Name: following endocardial tissue movements via cell photoconversion in the zebrafish embryo
Uploaded: 2018-02-20T14:00:00
Description: Watch this Scientific Journal Video about Following Endocardial Tissue Movements via Cell Photoconversion in the Zebrafish Embryo at JoVE.com

We would like to thank Anne-Laure Duchemin, Denise Duchemin, Nathalie Faggianelli and Basile Gurchenkov for helping to design and make the mold described in this protocol. This work was supported by FRM (DEQ20140329553), the ANR (ANR-15-CE13-0015-01, ANR-12-ISV2-0001-01), the EMBO Young Investigator Program, the European Community, ERC CoG N&#176;682938 Evalve and by the grant ANR-10-LABX-0030-INRT, a French State fund managed by the Agence Nationale de la Recherche under the frame program Investissements d'Avenir labeled ANR-10-IDEX-0002-02.

1. Preparing molds and mounting agarose
<ol>
	<li>Create a mold with the dimensions shown in Figure 1 using either a 3D printer or traditional mechanical workshop tools.</li>
	<li>Pipette about 1.5 mL of melted 1 % agarose into a 35-mm plastic mounting dish. Place the plastic mold in the liquid agarose, taking care to avoid trapping air between the mold and the agarose. Place the dish at 4 &#176;C, wait till the agarose hardens (this takes about 5 min), then remove the plastic mold.</li>
	...

<ol>
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W., Tristani-Firouzi, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Blebbistatin+effectively+uncouples+the+excitation-contraction+process+in+zebrafish+embryonic+heart.">Blebbistatin effectively uncouples the excitation-contraction process in zebrafish embryonic heart.</a> Cell Physiol Biochem. 25 (4-5), 419-424 (2010).</li><li>Pestel, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Real-time+3D+visualization+of+cellular+rearrangements+during+cardiac+valve+formation.">Real-time 3D visualization of cellular rearrangements during cardiac valve formation.</a> Development. 143 (12), 2217-2227 (2016).</li><li>Swinburne, I. A., Mosaliganti, K. R., Green, A. A., Megason, S. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Improved+Long-Term+Imaging+of+Embryos+with+Genetically+Encoded+alpha-Bungarotoxin.">Improved Long-Term Imaging of Embryos with Genetically Encoded alpha-Bungarotoxin.</a> PLoS One. 10 (8), e0134005 (2015).</li><li>Kaufmann, A., Mickoleit, M., Weber, M., Huisken, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multilayer+mounting+enables+long-term+imaging+of+zebrafish+development+in+a+light+sheet+microscope.">Multilayer mounting enables long-term imaging of zebrafish development in a light sheet microscope.</a> Development. 139 (17), 3242-3247 (2012).</li><li>Marchant, J. S., Stutzmann, G. E., Leissring, M. A., LaFerla, F. M., Parker, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multiphoton-evoked+color+change+of+DsRed+as+an+optical+highlighter+for+cellular+and+subcellular+labeling.">Multiphoton-evoked color change of DsRed as an optical highlighter for cellular and subcellular labeling.</a> Nat Biotechnol. 19 (7), 645-649 (2001).</li><li>Dempsey, W. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+single-cell+labeling+by+confined+primed+conversion.">In vivo single-cell labeling by confined primed conversion.</a> Nat Methods. 12 (7), 645-648 (2015).</li><li>Mohr, M. A., Pantazis, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single+neuron+morphology+in+vivo+with+confined+primed+conversion.">Single neuron morphology in vivo with confined primed conversion.</a> Methods Cell Biol. 133, 125-138 (2016).</li><li>Mohr, M. A., Argast, P., Pantazis, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Labeling+cellular+structures+in+vivo+using+confined+primed+conversion+of+photoconvertible+fluorescent+proteins.">Labeling cellular structures in vivo using confined primed conversion of photoconvertible fluorescent proteins.</a> Nat Protoc. 11 (12), 2419-2431 (2016).</li><li>Mohr, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rational+Engineering+of+Photoconvertible+Fluorescent+Proteins+for+Dual-Color+Fluorescence+Nanoscopy+Enabled+by+a+Triplet-State+Mechanism+of+Primed+Conversion.">Rational Engineering of Photoconvertible Fluorescent Proteins for Dual-Color Fluorescence Nanoscopy Enabled by a Triplet-State Mechanism of Primed Conversion.</a> Angew Chem Int Ed Engl. 56 (38), 11628-11633 (2017).</li><li>Dempsey, W. P., Fraser, S. E., Pantazis, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=PhOTO+zebrafish:+a+transgenic+resource+for+in+vivo+lineage+tracing+during+development+and+regeneration.">PhOTO zebrafish: a transgenic resource for in vivo lineage tracing during development and regeneration.</a> PLoS One. 7 (3), e32888 (2012).</li><li>Dempsey, W. 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Timing of photoconversion: Although Kaede remains brightly expressed in EdCs even at 96 hpf, it should be noted that as the embryo grows, laser light diffuses more before it reaches the AVC, making confined photoconversion of Kaede more difficult. At embryonic stages later than 55 hpf, the ballooning of the ventricle and the atrium also means that the violet laser beam used for photoconversion cannot reach AVC cells without first passing through either the atrium or ventricle. This means that beyond 55 hpf, in o...

