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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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&#160; 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 colspan="2">E3 medium for zebrafish embryos</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 thick 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 thin 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: 
			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, 
			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

Research

JoVE Journal

Biology

3.9K 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

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 ...

Metabolic Profile Analysis of Zebrafish Embryos

A growing goal in the field of metabolism is to determine the impact of genetics on different aspects of mitochondrial function. Understanding these relationships will help to understand the underlying etiology for a range of diseases linked with mitochondrial dysfunction, such as diabetes and obesity. Recent advances in instrumentation, has enabled the monitoring of distinct parameters of mitochondrial function in cell lines or tissue explants. Here we present a method for a rapid and sensitive analysis of mitochondrial function parameters in vivo during zebrafish embryonic development using the Seahorse bioscience XF 24 extracellular flux analyser. This protocol utilizes the Islet Capture microplates where a single embryo is placed in each well, allowing measurement of bioenergetics, including: (i) basal respiration; (ii) basal mitochondrial respiration (iii) mitochondrial respiration due to ATP turnover; (iv) mitochondrial uncoupled respiration or proton leak and (iv) maximum respiration. Using this approach embryonic zebrafish respiration parameters can be compared between wild type and genetically altered embryos (mutant, gene over-expression or gene knockdown) or those manipulated pharmacologically. It is anticipated that dissemination of this protocol will provide researchers with new tools to analyse the genetic basis of metabolic disorders in vivo in this relevant vertebrate animal model.

Zebrafish represent a powerful vertebrate model that has been under-utilised for metabolic studies. Here we describe a rapid way to measure the in vivo metabolic profile of developing zebrafish that allows the comparison of different mitochondrial function parameters between genetically or pharmacologically manipulated embryos, thereby increasing the applicability of this organism.

A growing goal in the field of metabolism is to determine the impact of genetics on different aspects of mitochondrial function. Understanding these ...

Intravital Microscopy of the Microcirculation in the Mouse Cremaster Muscle for the Analysis of Peripheral Stem Cell Migration

In the era of intravascular cell application protocols in the context of regenerative cell therapy, the underlying mechanisms of stem cell migration to nonmarrow tissue have not been completely clarified. We describe here the technique of intravital microscopy applied to the mouse cremaster microcirculation for analysis of peripheral bone marrow stem cell migration in vivo. Intravital microscopy of the M. cremaster has been previously introduced in the field of inflammatory research for direct observation of leucocyte interaction with the vascular endothelium. Since sufficient peripheral stem and progenitor cell migration includes similar initial steps of rolling along and firm adhesion at the endothelial lining it is conceivable to apply the M. cremaster model for the observation and quantification of the interaction of intravasculary administered stem cells with the endothelium. As various chemical components can be selectively applied to the target tissue by simple superfusion techniques, it is possible to establish essential microenvironmental preconditions, for initial stem cell recruitment to take place in a living organism outside the bone marrow.

Intravital microscopy of the mouse M. cremaster microcirculation offers a unique and well-standardized in vivo model for the analysis of peripheral bone marrow stem cell migration.

In the era of intravascular cell application protocols in the context of regenerative cell therapy, the underlying mechanisms of stem cell migration to ...

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 ...

Analysis of Oxidative Stress in Zebrafish Embryos

High levels of reactive oxygen species (ROS) may cause a change of cellular redox state towards oxidative stress condition. This situation causes oxidation of molecules (lipid, DNA, protein) and leads to cell death. Oxidative stress also impacts the progression of several pathological conditions such as diabetes, retinopathies, neurodegeneration, and cancer. Thus, it is important to define tools to investigate oxidative stress conditions not only at the level of single cells but also in the context of whole organisms. Here, we consider the zebrafish embryo as a useful in vivo system to perform such studies and present a protocol to measure in vivo oxidative stress. Taking advantage of fluorescent ROS probes and zebrafish transgenic fluorescent lines, we develop two different methods to measure oxidative stress in vivo: i) a &#8220;whole embryo ROS-detection method&#8221; for qualitative measurement of oxidative stress and ii) a &#8220;single-cell&#160;ROS detection method&#8221; for quantitative measurements of oxidative stress. Herein, we demonstrate the efficacy of these procedures by increasing oxidative stress in tissues by oxidant agents and physiological or genetic methods. This protocol is amenable for forward genetic screens and it will help address cause-effect relationships of ROS in animal models of oxidative stress-related pathologies such as neurological disorders and cancer.

Here we report a protocol to measure oxidative stress in living zebrafish embryos. This procedure allows reactive oxygen species (ROS) detection in both whole embryo tissues and single-cell populations. This protocol will accomplish both qualitative and quantitative analyses.

High levels of reactive oxygen species (ROS) may cause a change of cellular redox state towards oxidative stress condition. This situation causes ...

Setting Up a Simple Light Sheet Microscope for In Toto Imaging of C. elegans Development

Fast and low phototoxic imaging techniques are pre-requisite to study the development of organisms in toto. Light sheet based microscopy reduces photo-bleaching and phototoxic effects compared to confocal microscopy, while providing 3D images with subcellular resolution. Here we present the setup of a light sheet based microscope, which is composed of an upright microscope and a small set of opto-mechanical elements for the generation of the light sheet. The protocol describes how to build, align the microscope and characterize the light sheet. In addition, it details how to implement the method for in toto imaging of C. elegans embryos using a simple observation chamber. The method allows the capture of 3D two-colors time-lapse movies over few hours of development. This should ease the tracking of cell shape, cell divisions and tagged proteins over long periods of time.

Setting Up a Simple Light Sheet Microscope for In Toto Imaging of C. elegans Development

This protocol describes the setup of a light sheet microscope and its implementation for in vivo imaging of C. elegans embryos.

