We thank Madelyn Neufeld for the illustration in Figure 2h. This work was supported by the Max Planck Society, Morgridge Institute for Research, the Chan Zuckerberg Initiative, and the Human Frontier Science Program (HFSP).

All zebrafish (Danio rerio) adults and embryos were handled in accordance with protocols approved by the UW-Madison Institutional Animal Care and Use Committee (IACUC).
1. Preparation of zebrafish
<ol>
	<li>Handle zebrafish according to established protocols25,26 and institutional guidelines. Breed adult fish of desired transgenic line (see Discussion). Collect the embryos and keep them at 28 &#176;C in a Petri dish filled with fish medium, e.g., E327.</li>
	<li>Choose a method of immobilization (see Discussion).
	<ol>
		<li>If using &#945;-bungarotoxin mRNA to immobilize the fish, inject 30 pg mRNA22 into the yolk of one- or two-cell stage embryos using a bore glass needle mounted onto a micromanipulator and connected to a picoinjector28.</li>
		<li>If using tricaine, make 0.4% stock solution buffered to pH 7.0-7.4 with 1 M Tris base and store at -20 &#176;C until imaging.</li>
	</ol>
	</li>
	<li>Keep the eggs in an E3 filled Petri dish at 28 &#176;C and transfer the eggs every 24 h to a new dish with fresh E3 until imaging.</li>
	<li>To prevent pigment formation, if the zebrafish background is not albino, transfer fish at 24 h post-fertilization (hpf) to a new E3 dish with 0.2 mM tyrosinase inhibitor 1- phenyl 2-thiourea (see Discussion).</li>
</ol>
2. Preparation of FEP tubes
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62741/62741fig01.jpg" /> 
Figure 1: FEP tube cleaning and straightening. (a) FEP tubes on a cable drum. (b) FEP tubes before straightening. (c) FEP tubes in glass and steel autoclave-safe tubing. (d) Flushing of the FEP tubes after straightening and cool down. (e) FEP tube cut to the size of a centrifuge tube for sonication. (f) Flushing of FEP tubes after sonication. (g) Storage of the cleaned and straightened FEP tubes in a centrifuge tube. (h) Cutting the FEP tube prior to imaging. <a href="https://www.jove.com/files/ftp_upload/62741/62741fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>Straighten the FEP tubes (Figure 1a,b) by placing them in a glass or steel autoclave-safe tubing (Figure 1c) with the correct inner diameter to fit FEP tubes, usually 1.6 or 2.4 mm, and autoclave to 180 &#176;C for 2 h. Let the tubes cool down at room temperature for at least 5 h. Then, remove from the straightening tubes. 
	NOTE: Use gloves when manipulating the tubes and work with 50 cm tubing at a time.</li>
	<li>Clean the FEP tubes. 
	NOTE: Syringes with blunt needle tip of the inner FEP tube size are recommended for safety, but a regular needle will work.
	<ol>
		<li>Flush the tubes with 1 M NaOH twice with a 50 mL syringe (Figure 1d).</li>
		<li>Cut the FEP tubes to the size of a 50 mL centrifuge tube with a razor blade (Figure 1e), place cut tubes in 0.5 M NaOH filled centrifuge tubes, and ultrasonicate them for 10 min.</li>
		<li>Flush the FEP tubes with double-distilled H2O, then repeat flushing with 70% ethanol (Figure 1f).</li>
		<li>Transfer tubes to 70% ethanol and ultrasonicate for 10 min.</li>
		<li>Flush the tubes with double-distilled H2O and store them in centrifuge tubes in double-distilled H2O (Figure 1g).</li>
	</ol>
	</li>
</ol>
3. Preparation of 2% agarose dish
<ol>
	<li>In a glass flask, dissolve low melting point agarose powder in E3. Heat the solution in a microwave and stir it every 20 s, until all powder is dissolved.</li>
	<li>Pour agarose into a glass or plastic Petri dish to make a 1-2 mm coat. Wait until agarose is solidified.</li>
	<li>To store, gently pour E3 on the top of the agar to prevent drying. Wrap in paraffin film and keep at 4 &#176;C.</li>
</ol>
4. Preparation of embedding media
<ol>
	<li>Prepare enough E3 to fill the sample chamber. 
	NOTE: Avoid using methyl blue if media is in contact with objective lenses.</li>
	<li>If using tricaine, thaw stock solution and add 0.02% tricaine to E3.</li>
</ol>
5. Sample mounting
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62741/62741fig02.jpg" /> 
Figure 2: Embryo mounting in FEP tube. (a) Anesthetized pigment-free fish in mounting media. (b) A syringe with blunt end needle and FEP tube attached. (c) Once media and fish are taken up in the FEP tube, cut the tube at the edge of the needle. (d) Dipping the cut tube into a dish coated with 2% agarose to plug its end. (e) A zebrafish in a plugged FEP tube. (f) Gently cut the FEP tube at 30&#176; to create gas-exchange holes. (g) FEP tube with four holes above an embedded zebrafish. (h) Scheme of a zebrafish embedded in an FEP tube. Holes and agar plug are indicated. (i) Multiple embedded zebrafish ready for imaging. <a href="https://www.jove.com/files/ftp_upload/62741/62741fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>With a disposable glass pipette, transfer fish to embedding media (Figure 2a). If using tricaine, transfer fish to Petri dish filled with tricaine-containing E3, 10 min before imaging. In both cases, view under a stereomicroscope to verify that the fish stopped moving and that the heart is beating at the similar speed when compared to the control.</li>
