorescence microscopy (LSFM), in combination with chemical clearing protocols, has become the gold standard for analyzing fluorescently ...

Medicine

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 and Tissue Clearing for Comprehensive Cardiac Imaging

Light-Sheet Imaging to Reveal Cardiac Structure in Rodent Hearts

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

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

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

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

Rapid Whole-Mount High-Resolution Imaging of Small Animal Vasculature for Quantitative Studies

In small animal models of cardiovascular development and diseases, subject-specific computational simulations of blood flow&#160;enable quantitative assessments of hemodynamic metrics that are difficult to measure experimentally. Computational fluid dynamic simulations shed light on the critical roles of mechanics in cardiovascular function and disease progression. Acquiring high-quality volumetric images of the vessels of interest is central to the accuracy and reproducibility of morphological measurement and flow quantitation results. This study proposes a rapid, cost-effective, and accessible method for whole-mount high-resolution imaging of small animal vasculature using light-sheet fluorescence microscopy. The modified iDISCO+ (immunolabeling-enabled three-dimensional imaging of solvent-cleared organs) light-sheet sample preparation protocol involves (1) labeling vasculature with a fluorescent agent, (2) preserving the sample, and (3) rendering the sample transparent. Unlike classical iDISCO+, which uses immunohistochemical staining, the authors label vascular endothelium with FITC-tagged poly-L-lysine, an affordable non-specific fluorescent dye that is highly resistant to photo-bleaching, in a process termed &#34;endo-painting.&#34; The rapid labeling reduces sample preparation time from approximately four weeks to less than 3 days. Furthermore, the use of minimally hazardous solvent ethyl cinnamate (ECi) as the clearing agent and imaging solution makes the samples safer to handle and compliant with a wider range of imaging facilities. The proposed protocol is applied to obtain highly resolved light-sheet fluorescence microscopy image stacks of the cardiovascular system in chick embryos ranging from day 3 (HH18) to day 8 (HH34). This study further demonstrates the suitability of this method for vascular quantitation through 3D reconstruction and computational hemodynamic modeling of a day 5 (HH 26) chick embryo.

This protocol introduces a rapid method for quantitative whole-mount three-dimensional vascular imaging using light-sheet fluorescence microscopy. The efficacy of the method is demonstrated using the pharyngeal arch artery system of the chick embryo model, with hemodynamic forces quantified via computational fluid dynamics.

In small animal models of cardiovascular development and diseases, subject-specific computational simulations of blood flow&#160;enable quantitative ...

Local Field Fluorescence Microscopy: Imaging Cellular Signals in Intact Hearts

In the heart, molecular signaling studies are usually performed in isolated myocytes. However, many pathological situations such as ischemia and arrhythmias can only be fully understood at the whole organ level. Here, we present the spectroscopic technique of local field fluorescence microscopy (LFFM) that allows the measurement of cellular signals in the intact heart. The technique is based on a combination of a Langendorff perfused heart and optical fibers to record fluorescent signals. LFFM has various applications in the field of cardiovascular physiology to study the heart under normal and pathological conditions. Multiple cardiac variables can be monitored using different fluorescent indicators. These include cytosolic [Ca2+], intra-sarcoplasmic reticulum [Ca2+] and membrane potentials. The exogenous fluorescent probes are excited and the emitted fluorescence detected with three different arrangements of LFFM epifluorescence techniques presented in this paper. The central differences among these techniques are the type of light source used for excitation and on the way the excitation light is modulated. The pulsed LFFM (PLFFM) uses laser light pulses while continuous wave LFFM (CLFFM) uses continuous laser light for excitation. Finally, light-emitting diodes (LEDs) were used as a third light source. This non-coherent arrangement is called pulsed LED fluorescence microscopy (PLEDFM).

In the heart, molecular events coordinate the electrical and contractile function of the organ. A set of local field fluorescence microscopy techniques presented here enables the recording of cellular variables in intact hearts. Identifying mechanisms defining the cardiac function is critical in understanding how the heart works under pathological situations.

In the heart, molecular signaling studies are usually performed in isolated myocytes. However, many pathological situations such as ischemia and ...

Biology

Revealing Murine Pulmonary Veins Myocardial Architecture

Microdissection and Immunofluorescence Staining of Myocardial Sleeves in Murine Pulmonary Veins

Pulmonary veins (PVs) are the major source of ectopic beats in atrial arrhythmias and play a crucial role in the development and progression of atrial fibrillation (AF). PVs contain myocardial sleeves (MS) composed of cardiomyocytes. MS are implicated in the initiation and maintenance of AF, as they preserve similarities to the cardiac working myocardium, including the ability to generate ectopic electrical impulses. Rodents are widely used and may represent excellent animal models to study the pulmonary vein myocardium since cardiomyocytes are widely present all over the vessel wall. However, precise microdissection and preparation of murine PVs is challenging due to the small organ size and intricate anatomy.
We demonstrate a microscopy-guided microdissection protocol for isolating the murine left atrium (LA) together with the PVs. Immunofluorescence staining using cardiac Troponin-T (cTNT) and connexin 43 (Cx43) antibodies is performed to visualize the LA and PVs in full length. Imaging at 10x and 40x magnification provides a comprehensive view of the PV structure as well as detailed insights into the myocardial architecture, particularly highlighting the presence of connexin 43 within the MS.

Author Spotlight: Developing a Translational Model for Atrial Fibrillation Research Across Species 

This protocol demonstrates microscopy-guided isolation and immunofluorescence staining of murine pulmonary veins. We prepare tissue samples containing the left atrium, pulmonary veins, and the corresponding lungs and stain them for cardiac Troponin T and Connexin 43.

Pulmonary veins (PVs) are the major source of ectopic beats in atrial arrhythmias and play a crucial role in the development and progression of atrial ...

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 ...

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 ...