The authors would like to thank Jeffrey Farrell, James Gagnon, and Jennifer Bergmann for comments on the manuscript. KWR was supported by the National Science Foundation Graduate Research Fellowship Program during the development of the FDAP assay. This work was supported by grants from the NIH to AFS and by grants from the German Research Foundation (Emmy Noether Program), the Max Planck Society, and the Human Frontier Science Program (Career Development Award) to PM.

1. Generating a Photoconvertible Fusion Construct and Injecting Dechorionated Zebrafish Embryos
<ol>
	<li>Generate a functional construct containing the protein of interest fused to a green-to-red photoconvertible protein (see Discussion), then use in vitro transcription to generate capped mRNA encoding the fusion protein as in M&#252;ller et al., 201219.</li>
	<li>Use pronase to remove the chorions from about 30 zebrafish embryos at the one-cell stage. Alternatively, manually dechorionate embryos using forceps28. 
	Note: Embryos must be dechorionated for subsequent imaging. If desired, embryos can be injected through the chorion and dechorionated later, just prior to imaging.
	<ol>
		<li>Make a 5 mg/ml stock solution of pronase from Streptomyces griseus in standard zebrafish embryo medium19. Rock the solution gently at RT for 10 min to allow the protease to dissolve. Aliquot 2 ml into microcentrifuge tubes and freeze at -20 &#176;C.</li>
		<li>Transfer one-cell stage embryos to a 5 cm diameter glass or agarose-coated plastic Petri dish containing ~8 ml embryo medium. Add 2 ml of thawed pronase stock solution to the dish and incubate at RT for 5 to 10 min.</li>
		<li>Avoid exposing embryos to air or plastic, as contact with either will cause dechorionated embryos to rupture. Fill a 200 ml glass beaker with embryo medium. Transfer the embryos to the beaker by tilting the Petri dish while submerging it in the medium.</li>
		<li>After the embryos have settled to the bottom of the beaker, pour out most of the embryo medium, then pour fresh embryo medium into the beaker. The mild swirling of the medium pouring into the beaker causes embryos to lose their weakened chorions.</li>
		<li>Repeat step 1.2.4.</li>
	</ol>
	</li>
	<li>Transfer the dechorionated embryos to an agarose-coated injection dish29 using a glass Pasteur pipette with a flamed tip. Flaming the pipette tip prevents jagged edges from injuring embryos.</li>
	<li>Co-inject the mRNA and a 3 kDa Alexa488-dextran conjugate29,30 (Figure 1B; see Discussion for suggested mRNA and Alexa488-dextran amounts). Inject directly into the center of the cell (not the yolk) to ensure even distribution of mRNA and fluorescent dye once cleavage commences. 
	Note: The Alexa488 signal will be used during data analysis to generate compartmental masks in order to distinguish between intracellular and extracellular fluorescence.</li>
	<li>Transfer injected embryos to a 1&#8211;2% agarose-coated well of a six-well plastic dish filled with embryo medium. Incubate in the dark at 28 &#176;C until embryos have reached late sphere stage31 (approximately 5 hr post fertilization). Check embryos every one to 2 hr under a stereomicroscope and remove any debris generated by embryos that have died.</li>
</ol>
2. Mounting Zebrafish Embryos for Photoconversion and Imaging on an Inverted Confocal Microscope
<ol>
	<li>Use a stereomicroscope to identify one to five healthy embryos, and use a glass Pasteur pipette with a flamed tip to remove them from the dish.</li>
	<li>Gently eject the embryos into a microcentrifuge tube containing ~1 ml of melted 1% low melting point agarose in 1x Danieau&#8217;s embryo medium (see Materials List) (Figure 2A). 
	Note: Agarose should have a temperature between 40 and 42 &#176;C; higher temperatures could damage the embryos.</li>
	<li>Draw the embryos back into the pipette along with some agarose. Gently eject the agarose and embryos onto the cover glass of a glass-bottom dish (Figure 2B). Ensure that the thickness of the cover glass is compatible with the objective on the confocal microscope.
	<ol>
		<li>Re-use the glass pipette if desired. To clean the residual agarose out of the pipette and prevent clogging, quickly pipette embryo medium up and down. Place a 15 ml tube filled with ~5 ml of embryo medium next to the stereomicroscope for this purpose.</li>
	</ol>
	</li>
	<li>Use a metal probe to position the embryos so that the animal pole (blastoderm) faces the cover glass. Work quickly since the agarose will solidify in 20&#8211;30 sec. Use the stereomicroscope to monitor the embryos&#8217; positions and readjust as necessary until the agarose hardens.</li>
	<li>Repeat steps 2.1&#8211;2.4 until the desired number of embryos has been mounted. 