	<li>Cut the FEP tube to the ideal length with a razor blade (Figure 1h). 
	NOTE: The length should be adjusted to the microscope&#39;s sample holder; the typical length is about 3 cm.</li>
	<li>Prepare a syringe with a blunt end cannula. Fill the syringe with air, then mount the FEP tube onto the needle and gently flush out any remaining water by emptying the syringe (Figure 2b). 
	NOTE: Avoid making bubbles by slowly flushing out the air.</li>
	<li>With the syringe mounted FEP tube, first, take up media to fill the FEP tube, then take up an embryo head down. Keep the fish head as close to the tube end as possible. Avoid making any bubbles; if a bubble is present, discard the sample.</li>
	<li>With a razor blade, carefully cut the FEP tube at the edge of the blunt end cannula or needle (Figure 2c).</li>
	<li>Discard any liquid on the top of the agar-coated dish. Plunge the FEP tube straight into the agar (Figure 2d). Rotate the tube and take it out to release the plug from the agarose bed.</li>
	<li>Under a stereoscope, verify the presence of the agar plug at the end of the tube (Figure 2e).</li>
	<li>For long-term imaging, cut 3-5 holes into the FEP tube at each cardinal direction, at least 5 mm above the end of the fish.
	<ol>
		<li>Under a stereoscope, use a razor blade perpendicular to the axis of the tube to make a 30&#176; incision into the FEP tube until reaching the mounting media (Figure 2f).</li>
		<li>Make a second cut at 180&#176; to create a hole (Figure 2g,h).</li>
	</ol>
	</li>
	<li>Transfer mounted embryo head down into a 1.5 mL microcentrifuge tube with embedding media until ready to image (Figure 2i).</li>
</ol>
6. Sample positioning
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62741/62741fig03.jpg" /> 
Figure 3: Sample chamber. (a) FEP tube mounted on a sample holder. (b) The sample chamber with stages and objectives. (c) Top view of the media-filled sample chamber, with illumination and detection objective in a T-SPIM configuration. (d) Sample holder mounted on the microscope, with the sample in the chamber. <a href="https://www.jove.com/files/ftp_upload/62741/62741fig03large.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/62741/62741fig04.jpg" /> 
Figure 4: Embryo positioning for heart imaging. (a) 24 hpf Tg(kdrl:Hsa.HRAS-mCherry) zebrafish with eyes misaligned. (a&#39;) Same fish, with eyes aligned. (b) Same fish rotated -100 &#176; and (b&#39;) -50 &#176; for optimal heart imaging. (c) 48 hpf zebrafish with eyes misaligned. (c&#39;) Same fish, with eyes aligned. (d) Same fish rotated by 30&#176; for optimal heart imaging. Black arrows point to heart. Scale bar 100 &#181;m. <a href="https://www.jove.com/files/ftp_upload/62741/62741fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>At the microscope, mount the FEP tube in the sample holder (Figure 3a) and fill the imaging chamber with embedding media (Figure 3b,c). Next, place the sample holder on the stage with the sample dipping into the chamber (Figure 3d).</li>
	<li>Check the sample&#39;s health. Visually assess heart rate to evaluate overall fish wellness, as specific heart rate is stage and temperature dependent, by comparing to non-mounted control fish. If the heartbeat is too slow, discard the fish. 
	NOTE: Ensure gentle handling of embryos, careful transfer to embedding media, imaging immediately after embedding, avoiding rapid temperature changes, avoiding tricaine, and lowering the exposure time to tricaine.</li>
	<li>For reproducible imaging, always use the same sample position. Aligning the eyes and imaging at an angle is recommended.</li>
	<li>Rotate the fish so that both eyes (Figure 4a,c) are in the focal plane (Figure 4a&#39;,c&#39;)</li>
	<li>From that position, further rotate the fish approximately 50 &#176;-100 &#176; clockwise for 24 hpf imaging (Figure 4b, b&#39;), and approximately 20&#176;-30&#176; counterclockwise for 48 hpf imaging (Figure 4d). 
	&#8203;NOTE: The early heart, before 30 hpf, can be difficult to image due to its hidden position (Figure 4b).</li>
</ol>
7. Image acquisition
<ol>
	<li>Choose the illumination side that gives the best image quality and adapt the laser power to every fish. 
	NOTE: Record the laser power used for subsequent image analysis.</li>
	<li>At each z-plane, record 4-5 heartbeats at 300 frames per second (fps) or more. 
	&#8203;NOTE: The field of view can be cropped to increase acquisition speed. For example, at 48 hpf the zebrafish heart beats two to three times per second, therefore, at 300 fps, between 300 and 600 frames are required to acquire four to six heartbeats.</li>
	<li>To record the beating heart, move the sample stepwise through the light sheet. Use a z-spacing of 1-2 &#181;m, covering the entire depth of the heart.</li>
</ol>
8. Image processing
<ol>
	<li>Synchronize recorded movie to reconstruct a 4D (x,y,z, time) heart using a Fiji (Image J229,30) plugin as previously described6.</li>
	<li>To explore data and generate movies of the rendered zebrafish heart, load the 4D file (x,y,z, time) into a 3D rendering software.</li>
</ol>