The zebrafish is currently one of the most important vertebrate models to study cellular and developmental processes in vivo. This is largely due to the zebrafish's optical transparency and amenability to genetics, which makes it a powerful model for applying optical techniques involving genetically encoded photoresponsive protein technologies1. Specific to the study of heart development, zebrafish receive sufficient oxygen via diffusion such that even mutants without heartbeat can survive through the first week of development, permitting analyses on the effects of developmental genes and perturbed blood flow on heart morphogenesis not possible in most vertebrates2.
The zebrafish heart tube is formed by 24 hours post fertilization (hpf). Shortly after its formation, the heart tube starts actively beating. By 36 hpf, a clear constriction separates the atrium from the ventricle. This region of constriction is called the atrioventricular canal (AVC), and cells in this region change from a squamous morphology to a cuboidal morphology3. Zebrafish atrioventricular valve morphogenesis starts around 48 h post fertilization. By 5 days post fertilization, two valve leaflets extend into the AVC and prevent the back flow of blood from the ventricle to the atrium during the cardiac cycle4. Tracking cells during AVC and valve formation is challenging as the rapid beating of the heart makes it difficult to follow cells via traditional time-lapse microscopy5,6. This protocol, adapted from Steed et al., 20167, describes a method that uses the Tg(fli1a:gal4FFubs, UAS:kaede)8 zebrafish transgenic line, in which the photoconvertible protein Kaede is expressed in endothelial cells, including the endocardium. The drug 2,3-butanedione-2-monoxime (BDM) is used to temporarily stop the heart beating, allowing accurate photoconversion of EdCs between 36 and 55 hpf, and high-resolution imaging of photoconverted EdCs at specific developmental time points. It has previously been shown that EdCs photoconverted using this method can remain distinguishable from their unconverted neighbors for five days or more after the time of photoconversion7. This protocol also details a method used for image analysis of photoconverted EdCs in embryos younger than 48 hpf, which has successfully been used to follow tissue movements during AVC development (Boselli et al., in press)9. We hope that readers would find this protocol useful for studying AVC and valve development in normal embryos, and in mutants, morphants, or drug-treated embryos. For a more general protocol relating to cell tracking using photoconvertible proteins during zebrafish development, please view the article by Lombardo et al., 201210.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Materials</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Necessary equipment for raising fish and collecting eggs (see the Zebrafish Book22 for details).</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Stereomicroscope</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Upright confocal microscope (Equipped with lasers operating at 488 nm, 561 nm, and 405 nm, a tunable multiphoton laser, and a Leica HCX IRAPO L, 25 &#215;, N.A. 0.95 objective)</td>
			<td>Leica</td>
			<td>Leica SP8</td>
			<td></td>
		</tr>
		<tr>
			<td>Heat block</td>
			<td>ThermoScientific</td>
			<td>88870001</td>
			<td></td>
		</tr>
		<tr>
			<td>35 mm x 10 mm tissue culture dish</td>
			<td>Falcon</td>
			<td>353001</td>
			<td></td>
		</tr>
		<tr>
			<td>6 well plate</td>
			<td>Falcon</td>
			<td>353046</td>
			<td></td>
		</tr>
		<tr>
			<td>Petri dish</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Glass pipette</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Mold</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Reagents</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>8 mg/mL Tricaine stock solution&#160;</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>1 M BDM stock solution</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>UltraPure low melting point agarose</td>
			<td>Invitrogen</td>
			<td>16520-050</td>
			<td></td>
		</tr>
		<tr>
			<td>Agarose</td>
			<td>Lonza</td>
			<td>50004</td>
			<td></td>
		</tr>
		<tr>
			<td>PTU (1-phenyl-2-thiourea)</td>
			<td>Sigma Aldrich</td>
			<td>P7629</td>
			<td></td>
		</tr>
		<tr>
			<td>Embryo medium: 30x stock solution</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Software</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Matlab equipped with the Image Analysis, Curve Fitting, Bio-Formats Toolboxes</td>
			<td>MathWorks</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>8 mg/mL Tricaine stock solution</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>200 mg of tricaine powder (Ethyl 3-aminobenzoate methanesulfonic acid)</td>
			<td>Sigma-Aldrich</td>
			<td>E10521</td>
			<td></td>
		</tr>
		<tr>
			<td>25 mL Danieau</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Adjust to pH 7, aliquot and store at -20 &#176;C&#160;</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>1 M BDM stock solution</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>1 g BDM</td>
			<td>Sigma-Aldrich</td>
			<td>112135</td>
			<td></td>
		</tr>
		<tr>
			<td>10 mL ddH2O</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Aliquot and store at -20 &#176;C</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Embryo medium: 30x stock solution</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>50.7 g NaCl</td>
			<td>Sigma-Aldrich</td>
			<td>7647-14-5</td>
			<td></td>
		</tr>
		<tr>
			<td>0.78 g KCl</td>
			<td>Sigma-Aldrich</td>
			<td>7447-40-7</td>
			<td></td>
		</tr>
		<tr>
			<td>1.47 g Magnesium sulfate</td>
			<td>Sigma-Aldrich</td>
			<td>&#160;7487-88-9</td>
			<td></td>
		</tr>
		<tr>
			<td>2.1 g Calcium nitrite tetrahydrate</td>
			<td>Sigma-Aldrich</td>
			<td>13477-34-4</td>
			<td></td>
		</tr>
		<tr>
			<td>19.52 g HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid)</td>
			<td>Sigma-Aldrich</td>
			<td>7365-45-9</td>
			<td></td>
		</tr>
		<tr>
			<td>Adjust to pH 7.2 and store at room temperature</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Mold</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>PlasCLEAR resin</td>
			<td>Asiga</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Plus 39&#160;Freeform Pico 3D printer</td>
			<td>Asiga</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