	Note: In a typical experiment, four agarose drops containing four or five embryos each will fit easily on the cover glass. About 16 embryos can be imaged during a single ideal experiment (Figure 2C).</li>
	<li>When the agarose has solidified, fill the glass-bottom dish with 1x Danieau&#8217;s embryo medium.</li>
</ol>
3. Photoconverting and Measuring the Decrease of the Photoconverted Signal
A 25X or 40X water objective is appropriate for the size and refractive index of zebrafish embryos. It is best to use immersion oil with the same refractive index as water rather than actual water, since water will evaporate during the course of the five-hour experiment. Ensure that the immersion oil is designed to be used with a water (not oil) objective.
<ol>
	<li>Place a large drop of immersion oil on the objective to ensure that the oil film between the objective and cover glass will not break as the stage moves to different embryo positions during imaging. Securely place the glass-bottom dish onto the stage so that the dish will not shift when the stage moves. If possible, use a heated stage at 28 &#176;C, the optimal temperature for zebrafish development.</li>
	<li>Define each embryo&#8217;s position in the confocal microscope&#8217;s software package. Adjust the z-depth for each embryo, and attempt to target roughly the same plane in each embryo. 
	Note: About 30 &#956;m from the animal pole is a good depth since at this depth the enveloping layer of the embryo can be avoided, imaging area is maximized, and light scattering is minimal. A single optical slice with a thickness of ~3.3 &#956;m provides sufficient data; there is no need to acquire a z-stack (see Section 5).</li>
	<li>Collect two signals during the experiment: the &#8220;green&#8221; signal from the Alexa488-dextran conjugate&#8212;which will be used during data analysis to isolate extracellular and intracellular fluorescence&#8212;and the &#8220;red&#8221; signal from the fusion protein after it is photoconverted.
	<ol>
		<li>Excite Alexa488 using a 488 nm laser, and collect emitted fluorescence between ~500 and 540 nm. Note: After photoconversion, many green-to-red photoconvertible proteins (e.g., Dendra2) can be excited with a 543 nm laser and emit fluorescence between ~550 and 650 nm. Adjust as necessary based on the photoconvertible protein used.</li>
	</ol>
	</li>
	<li>Acquire &#8220;pre-photoconversion&#8221; images, and configure the confocal microscope&#8217;s software to image each of the previously defined positions (from step 3.2) with the appropriate imaging conditions every 10 or 20 min for a five-hour time course (see Section 5 and Discussion).</li>
	<li>To photoconvert the fusion protein, switch to a 10X objective and expose groups of embryos to UV light from a mercury arc lamp with a ~300&#8211;400 nm excitation filter at 100% output for 2 min. Shift the focus along the z-axis to promote uniform photoconversion (see Section 5). Ensure that the immersion oil does not drip onto the 10x objective during photoconversion. 
	Note: The shifting of focus during photoconversion could be automated in order to avoid variability among experimenters.</li>
	<li>Switch back to the 25X or 40X objective immediately after photoconversion. Ensure that the previously defined positions from step 3.2 are still accurate. If the dish shifted during photoconversion, re-define the positions of the embryos.
	<ol>
		<li>Start the program created in step 3.4 and allow imaging to continue for 5 hr. Note the time elapsed between photoconversion and the start of imaging for each embryo.</li>
	</ol>
	</li>
	<li>Occasionally check on the experiment. Monitor the level of Danieau&#8217;s medium and add more if necessary. Restart the software if it has stalled.</li>
	<li>In order to determine the background fluorescence values that will be used during data analysis to estimate the asymptote of an exponentially decreasing model, include some embryos that have been injected with Alexa488-dextran but not mRNA in the experiment. To determine the instrument noise, which will also be used during subsequent data analysis, acquire an image in the absence of a sample.</li>
</ol>
4. Analyzing the Data Using PyFDAP
<ol>
	<li>Visually inspect the time course data sets from each embryo. Discard data sets from embryos that died during imaging, that shifted significantly, that have very low levels of photoconverted signal, or that contain regions of cells that look unusual and have stopped moving and dividing (typical of injured or sick embryos). 
	Note: Occasionally, bubbles in the immersion oil or other artifacts will appear in one or two images in an otherwise usable data set. Note any images that contain artifacts; they will be discarded later, and the remaining time points from such a data set can still be analyzed.</li>
	<li>Use the Python-based software package PyFDAP to analyze the FDAP data. PyFDAP calculates half-lives by determining the average intracellular and extracellular red fluorescence intensity in each image and fitting the data with an exponentially decreasing function56 (Figure 3).