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Y., Lamperti, P., Steed, E., Boselli, F., Vermot, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=following+endocardial+tissue+movements+via+cell+photoconversion+in+the+zebrafish+embryo.">following endocardial tissue movements via cell photoconversion in the zebrafish embryo.</a> Journal of Visualized Experiments. (132), e57290 (2018).</li><li>Nagel, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Channelrhodopsin-2,+a+directly+light-gated+cation-selective+membrane+channel.">Channelrhodopsin-2, a directly light-gated cation-selective membrane channel.</a> Proccedings of the National Academy of Sciences. 100 (24), 13940-13945 (2003).</li><li>Knollmann, B. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pacing+lightly:+optogenetics+gets+to+the+heart.">Pacing lightly: optogenetics gets to the heart.</a> Nature Methods. 7 (11), 889-891 (2010).</li><li>Bruegmann, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optogenetic+control+of+heart+muscle+in+vitro+and+in+vivo.">Optogenetic control of heart muscle in vitro and in vivo.</a> Nature Methods. 7 (11), 897-900 (2010).</li></ol>

The authors have nothing to disclose.

Transgenic lines to image the heart
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/62741/62741fig06.jpg" /> 
Figure 6: Comparison of cytoplasmic- and membrane-marker zebrafish transgenic lines. Anterior-ventral view of 48 hpf zebrafish hearts imaged with LSFM. White arrows indicate structures visible only with a membrane-marker transgenic line. (a) Tg(kdrl:EGFP)32 signal in cyan in the heart and (a&#39;) in the ventricle. (b) Tg(kdrl:Hsa.HRAS-mCherry; myl7:dsRed)33 signal in red in the heart and (b&#39;) in the ventricle.( c,c&#39;) merge of both Tg(kdrl:Hsa.HRAS-mCherry; myl7:dsRed) and Tg(kdrl:EGFP) signal. Scale bar 50 &#181;m. <a href="https://www.jove.com/files/ftp_upload/62741/62741fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Imaging the zebrafish heart requires precise heart-cell labeling. While the myocardial thickness is relatively constant throughout the cells, endocardial cells are thick around the nucleus but have thin membrane protrusions, in some regions thinner than 2 &#181;m. Cytoplasmic transgenic lines such as Tg(kdrl:EGFP)32 effectively label the regions around endocardial nuclei, but further away, the thin cytoplasm might not emit enough photons to be detected with such short exposure times, leading to artificial holes in the data (Figure 6a). In contrast, membrane marker transgenic lines such as Tg(kdrl:Hsa.HRAS-mCherry)33 can effectively label the endocardium and reveal more details (Figure 6b,c). For each experiment, carefully choose the most appropriate transgenic line.
Zebrafish immobilization 
The choice of immobilization technique depends on the length of the experiment and the age of the fish to image. Tricaine has commonly been used for zebrafish immobilization, mostly due to its ease of use. Indeed, simply adding 130 mg/L tricaine to the fish media results in their anesthetization in 10 min. As it can lead to developmental defects and affect heart physiology20,22, we recommend using tricaine only for short experiments (less than 30 min). For longer imaging, &#945;-bungarotoxin mRNA injections at the one- or two-cell stage paralyzes fish up to 3 days post fertilization (dpf) without affecting cardiovascular development or physiology22.
Choosing the right FEP tubes 
FEP tubes are available in various diameters and thicknesses. To image 0-5 dpf fish, 0.8 mm is a good inner diameter; choose either thick wall 0.8 x 1.6 mm tubes or thin wall 0.8 x 1.2 mm tubes. We recommend thin-walled tubes; however, thicker walls offer increased stability and rigidity, which can be important if the sample chamber has flowing media that could disrupt and move a thin tube. For larger samples, 1.6 x 2.4 mm and 2 x 3 mm can be used.
Temperature and gas exchanges 
An essential aspect of the zebrafish embryo&#39;s well-being is temperature. Ideally, keep the fish at 28.5 &#176;C while imaging, as the environment&#39;s temperature affects development and heart rate34.
In our experience, oxygen exchange through the 2% agarose plug only maintains a stable heart rate until 3-4 dpf. Therefore, cutting holes in the tube ensures oxygen diffusion. It can also be necessary for drug delivery to the sample if desired.
Suspension of heartbeat. 
The fast acquisition speeds of appropriately equipped light sheet microscopes allow recording of the beating heart in vivo. However, to acquire an undisturbed z-stack, one can slow down or stop the heart. However, stopping the heart leads to heart muscle relaxation and might result in the collapse of the heart6. Heartbeat suspension can be done by using morpholinos, low temperatures, an inhibitor of muscle contraction or optogenetics. These methods each have their drawbacks and must be carefully evaluated for every experiment.
The injection of 4 ng of silent heart (sih) morpholino at the one cell stage can stop the heartbeat by targeting the gene tnnt2a crucial for sarcomere formation35. sih zebrafish do not have a heartbeat and only survive until 7 dpf, when the embryos start to rely on circulating blood for oxygenation. As heart morphogenesis is driven by both genetic and biomechanical forces36, these fish present heart malformations around 3 dpf.