An example of an embryo photoconverted at 48 hpf and imaged again at 80 hpf is shown in Figure 2, Movies 1 and 2. Exposing Kaede to 405 nm light irreversibly converts from the protein from its fluorescent green form to fluorescent red form, enabling the behavior of cells labeled with either the green or their red form to be followed with respect to their differentially colored neighbors during valve formation. It can be seen ...

Watch this Scientific Journal Video about Following Endocardial Tissue Movements via Cell Photoconversion in the Zebrafish Embryo at JoVE.com

Following Endocardial Tissue Movements via Cell Photoconversion in the Zebrafish Embryo

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 ...

The overall goal of this procedure is to follow endocardial cells during atrioventricular canal and heart valve formation. This method can help to address key question in the field of cardiac development, such as to understand how endocardial cells behave in different types of mutants. Tracking cells during AVC and valve formation is challenging because of the rapid beat of the heart.It makes it difficult to follow cells via traditional time-lapse microscopy. The main advantage of this technique is that it bypasses some of these difficulties. Demonstrating the procedure will be Renee, a postdoc in my lab.After creating a mold according to the text protocol, pipette about 1.5 milliliters of melted 1%agarose into a 35-millimeter plastic mounting dish. Then, position the plastic mold in the liquid agarose, taking care to avoid trapping air between the mold and the agarose. Place the dish at four degrees Celsius until the agarose hardens.Then, remove the plastic mold from the agarose. Next, add two milliliters of previously prepared mounting medium to the mounting dish, and allow 15 to 30 minutes for the solution to diffuse into the agarose. In the meantime, melt a one-milliliter aliquot of 0.7%low melting point, or LMP, agarose by placing the tube in a 70-degree Celsius heat block for about five minutes.Pipette the LMP agarose up and down to cool it slightly. Then, add 25 microliters of eight milligram per milliliter tricaine and 40 microliters of one molar BDM solutions to the tube. Mix by pipetting up and down.Then, transfer the tube to a 38-degree Celsius heat block to keep the LMP agarose in its liquid state. Transfer the embryos to a separate dish containing two milliliters of mounting medium. Then, when the hearts have stopped beating, transfer the embryos to the mounting dish and use forceps to arrange them in the wells so that they lie at a 25-degree angle, tails pointing toward the deeper part of the well, with the yolk facing up.When the embryos are roughly in place, remove the medium, and embed them in approximately 200 microliters of 0.7%LMP agarose containing tricaine and BDM. Then, readjust the embryo positions, tilting them to the left or right as necessary. Press down on the walls of the indented square to allow excess agarose to overflow to the side areas of the mounting dish.Wait approximately five minutes for the LMP agarose to set, then add mounting medium to the dish. The embryos are now ready to be photoconverted. To carry out photoconversion, on an upright confocal microscope equipped with a 405, 488, and 561-nanometer laser sources, use transmitted white light and an objective lens to locate the embryo's heart.Using the 561-nanometer laser, check for red Kaede fluorescence. Then, use the 488-nanometer laser to visualize the endocardium. Next, on the