Fast and low phototoxic imaging techniques are pre-requisite to study the development of organisms in toto. Light sheet based microscopy reduces ...

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and temporal resolution to study in vivo hematopoiesis using the murine bone marrow transplant experimental model. Genetic combinatorial marking using lentiviral vectors encoding fluorescent proteins (FPs) enabled cell fate mapping through advanced microscopy imaging. Vectors encoding five different FPs: Cerulean, EGFP, Venus, tdTomato, and mCherry were used to concurrently transduce HSPCs, creating a diverse palette of color marked cells. Imaging using confocal/two-photon hybrid microscopy enables simultaneous high resolution assessment of uniquely marked cells and their progeny in conjunction with structural components of the tissues. Volumetric analyses over large areas reveal that spectrally coded HSPC-derived cells can be detected non-invasively in various intact tissues, including the bone marrow (BM), for extensive periods of time following transplantation. Live studies combining video-rate multiphoton and confocal time-lapse imaging in 4D demonstrate the possibility of dynamic cellular and clonal tracking in a quantitative manner.

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

Combinatorial 5 fluorescent proteins marking of hematopoietic stem and progenitor cells allows in vivo clonal tracking via confocal and two-photon microscopy, providing insights into bone marrow hematopoietic architecture during regeneration. This method allows non-invasive fate mapping of spectrally-coded HSPCs-derived cells in intact tissues for extensive periods of time following transplantation.

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and ...

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) maintain the turnover of all mature blood cell lineages. The coordination of the complex signals leading to specific HSC fates relies upon the interaction between HSCs and the intricate bone marrow microenvironment, which is still poorly understood[1-2].
We describe how by combining a newly developed specimen holder for stable animal positioning with multi-step confocal and two-photon in vivo imaging techniques, it is possible to obtain high-resolution 3D stacks containing HSPCs and their surrounding niches and to monitor them over time through multi-point time-lapse imaging. High definition imaging allows detecting ex vivo labeled hematopoietic stem and progenitor cells (HSPCs) residing within the bone marrow. Moreover, multi-point time-lapse 3D imaging, obtained with faster acquisition settings, provides accurate information about HSPC movement and the reciprocal interactions between HSPCs and stroma cells.
Tracking of HSPCs in relation to GFP positive osteoblastic cells is shown as an exemplary application of this method. This technique can be utilized to track any appropriately labeled hematopoietic or stromal cell of interest within the mouse calvarium bone marrow space.

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

The nature of the interactions between hematopoietic stem and progenitor cells (HSPCs) and bone marrow niches is poorly understood. Custom hardware modifications and a multi-step acquisition protocol allow the use of two-photon and confocal microscopy to image ex vivo labeled HSPCs homed within bone marrow areas, tracking interactions and movement.

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) ...

Light Sheet Fluorescence Microscopy of Plant Roots Growing on the Surface of a Gel

One of the key questions in understanding plant development is how single cells behave in a larger context of the tissue. Therefore, it requires the observation of the whole organ with a high spatial- as well as temporal resolution over prolonged periods of time, which may cause photo-toxic effects. This protocol shows a plant sample preparation method for light-sheet microscopy, which is characterized by mounting the plant vertically on the surface of a gel. The plant is mounted in such a way that the roots are submerged in a liquid medium while the leaves remain in the air. In order to ensure photosynthetic activity of the plant, a custom-made lighting system illuminates the leaves. To keep the roots in darkness the water surface is covered with sheets of black plastic foil. This method allows long-term imaging of plant organ development in standardized conditions.

This protocol shows a plant sample preparation method for light-sheet microscopy. The setup is characterized by mounting the plant vertically on the surface of a gel and letting it grow in controlled bright conditions. This allows long-term observation of plant organ development in standardized conditions.

One of the key questions in understanding plant development is how single cells behave in a larger context of the tissue. Therefore, it requires the ...

Analysis of the Gap Junction-dependent Transfer of miRNA with 3D-FRAP Microscopy

Small antisense RNAs, like miRNA and siRNA, play an important role in cellular physiology and pathology and, moreover, can be used as therapeutic agents in the treatment of several diseases. The development of new, innovative strategies for miRNA/siRNA therapy is based on an extensive knowledge of the underlying mechanisms. Recent data suggest that small RNAs are exchanged between cells in a gap junction-dependent manner, thereby inducing gene regulatory effects in the recipient cell. Molecular biological techniques and flow cytometric analysis are commonly used to study the intercellular exchange of miRNA. However, these methods do not provide high temporal resolution, which is necessary when studying the gap junctional flux of molecules. Therefore, to investigate the impact of miRNA/siRNA as intercellular signaling molecules, novel tools are needed that will allow for the analysis of these small RNAs at the cellular level. The present protocol describes the application of three-dimensional fluorescence recovery after photobleaching (3D-FRAP) microscopy to elucidating the gap junction-dependent exchange of miRNA molecules between cardiac cells. Importantly, this straightforward and non-invasive live-cell imaging approach allows for the visualization and quantification of the gap junctional shuttling of fluorescently labeled small RNAs in real time, with high spatio-temporal resolution. The data obtained by 3D-FRAP confirm a novel pathway of intercellular gene regulation, where small RNAs act as signaling molecules within the intercellular network.

Here, we describe the application of three-dimensional fluorescence recovery after photobleaching (3D-FRAP) for the analysis of the gap junction-dependent shuttling of miRNA. In contrast to commonly applied methods, 3D-FRAP allows for the quantification of the intercellular transfer of small RNAs in real time, with high spatio-temporal resolution.

Small antisense RNAs, like miRNA and siRNA, play an important role in cellular physiology and pathology and, moreover, can be used as therapeutic agents ...