As the flow of Ca2+ is temperature sensitive, temperature influences heart rate in embryonic zebrafish21. Consequently, lowering the temperature in the imaging chamber slows down the heartbeat. Stopping the heartbeat requires temperatures below 15 &#176;C. As zebrafish are usually kept at 28.5 &#176;C, such low temperatures can only be maintained for brief periods (less than 10 min).
Drugs such as chemical inhibitors of muscle contractions, 2,3-Bu-tanedione 2-monoxime (BDM), can be added to the zebrafish media (50 nM37,38) to suspend the heartbeat temporarily. BDM is convenient to use as it stops heart contraction in under 15 minutes and can be washed away to restore cardiac function. However, as BDM alters the cardiac action potential, it must be used with a caution37.
Finally, the heart of transgenic zebrafish expressing light-gated ion channels or pumps such as channelrhodopsin or halorhodopsin in their myocardium can be manipulated and stopped by illuminating the pacemaker at the inflow tract with light39,7,40,41,9.
Outlook 
The presented optimized tools and solutions to study the zebrafish heart in vivo allow long term, gentle imaging of ultrafast cardiac dynamics. The sample embedding can be adapted to suit different imaging modalities, such as confocal microscopy, two-photon microscopy, or optical projection tomography (OPT). Light sheet microscopy, however, is likely the preferred technique that offers optical sectioning at a speed sufficient to capture the dynamics of the heart. While this protocol focuses on zebrafish embryonic heart imaging, we believe that it could also be applied to various other samples and experiments. It will be interesting to see in the future if similar embedding and imaging techniques can also be used at later stages during development when the heart is more hidden and the larva less translucent.

To uncover the complex events and interactions in the early beating heart, in vivo imaging of the whole organ is required. With its minimal phototoxicity1,2,3, low photobleaching4, and high speed, light sheet microscopy has evolved as the primary imaging tool for embryonic and cardiac development5,6. It has delivered insights into cardiac form and function at a high spatial and temporal resolution7,8,9 and has allowed researchers to image and track fluorescently labeled parts of the heart at high speed, study hemodynamic forces, and follow the heart development directly inside the body of developing embryos6,10,11,12.
To precisely and reproducibly constrain zebrafish in the field of view, a variety of embedding protocols for light sheet exist, for the short and long term, as well as single or multi-sample13,14,15,16,17,18,19. The most common protocol involves tricaine immobilization and agarose mounting inside a glass or plastic tube. However, as the heart rate can change due to the temperature, anesthetics, and embedding material used20,21,22, zebrafish cardiac imaging requires specific, gentle protocols to ensure sample health6,8,11,12,20,21,22,23. For short-term imaging (up to an hour), one can anesthetize the fish in 130 mg/L tricaine and embed it in Fluorinated Ethylene Propylene (FEP) tubes with 0.1% agarose solution and a plug, as described in Weber et al. 201416. However, as tricaine can lead to developmental defects20,22, different protocols must be used for long-term imaging.
Here we describe a new strategy for long-term cardiac imaging. While many light sheet implementations exist24, we recommend using a hanging sample in a T-SPIM microscope (one detection and two illumination lenses in a horizontal plane with the sample hanging vertically in the common focus). This gives the necessary freedom of movement and rotation for the precise sample positioning. The fish are immobilized by injecting 30 pg &#945;-bungarotoxin mRNA at the one- or two-cell stage. &#945;-bungarotoxin is a snake venom that paralyzes muscles without affecting cardiovascular development or physiology22. For precise, distortion-free imaging, we recommend mounting fish in tubes made of FEP, a polymer with a refractive index almost identical to water. We discuss how to best prepare the FEP tubes by straightening and cleaning them prior to imaging. The fish are then mounted in these tubes, head down, in media, and the bottom of the tube is sealed with a 2% agarose plug, on which fish heads rest. Cutting holes in the FEP tube facilitates gas exchange and ensures fish growth. The embedded fish can be kept in media until mounted onto a sample holder right before imaging. We also suggest a data acquisition and analysis pipeline for reproducible high-speed imaging. Further, we discuss the use of cytoplasmic versus membrane marker transgenic lines for robust heart cell labeling, as well as different options to stop the heart. These mounting techniques ensure sample health while allowing to constrain the heart precisely and reproducibly in the field of view.