	<ol>
		<li>Download PyFDAP (see Materials List).</li>
		<li>Use PyFDAP to separate intracellular and extracellular photoconverted signal (Figure 3A,B). Use the Alexa488 signal, which is strictly intracellular, to create an intracellular mask. Apply this mask to the corresponding red channel image to prevent intracellular pixels from being considered when calculating average extracellular intensity. To measure average intracellular intensity, invert the mask.</li>
		<li>In PyFDAP, display the masked images generated in step 4.2.2. Visually inspect these images and discard data sets in which masks do not accurately distinguish intracellular from extracellular space (this should be rare; note that cell membranes are included in images in which extracellular space has been masked, but they could be removed by altering the thresholding algorithm or by introducing a membrane mask (e.g., using membrane-CFP)). Also discard any single images containing artifacts (e.g., bubbles in the immersion oil) identified in step 4.1.</li>
		<li>Use PyFDAP to calculate average extracellular and intracellular fluorescence intensities for each image. PyFDAP calculates these averages by summing the intensities of pixels that fall outside of the mask and dividing by the total number of pixels summed.</li>
		<li>Fit the fluorescence data (Figure 3C) with the following exponential function: 
		<img fo:content-width="1in" src="/files/ftp_upload/52266/52266eq1.jpg" width="159" /> 
		where t is time post-photoconversion, c(t) is intensity at a given value of t, c0 is the intensity at t = 0, k is the clearance rate constant, and y0 is the asymptote that the function approaches as fluorescence decreases (Figure 1D). y0 can be constrained based on the measurements in step 3.819.</li>
		<li>Use PyFDAP to calculate the extracellular and intracellular protein half-lives (&#964;) from the clearance rate constants (k) using the following relationship: 
		<img fo:content-width="1in" src="/files/ftp_upload/52266/52266eq2.jpg" width="153" /></li>
	</ol>
	</li>
</ol>
5. Control Experiments to Assess Photobleaching, Inadvertent Photoconversion, and Photoconversion Uniformity
<ol>
	<li>Assessing photobleaching 
	Note: Photobleaching could cause an artifactual decrease in fluorescence intensity that reflects the bleaching properties of the fluorescent protein in addition to the clearance of the protein of interest.
	<ol>
		<li>To assess possible photobleaching, perform one set of FDAP experiments with 10 min intervals between imaging and a second set with 20 min intervals between imaging (Figure 4). Analyze the data from both sets of experiments using PyFDAP as described in Section 4.</li>
		<li>Compare the half-lives from the 10 and 20 min interval experiments. Longer half-lives from 20 min interval experiments indicate significant photobleaching. If the half-lives from both experiments are identical, photobleaching is not a significant concern.</li>
		<li>Alternatively, assess photobleaching by acquiring a series of ~30 images immediately after photoconversion. A significant decrease in fluorescence intensity indicates significant photobleaching.</li>
		<li>If photobleaching is detected, use lower laser power, decrease imaging time, or consider using a more photostable photoconvertible protein32.</li>
	</ol>
	</li>
	<li>Assessing inadvertent photoconversion. 
	Note: Dendra2 can be photoconverted using 488 nm illumination27. When exciting Alexa488 with the 488 nm laser as described in step 3.3.1, inadvertent photoconversion and therefore an artifactual increase in the apparent half-life of the protein of interest is possible. However, we and others33 have found that 488 nm illumination is an inefficient method of photoconversion in zebrafish embryos.
	<ol>
		<li>Use the control experiment described in step 5.1.1 to detect inadvertent photoconversion. Compare the half-lives from the 10 and 20 min interval experiments. Shorter half-lives from 20 min interval experiments indicate significant inadvertent photoconversion. If the half-lives from both experiments are identical, inadvertent photoconversion is not a significant concern.</li>
		<li>If inadvertent photoconversion is detected, use a lower 488 nm laser power and shorter imaging times to avoid inadvertently photoconverting Dendra2.</li>
	</ol>
	</li>
	<li>Assessing photoconversion uniformity. 
	Note: If photoconversion is biased toward the animal pole of the embryo, the decrease in fluorescence will be influenced by protein diffusion or cell movement into deeper planes (Figure 5A).
	<ol>
		<li>To determine whether photoconversion is uniform, express a secreted photoconvertible protein (for experiments with extracellular fusion proteins) or a cytoplasmic photoconvertible protein (for experiments with intracellular fusion proteins). Photoconvert as usual, then acquire a z-stack encompassing most of the blastoderm every 20 min for 80 min.</li>
		<li>If photoconversion is biased toward the animal pole, the fluorescence intensity in deeper planes will increase over time due to diffusion or cell movement (Figure 5B). If non-uniform photoconversion is detected, focus deeper into the embryos during photoconversion.</li>
	</ol>
	</li>
</ol>

The authors have no conflicts to disclose.