microscope acquisition software, enter FRAP mode.Then, select approximately 1%laser power for both the 488 and the 561-nanometer lasers. While focusing on the plane of interest, select a region of interest, or ROI. Then, using the 405-nanometer diode laser at 25%laser power, photoconvert the cells, scanning over the ROI three times.Should insufficient Kaede be photoconverted in the ROI, scan the 405-nanometer laser over the area three times again. Return to the TCS SP8 mode on the software, and using the 561 and 488-nanometer lasers sequentially, in Bidirectional mode, acquire a Z-stack. After photoconversion, use a glass pipette to press down slightly just above the head of the embryo to break the LMP agarose.Then, gently suck up the embryo. To wash away the BDM containing medium, eject the embryo from the glass pipette into an 85-millimeter Petri dish containing embryo medium with PTU. Transfer the embryo to a well in a six-well plate containing embryo medium with PTU.During this step, make sure to keep note of which embryo goes into which well, as this is essential to correlate the position of photoconverted cells at later developmental stages. Now that the embryos have been removed from the BDM containing medium, their hearts should start beating again in about five minutes. Return embryos to the dark at 28.5 degrees Celsius to allow them to continue to develop normally.At the desired stage, stop the heart and re-embed the embryos, as demonstrated earlier in this video, with the important difference that the embryos must be treated with BDM in separate, marked dishes to keep track of them. For embryos up to 62 hpf, acquire Z-stacks of the endocardium using 561 and 488 nanometers for red and green fluorescent signals, respectively. For embryos older than 62 hpf, use a multiphoton laser set to 940 nanometers to image green Kaede instead of the 488-nanometer laser.For embryos younger than 48 hpf, it is possible to project the three-dimensional AVC onto a two-dimensional image for analysis. First, threshold the image. Then, erase points that lie outside of the heart.To perform manual segmentation, orient the image on the left to match the image on the right. Segment the side of the heart in which the orange vector passes through, and add points in the direction of the blue vector. Next, interpolate the cross-sections, and project the color intensity onto the surface.Finally, adjust the orientation of the AVC, and project it onto a 2D surface. An example of an embryonic heart just after photoconversion at 48 hpf is shown here. Cytoplasmic Kaede in several cells of the superior side of the AVC has been converted from green to red.In the same embryo at 80 hpf, it's evident that the photoconverted cells have proliferated and have migrated to the inner side of the superior atrioventricular valve. An example of a 36 hpf embryo where the EdCs in the atrium and ventricle have been photoconverted is shown here. The AVC is then projected into two dimensions for easier visualization and analysis and allows one to quantify the distance between the two photoconverted regions.While attempting this procedure, it's important to keep the time the heart is stopped to a minimum to avoid affecting normal development. With practice, photoconversion can be completed within about 30 minutes provided that everything is prepared in advance. After watching this video, you should have a good understanding of how to track endocardial cells after photoconversion in live zebrafish embryo and analyze your data sets.