<table><tbody><tr><td>1.5mL Eppendorf</td><td>Eppendorf</td><td>22364111</td><td>To carry embedded samples</td></tr><tr><td>15mL Falcon tubes</td><td>Falcon</td><td>352095</td><td>To carry embedded samples</td></tr><tr><td>50mL centrifuge tubes</td><td>Falcon</td><td>352070</td><td>50ml tubes for sonication step, and storing cleaned, straightened FEP tubes</td></tr><tr><td>50mL syringe</td><td>BD</td><td>309654</td><td>50ml syringe for FEP&amp;nbsp; cleaning</td></tr><tr><td>Agarose, low gelling temperature</td><td>Sigma Aldrich</td><td>39346-81-1</td><td>To make plug</td></tr><tr><td>Blunt Tip Needles, 21 gauge</td><td>VWR</td><td>89500-304</td><td>Blunt end needle for 0.8 inner diameter FEP tube</td></tr><tr><td>Borosilicate glass tube</td><td>McMaster-Carr</td><td>8729K33</td><td>Tubing for FEP tube straightening 9.5mm outer diameter, 5.6 inner diameter, 30cm long, other sizes available</td></tr><tr><td>Borosilicate glass tube</td><td>McMaster-Carr</td><td>8729K31</td><td>Tubing for FEP tube straightening 6.35mm outer diameter, 4mm inner diameter, 30cm long, other sizes available</td></tr><tr><td>Conventional needles, 21 Gauge</td><td>BD</td><td>305165</td><td>Conventional needle for 0.8 inner diameter FEP tube</td></tr><tr><td>Disposable glass pipette</td><td>Grainger</td><td>52NK56</td><td>To transfer fish, use with pipette pump</td></tr><tr><td>E3 medium for zebrafish embryos</td><td></td><td></td><td></td></tr><tr><td>Ethanol</td><td>Sigma Aldrich</td><td>64-17-5</td><td>Ethanol for FEP cleaning</td></tr><tr><td>FEP tube, 0.8 / 1.2 mm</td><td>ProLiquid</td><td>2001048</td><td>FEP tube with thin wall, other sizes available</td></tr><tr><td>FEP tube, 0.8 / 1.6 mm</td><td>Bola</td><td>S1815-04</td><td>FEP tube with thick wall, other sizes available</td></tr><tr><td>FEP tube, 2/3mm</td><td>BGB</td><td>211581</td><td>Large FEP tube with thick wall, other sizes available</td></tr><tr><td>Hot Hand Rubber Mitt</td><td>Cole-Parmer</td><td>691000</td><td>To carry hot equipment after autoclaving</td></tr><tr><td>Omnifix 1mL Syringes</td><td>B Braun</td><td>9161406V</td><td>1ml syringe for embedding</td></tr><tr><td>Petri dish, small</td><td>Dot Scientific</td><td>PD-94050</td><td>To make agarose plug</td></tr><tr><td>Pipette pumps</td><td>Argos Technologies</td><td>04395-05</td><td>To transfer fish, use with disposable glass pipette</td></tr><tr><td>PTU</td><td>Sigma Aldrich</td><td>103-85-5</td><td>Also known as:&lt;br/&gt; N-Phenylthiourea, 1-Phenyl-2-thiourea, Phenylthiocarbamide</td></tr><tr><td>Razor blades</td><td>Azpack</td><td>11904325</td><td>To cut FEP tubes</td></tr><tr><td>Sodium Hydroxide</td><td>Dot Scientific</td><td>DSS24000</td><td>NaOH for FEP cleaning</td></tr><tr><td>Tricaine</td><td>Sigma Aldrich</td><td>E10521</td><td>Also known as: MS-222,&lt;br/&gt; Ethyl 3-aminobenzoate methanesulfonate, Tricaine</td></tr></tbody></table>

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.

<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/62741/62741fig05.jpg" /> 
Figure 5: The 48 hpf zebrafish heart. (a) Still of one z-frame, anterior-ventral view of 48 hpf Tg(kdrl:Hsa.HRAS-mCherry; myl7:lck-EGFP) zebrafish, imaged with LSFM, (b) 3D reconstruction of movie stacks, cut view through the atrium. (c) Montage of four frames over a full heartbeat at one z-plane. Pie charts indicate the time during heartbeat. Scale bar 50 &#181;m. <a href="https://www.jove.com/files/ftp_upload/62741/62741fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
We have recorded the 48 hpf beating heart of Tg(kdrl:Hsa.HRAS-mCherry; myl7:lck-EGFP) zebrafish according to the protocol detailed above (Figure 5). A 488 nm and a 561 nm laser light sheet illuminated the sample simultaneously. The emitted fluorescence was detected perpendicularly using a 16x/0.8 W objective lens and a scientific metal oxide semiconductor (sCMOS) camera.
At 48 hpf, the heart has just undergone looping and has two chambers, the ventricle and the atrium but has yet to develop valves. In our movies, the different heart structures such as inflow tract, ventricle, atrioventricular canal (AVC), atrium, and outflow track are easily distinguishable (Figure 5a,b). These data show the precise beating and reveal complexinteractions between the heart&#39;s two cell layers: the myocardium, a single-cell muscle layer contracting and generatingforce (Figure 5c, red), and the endocardium, a single cell layer that connects the heart to the vasculature (Figure 5c, cyan).
The heartbeat reconstruction in x,y,z (3D) + time (4D) + color (5D) was performed according to Mickoleit et al.6. The reconstruction is based on two hypotheses: the motion of the heart is repetitive, and data should be acquired with a small z-step. The output is a reconstructed single heartbeat in 5D, measuring 30 GB to 80 GB per heartbeat. To render the data, we used the free, open-source tool FluoRender for in depth rendering31 as it was designed to handle multidimensional datasets and easily renders 5D movies of both cell layers and individual layers (Figure 5b).

Watch this Scientific Journal Video about Light Sheet Microscopy of Fast Cardiac Dynamics in Zebrafish Embryos at JoVE.com

Light Sheet Microscopy of Fast Cardiac Dynamics in Zebrafish Embryos

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 ...

light-sheet-microscopy-of-fast-cardiac-dynamics-in-zebrafish-embryos

Universidad San Sebastian

Research

JoVE Journal

Biology

4.0K Views.  Morgridge Institute for Research, Madison, WI. 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.

Video: Light Sheet Microscopy of Fast Cardiac Dynamics in Zebrafish Embryos

Imaging Subcellular Structures in the Living Zebrafish Embryo

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such imaging experiments in higher vertebrates such as mammals generally requires surgical access to the system under study. The optical accessibility of embryonic and larval zebrafish allows such invasive procedures to be circumvented and permits imaging in the intact organism. Indeed the zebrafish is now a well-established model to visualize dynamic cellular behaviors using in vivo microscopy in a wide range of developmental contexts from proliferation to migration and differentiation. A more recent development is the increasing use of zebrafish to study subcellular events including mitochondrial trafficking and centrosome dynamics. The relative ease with which these subcellular structures can be genetically labeled by fluorescent proteins and the use of light microscopy techniques to image them is transforming the zebrafish into an in vivo model of cell biology. Here we describe methods to generate genetic constructs that fluorescently label organelles, highlighting mitochondria and centrosomes as specific examples. We use the bipartite Gal4-UAS system in multiple configurations to restrict expression to specific cell-types and provide protocols to generate transiently expressing and stable transgenic fish. Finally, we provide guidelines for choosing light microscopy methods that are most suitable for imaging subcellular dynamics.

Imaging the dynamic behavior of organelles and other subcellular structures in vivo can shed light on their function in physiological and disease conditions. Here, we present methods for genetically tagging two organelles, centrosomes and mitochondria, and imaging their dynamics in living zebrafish embryos using wide-field and confocal microscopy.

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such ...