following-endocardial-tissue-movements-via-cell-photoconversion

Research

JoVE Journal

Developmental Biology

Measuring Protein Stability in Living Zebrafish Embryos Using Fluorescence Decay After Photoconversion (FDAP)

Protein stability influences many aspects of biology, and measuring the clearance kinetics of proteins can provide important insights into biological systems. In FDAP experiments, the clearance of proteins within living organisms can be measured. A protein of interest is tagged with a photoconvertible fluorescent protein, expressed in vivo and photoconverted, and the decrease in the photoconverted signal over time is monitored. The data is then fitted with an appropriate clearance model to determine the protein half-life. Importantly, the clearance kinetics of protein populations in different compartments of the organism can be examined separately by applying compartmental masks. This approach has been used to determine the intra- and extracellular half-lives of secreted signaling proteins during zebrafish development. Here, we describe a protocol for FDAP experiments in zebrafish embryos. It should be possible to use FDAP to determine the clearance kinetics of any taggable protein in any optically accessible organism.

Measuring Protein Stability in Living Zebrafish Embryos Using Fluorescence Decay After Photoconversion FDAP

Protein levels in cells and tissues are often tightly regulated by the balance of protein production and clearance. Using Fluorescence Decay After Photoconversion (FDAP), the clearance kinetics of proteins can be experimentally measured in vivo.

Protein stability influences many aspects of biology, and measuring the clearance kinetics of proteins can provide important insights into biological ...

Microbead Implantation in the Zebrafish Embryo

The zebrafish has emerged as a valuable genetic model system for the study of developmental biology and disease. Zebrafish share a high degree of genomic conservation, as well as similarities in cellular, molecular, and physiological processes, with other vertebrates including humans. During early ontogeny, zebrafish embryos are optically transparent, allowing researchers to visualize the dynamics of organogenesis using a simple stereomicroscope. Microbead implantation is a method that enables tissue manipulation through the alteration of factors in local environments. This allows researchers to assay the effects of any number of signaling molecules of interest, such as secreted peptides, at specific spatial and temporal points within the developing embryo. Here, we detail a protocol for how to manipulate and implant beads during early zebrafish development.

The zebrafish is an excellent model system for genetic and developmental studies. Bead implantation is a valuable tissue manipulation technique that can be used to interrogate developmental mechanisms by introducing alterations in local cellular environments. This protocol describes how to perform microbead implantation in the zebrafish embryo.

The zebrafish has emerged as a valuable genetic model system for the study of developmental biology and disease. Zebrafish share a high degree of genomic ...

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 ...

Observing Mitotic Division and Dynamics in a Live Zebrafish Embryo

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin condensation, microtubule-kinetochore attachment, chromosome segregation, and cytokinesis in a small time frame. Errors in the delicate process can result in human disease, including birth defects and cancer. Traditional approaches investigating human mitotic disease states often rely on cell culture systems, which lack the natural physiology and developmental/tissue-specific context advantageous when studying human disease. This protocol overcomes many obstacles by providing a way to visualize, with high resolution, chromosome dynamics in a vertebrate system, the zebrafish. This protocol will detail an approach that can be used to obtain dynamic images of dividing cells, which include: in vitro transcription, zebrafish breeding/collecting, embryo embedding, and time-lapse imaging. Optimization and modifications of this protocol are also explored. Using H2A.F/Z-EGFP (labels chromatin) and mCherry-CAAX (labels cell membrane) mRNA-injected embryos, mitosis in AB wild-type, auroraBhi1045, and esco2hi2865 mutant zebrafish is visualized. High resolution live imaging in zebrafish allows one to observe multiple mitoses to statistically quantify mitotic defects and timing of mitotic progression. In addition, observation of qualitative aspects that define improper mitotic processes (i.e., congression defects, missegregation of chromosomes, etc.) and improper chromosomal outcomes (i.e., aneuploidy, polyploidy, micronuclei, etc.) are observed. This assay can be applied to the observation of tissue differentiation/development and is amenable to the use of mutant zebrafish and pharmacological agents. Visualization of how defects in mitosis lead to cancer and developmental disorders will greatly enhance understanding of the pathogenesis of disease.

Mitosis is critical to every living organism and defects often lead to cancer and developmental disorders. Using this imaging protocol and zebrafish as a model system, researchers can visualize mitosis in a live vertebrate organism and the multitude of defects that arise when mitotic processes are defective.

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin ...

Visualizing the Interrenal Steroidogenic Tissue and Its Vascular Microenvironment in Zebrafish

This protocol introduces how to detect differentiated interrenal steroidogenic cells through a simple whole-mount enzymatic activity assay. Identifying differentiated steroidogenic tissues through chromogenic histochemical staining of 3-&#946;-Hydroxysteroid dehydrogenase /&#916;5-4 isomerase (3&#946;-Hsd) activity-positive cells is critical for monitoring the morphology and differentiation of adrenocortical and interrenal tissues in mammals and teleosts, respectively. In the zebrafish model, the optical transparency and tissue permeability of the developing embryos and larvae allow for whole-mount staining of 3&#946;-Hsd activity. This staining protocol, as performed on transgenic fluorescent reporter lines marking the developing pronephric and endothelial cells, enables the detection of the steroidogenic interrenal tissue in addition to the kidney and neighboring vasculature. In combination with vibratome sectioning, immunohistochemistry, and confocal microscopy, we can visualize and assay the vascular microenvironment of interrenal steroidogenic tissues. The 3&#946;-Hsd activity assay is essential for studying the cell biology of the zebrafish interrenal gland because to date, no suitable antibody is available for labeling zebrafish steroidogenic cells. Furthermore, this assay is rapid and simple, thus providing a powerful tool for mutant screens targeting adrenal (interrenal) genetic disorders as well as for determining disruption effects of chemicals on steroidogenesis in pharmaceutical or toxicological studies.

The interrenal gland in zebrafish is the teleostean counterpart of the mammalian adrenal gland. This protocol introduces how to perform the 3-&#946;-hydroxysteroid dehydrogenase (&#916;5-4 isomerase; 3&#946;-Hsd) enzymatic activity assay, which detects differentiated steroidogenic cells in the developing zebrafish.

This protocol introduces how to detect differentiated interrenal steroidogenic cells through a simple whole-mount enzymatic activity assay. Identifying ...