Developmental Biology

Using Light Sheet Fluorescence Microscopy to Image Zebrafish Eye Development

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM compared to confocal systems are its low phototoxicity, gentle mounting strategies, fast acquisition with high signal to noise ratio and the possibility of imaging samples from various angles (views) for long periods of time. Imaging from multiple views unleashes the full potential of LSFM, but at the same time it can create terabyte-sized datasets. Processing such datasets is the biggest challenge of using LSFM. In this protocol we outline some solutions to this problem. Until recently, LSFM was mostly performed in laboratories that had the expertise to build and operate their own light sheet microscopes. However, in the last three years several commercial implementations of LSFM became available, which are multipurpose and easy to use for any developmental biologist. This article is primarily directed to those researchers, who are not LSFM technology developers, but want to employ LSFM as a tool to answer specific developmental biology questions. 
Here, we use imaging of zebrafish eye development as an example to introduce the reader to LSFM technology and we demonstrate applications of LSFM across multiple spatial and temporal scales. This article describes a complete experimental protocol starting with the mounting of zebrafish embryos for LSFM. We then outline the options for imaging using the commercially available light sheet microscope. Importantly, we also explain a pipeline for subsequent registration and fusion of multiview datasets using an open source solution implemented as a Fiji plugin. While this protocol focuses on imaging the developing zebrafish eye and processing data from a particular imaging setup, most of the insights and troubleshooting suggestions presented here are of general use and the protocol can be adapted to a variety of light sheet microscopy experiments.

Light sheet fluorescence microscopy is an excellent tool for imaging embryonic development. It allows recording of long time-lapse movies of live embryos in near physiological conditions. We demonstrate its application for imaging zebrafish eye development across wide spatio-temporal scales and present a pipeline for fusion and deconvolution of multiview datasets.

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM ...

Following Endocardial Tissue Movements via Cell Photoconversion in the Zebrafish Embryo

During embryogenesis, cells undergo dynamic changes in cell behavior, and deciphering the cellular logic behind these changes is a fundamental goal in the field of developmental biology. The discovery and development of photoconvertible proteins have greatly aided our understanding of these dynamic changes by providing a method to optically highlight cells and tissues. However, while photoconversion, time-lapse microscopy, and subsequent image analysis have proven to be very successful in uncovering cellular dynamics in organs such as the brain or the eye, this approach is generally not used in the developing heart due to challenges posed by the rapid movement of the heart during the cardiac cycle. This protocol consists of two parts. The first part describes a method for photoconverting and subsequently tracking endocardial cells (EdCs) during zebrafish atrioventricular canal (AVC) and atrioventricular heart valve development. The method involves temporally stopping the heart with a drug in order for accurate photoconversion to take place. Hearts are allowed to resume beating upon removal of the drug and embryonic development continues normally until the heart is stopped again for high-resolution imaging of photoconverted EdCs at a later developmental time point. The second part of the protocol describes an image analysis method to quantify the length of a photoconverted or non-photoconverted region in the AVC in young embryos by mapping the fluorescent signal from the three-dimensional structure onto a two-dimensional map. Together, the two parts of the protocol allows one to examine the origin and behavior of cells that make up the zebrafish AVC and atrioventricular heart valve, and can potentially be applied for studying mutants, morphants, or embryos that have been treated with reagents that disrupt AVC and/or valve development.

This protocol describes a method for the photoconversion of Kaede fluorescent protein in endocardial cells of the living zebrafish embryo that enables the tracking of endocardial cells during atrioventricular canal and atrioventricular heart valve development.

During embryogenesis, cells undergo dynamic changes in cell behavior, and deciphering the cellular logic behind these changes is a fundamental goal in the ...

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

The morphogenesis of the Drosophila embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell shape changes during embryonic development. One of the challenges faced in studying this structure is that the lumen of the heart tube, as well as the membrane features that are crucial to heart tube formation, are difficult to visualize in whole mount embryos, due to the small size of the heart tube and intra-lumenal space relative to the embryo. The use of transmission electron microscopy allows for higher magnification of these structures and gives the advantage of examining the embryos in cross section, which easily reveals the size and shape of the lumen. In this video, we detail the process for reliable fixation, embedding, and sectioning of late stage Drosophila embryos in order to visualize the heart tube lumen as well as important cellular structures including cell-cell junctions and the basement membrane.

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

We describe a process for fixation, embedding, sectioning, and imaging of late stage Drosophila embryos for Trasmission Electron Microscopy of the embryonic heart tube. This technique allows for the visualization of the heart tube lumen as well as the basement membrane, which lines the lumen of the heart.

The morphogenesis of the Drosophila  embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell ...

Two-Photon-Based Photoactivation in Live Zebrafish Embryos

Photoactivation of target compounds in a living organism has proven a valuable approach to investigate various biological processes such as embryonic development, cellular signaling and adult physiology. In this respect, the use of multi-photon microscopy enables quantitative photoactivation of a given light responsive agent in deep tissues at a single cell resolution. As zebrafish embryos are optically transparent, their development can be monitored in vivo. These traits make the zebrafish a perfect model organism for controlling the activity of a variety of chemical agents and proteins by focused light. Here we describe the use of two-photon microscopy to induce the activation of chemically caged fluorescein, which in turn allows us to follow cell's destiny in live zebrafish embryos. We use embryos expressing a live genetic landmark (GFP) to locate and precisely target any cells of interest. This procedure can be similarly used for precise light induced activation of proteins, hormones, small molecules and other caged compounds.

Multiphoton microscopy allows control of low energy photons with deep optical penetration and reduced phototoxicity. We describe the use of this technology for live cell labeling in zebrafish embryos. This protocol can be readily adapted for photo-induction of various light-responsive molecules.

Photoactivation of target compounds in a living organism has proven a valuable approach to investigate various biological processes such as embryonic ...

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 ...

Bioengineering

Refined Multiview Light Sheet Microscopy Protocol for Zebrafish Embryos

Light Sheet Microscopy Imaging and Mounting Strategies for Early Zebrafish Embryos

Light sheet microscopy has become the methodology of choice for live imaging of zebrafish embryos over long time scales with minimal phototoxicity. In particular, a multiview system, which allows sample rotation, enables imaging of entire embryos from different angles. However, in most imaging sessions with a multiview system, sample mounting is a troublesome process as samples are usually prepared in a polymer tube. To aid in this process, this protocol describes basic mounting strategies for imaging early zebrafish development between the 70% epiboly and early somite stages. Specifically, the study provides statistics on the various positions the embryos default to at the 70% epiboly and bud stages within the chorion. Furthermore, it discusses the optimum number of angles and the interval between angles required for imaging whole zebrafish embryos at the early stages of development so that cellular-scale information can be extracted by fusing the different views. Finally, since the embryo covers the entire field of view of the camera, which is required to obtain a cellular-scale resolution, this protocol details the process of using bead information from above or below the embryo for the registration of the different views.

Author Spotlight: Strategies for Mounting Zebrafish Embryos for High-Resolution Multiview Light-Sheet Microscopy &#8212; Techniques for Imaging and Image Reconstruction

A sample preparation strategy for imaging early zebrafish embryos within an intact chorion using a light-sheet microscope is described. It analyzes the different orientations that embryos acquire within the chorion at the 70% epiboly and bud stages and details imaging strategies for obtaining cellular-scale resolution throughout the embryo using the light-sheet system.

Light sheet microscopy has become the methodology of choice for live imaging of zebrafish embryos over long time scales with minimal phototoxicity. In ...

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 ...

Multilayer Mounting for Long-term Light Sheet Microscopy of Zebrafish

Light sheet microscopy is the ideal imaging technique to study zebrafish embryonic development. Due to minimal photo-toxicity and bleaching, it is particularly suited for long-term time-lapse imaging over many hours up to several days. However, an appropriate sample mounting strategy is needed that offers both confinement and normal development of the sample. Multilayer mounting, a new embedding technique using low-concentration agarose in optically clear tubes, now overcomes this limitation and unleashes the full potential of light sheet microscopy for real-time developmental biology.

The development of zebrafish can be followed over days with light sheet microscopy when embryos are embedded in optically clear polymer tubes with low-concentration agarose.

Light sheet microscopy is the ideal imaging technique to study zebrafish embryonic development. Due to minimal photo-toxicity and bleaching, it is ...

Light Sheet Microscopy Sample Preparation: Mounting Live Zebrafish Embryos for Long-Term Imaging

Source: Icha J., et al. <a href="https://www.jove.com/v/53966/" target="_blank">Using Light Sheet Fluorescence Microscopy to Image Zebrafish Eye Development</a>. J. Vis. Exp. (2016).
This article describes sample preparation for light-sheet fluorescence microscopy. It also demonstrates a method to mount live zebrafish embryos for long term imaging, which allows recording of long time-lapse movies of live embryos in near-physiological conditions.

Source: Icha J., et al. Using Light Sheet Fluorescence Microscopy to Image Zebrafish Eye Development. J. Vis. Exp. (2016).
This article describes sample ...

Light Sheet Microscopy of Fast Cardiac Dynamics in Zebrafish Embryos

Summary

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Chapters in this video

Imaging Subcellular Structures in the Living Zebrafish Embryo

Using Light Sheet Fluorescence Microscopy to Image Zebrafish Eye Development

Following Endocardial Tissue Movements via Cell Photoconversion in the Zebrafish Embryo

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

Two-Photon-Based Photoactivation in Live Zebrafish Embryos

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

Light Sheet Microscopy Imaging and Mounting Strategies for Early Zebrafish Embryos

4D Light-sheet Imaging of Zebrafish Cardiac Contraction

Multilayer Mounting for Long-term Light Sheet Microscopy of Zebrafish

Light Sheet Microscopy Sample Preparation: Mounting Live Zebrafish Embryos for Long-Term Imaging