Use of Immunolabeling to Analyze Stable, Dynamic, and Nascent Microtubules in the Zebrafish Embryo

Microtubules (MTs) are dynamic and fragile structures that are challenging to image in vivo, particularly in vertebrate embryos. Immunolabeling methods are described here to analyze distinct populations of MTs in the developing neural tube of the zebrafish embryo. While the focus is on neural tissue, this methodology is broadly applicable to other tissues. The procedures are optimized for early to mid-somitogenesis-stage embryos (1 somite to 12 somites), however they can be adapted to a range of other stages with relatively minor adjustments. The first protocol provides a method to assess the spatial distribution of stable and dynamic MTs and perform a quantitative analysis of these populations with image-processing software. This approach complements existing tools to image microtubule dynamics and distribution in real-time, using transgenic lines or transient expression of tagged constructs. Indeed, such tools are very useful, however they do not readily distinguish between dynamic and stable MTs. The ability to image and analyze these distinct microtubule populations has important implications for understanding mechanisms underlying cell polarization and morphogenesis. The second protocol outlines a technique to analyze nascent MTs specifically. This is accomplished by capturing the de novo growth properties of MTs over time, following microtubule depolymerization with the drug nocodazole and a recovery period after drug washout. This technique has not yet been applied to the study of MTs in zebrafish embryos, but is a valuable assay for investigating the in vivo function of proteins implicated in microtubule assembly.

Immunolabeling methods to analyze distinct populations of microtubules in the developing zebrafish brain are described here, which are broadly applicable to other tissues. The first protocol outlines an optimized method for immunolabeling stable and dynamic microtubules. The second protocol provides a method to image and quantify nascent microtubules specifically.

Microtubules (MTs) are dynamic and fragile structures that are challenging to image in vivo, particularly in vertebrate embryos. Immunolabeling methods ...

Live Imaging of Mouse Secondary Palate Fusion

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to cleft secondary palate, a common congenital anomaly in humans. Secondary palate fusion has been extensively studied leading to several proposed cellular mechanisms that may mediate this process. However, these studies have been mostly performed on fixed embryonic tissues at progressive timepoints during development or in fixed explant cultures analyzed at static timepoints. Static analysis is limited for the analysis of dynamic morphogenetic processes such a palate fusion and what types of dynamic cellular behaviors mediate palatal fusion is incompletely understood. Here we describe a protocol for live imaging of ex vivo secondary palate fusion in mouse embryos. To examine cellular behaviors of palate fusion, epithelial-specific Keratin14-cre was used to label palate epithelial cells in ROSA26-mTmGflox reporter embryos. To visualize filamentous actin, Lifeact-mRFPruby reporter mice were used. Live imaging of secondary palate fusion was performed by dissecting recently-adhered secondary palatal shelves of embryonic day (E) 14.5 stage embryos and culturing in agarose-containing media on a glass bottom dish to enable imaging with an inverted confocal microscope. Using this method, we have detected a variety of novel cellular behaviors during secondary palate fusion. An appreciation of how distinct cell behaviors are coordinated in space and time greatly contributes to our understanding of this dynamic morphogenetic process. This protocol can be applied to mutant mouse lines, or cultures treated with pharmacological inhibitors to further advance understanding of how secondary palate fusion is controlled.

Here, we present a protocol for live imaging of mouse secondary palate fusion using confocal microscopy. This protocol can be used in combination with a variety of fluorescent reporter mouse lines, and with pathway inhibitors for mechanistic insight. This protocol can be adapted for live imaging in other developmental systems.

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to ...

AFM and Microrheology in the Zebrafish Embryo Yolk Cell

Elucidating the factors that direct the spatio-temporal organization of evolving tissues is one of the primary purposes in the study of development. Various propositions claim to have been important contributions to the understanding of the mechanical properties of cells and tissues in their spatiotemporal organization in different developmental and morphogenetic processes. However, due to the lack of reliable and accessible tools to measure material properties and tensional parameters in vivo, validating these hypotheses has been difficult. Here we present methods employing atomic force microscopy (AFM) and particle tracking with the aim of quantifying the mechanical properties of the intact zebrafish embryo yolk cell during epiboly. Epiboly is an early conserved developmental process whose study is facilitated by the transparency of the embryo. These methods are simple to implement, reliable, and widely applicable since they overcome intrusive interventions that could affect tissue mechanics. A simple strategy was applied for the mounting of specimens, AFM recording, and nanoparticle injections and tracking. This approach makes these methods easily adaptable to other developmental times or organisms.

The lack of tools to measure material properties and tensional parameters in vivo prevents validating their roles during development. We employed atomic force microscopy (AFM) and nanoparticle-tracking to quantify mechanical features on the intact zebrafish embryo yolk cell during epiboly. These methods are reliable and widely applicable avoiding intrusive interventions.

Elucidating the factors that direct the spatio-temporal organization of evolving tissues is one of the primary purposes in the study of development. ...

Single-cell Photoconversion in Living Intact Zebrafish

Animal and plant tissue is composed of distinct populations of cells. These cells interact over time to build and maintain the tissue and can cause disease when disrupted. Scientists have developed clever techniques to investigate characteristics and natural dynamics of these cells within intact tissue by expressing fluorescent proteins in subsets of cells. However, at times, experiments require more selected visualization of cells within the tissue, sometimes at the single-cell or population-of-cells manner. To achieve this and visualize single cells within a population of cells, scientists have utilized single-cell photoconversion of fluorescent proteins. To demonstrate this technique, we show here how to direct UV light to an Eos-expressing cell of interest in an intact, living zebrafish. We then image those photoconverted Eos+ cells 24 h later to determine how they changed in the tissue. We describe two techniques: single cell photoconversion and photoconversions of populations of cell. These techniques can be used to visualize cell-cell interactions, cell-fate and differentiation, and cell migrations, making it a technique that is applicable in numerous biological questions.

Here, we present a protocol to show how cell photoconversion is achieved through UV exposure to specific areas expressing the fluorescent protein, Eos, in living animals.

Animal and plant tissue is composed of distinct populations of cells. These cells interact over time to build and maintain the tissue and can cause ...

Microinjection-based System for In Vivo Implantation of Embryonic Cardiomyocytes in the Avian Embryo

Interpreting the relative impact of cell autonomous patterning versus extrinsic microenvironmental influence on cell lineage determination represents a general challenge in developmental biology research. In the embryonic heart, this can be particularly difficult as regional differences in the expression of transcriptional regulators, paracrine/juxtacrine signaling cues, and hemodynamic force are all known to influence cardiomyocyte maturation. A simplified method to alter a developing cardiomyocyte's molecular and biomechanical microenvironment would, therefore, serve as a powerful technique to examine how local conditions influence cell fate and function. To address this, we have optimized a method to physically transplant juvenile cardiomyocytes into ectopic locations in the heart or the surrounding embryonic tissue. This allows us to examine how microenvironmental conditions influence cardiomyocyte fate transitions at single cell resolution within the intact embryo. Here, we describe a protocol in which embryonic myocytes can be isolated from a variety of cardiac sub-domains, dissociated, fluorescently labeled, and microinjected into host embryos with high precision. Cells can then be directly analyzed in situ using a variety of imaging and histological techniques. This protocol is a powerful alternative to traditional grafting experiments that can be prohibitively difficult in a moving tissue such as the heart. The general outline of this method can also be adapted to a variety of donor tissues and host environments, and its ease of use, low cost, and speed make it a potentially useful application for a variety of developmental studies.

In this method, embryonic cardiac tissues are surgically microdissected, dissociated, fluorescently labeled, and implanted into host embryonic tissues. This provides a platform for studying the individual or tissue level developmental organization under ectopic hemodynamic conditions, and/or altered paracrine/juxtacrine environments.

Interpreting the relative impact of cell autonomous patterning versus extrinsic microenvironmental influence on cell lineage determination represents a ...