The success of an FDAP experiment relies on the generation of a functional photoconvertible fusion protein. Tagging a protein can affect its biological activity and/or biophysical properties, including its localization, solubility, and stability36-41. Be prepared to test the activity of several different fusion constructs in order to find one that is active. We have found that changing the position of the photoconvertible protein relative to the protein of interest or using longer linkers (e.g., using...

The levels of proteins in cells and organisms are determined by their rates of production and clearance. Protein half-lives can range from minutes to days1-4. In many biological systems, the stabilization or clearance of key proteins has important effects on cellular activity. Modulation of intracellular protein stability is required for cell cycle progression5,6, developmental signaling7-9, apoptosis10, and normal function and maintenance of neurons11,12. Extracellular protein stability affects the distribution and availability of secreted proteins, such as morphogens13,14, within a tissue.
Over the last few decades, protein stability has mainly been assessed in cell culture using radioactive pulse-labeling or cycloheximide chase experiments15. In such pulse-chase experiments, cells are either transiently exposed to a &#8220;pulse&#8221; of radioactive amino acid precursors that are incorporated into newly synthesized proteins, or they are exposed to cycloheximide, which inhibits protein synthesis. Cultured cells are then collected at different time points, and either immunoprecipitation followed by autoradiography (in radioactive pulse-chase experiments) or western blotting (in cycloheximide experiments) is used to quantify the clearance of protein over time.
Conventional protein stability assays have several shortcomings. First, proteins in these assays are often not expressed in their endogenous environments, but rather in tissue culture and sometimes in cells from different species. For proteins whose stability is context-dependent, this approach is problematic. Second, it is not possible to follow protein clearance in individual cells or organisms over time, and the data from these assays reflects an average of different populations of cells at different time points. Since individual cells may have started with different amounts of protein, may have taken up the radioactive label or cycloheximide at different times, or may have different clearance kinetics, such aggregate data could be misleading. Finally, in the case of cycloheximide chase experiments, addition of the protein synthesis inhibitor may have unintended physiological effects that could artificially alter protein stability16-18. These shortcomings can be avoided by using Fluorescence Decay After Photoconversion (FDAP), a technique that utilizes photoconvertible proteins to measure protein clearance dynamically in living organisms19-25 (see Discussion for limitations of the FDAP technique).
Photoconvertible proteins are fluorescent proteins whose excitation and emission properties change after exposure to specific wavelengths of light26. One commonly used variant is Dendra2, a &#8220;green-to-red&#8221; photoconvertible protein that initially has excitation and emission properties similar to green fluorescent proteins, but after exposure to UV light&#8212;&#8220;photoconversion&#8221;&#8212;its excitation/emission properties become similar to those of red fluorescent proteins23,27. Importantly, new protein produced after photoconversion will not have the same excitation/emission properties as the photoconverted protein, allowing decoupling of production and clearance upon photoconversion and observation of only a pool of photoconverted protein. Tagging proteins of interest with photoconvertible proteins thus provides a convenient way to pulse-label proteins in intact, optically accessible living organisms.
In FDAP assays (Figure 1A), a protein of interest is tagged with a photoconvertible protein and expressed in a living organism (Figure 1B). The fusion protein is photoconverted, and the decrease in photoconverted signal over time is monitored by fluorescence microscopy (Figure 1C). The data is then fitted with an appropriate model to determine the half-life of the fusion protein (Figure 1D).
The FDAP assay described here was designed to determine the extracellular half-lives of secreted signaling proteins in zebrafish embryos during early embryogenesis19. However, this approach can be adapted to any transparent model organism that tolerates live imaging, and could be used to monitor the clearance of any taggable intracellular or extracellular protein. Variations of the technique described here have been performed in cultured cells20,23 and Drosophila22 and mouse21 embryos.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>PyFDAP (download from the following website: http://people.tuebingen.mpg.de/mueller-lab)</td>
			<td></td>
			<td></td>
			<td>Install and operate using the instructions provided on the PyFDAP website; PyFDAP is compatible with Linux, Mac, and Windows operating systems.</td>
		</tr>
		<tr>
			<td>mMessage mMachine Sp6 Transcription Kit (Life Technologies, AM1340)</td>
			<td></td>
			<td></td>
			<td>To generate capped mRNA for injection into embryos</td>
		</tr>
		<tr>
			<td>Alexa488-dextran conjugate, 3 kDa (Life Technologies, D34682)</td>
			<td></td>
			<td></td>
			<td>Co-inject with mRNA to create intracellular and extracellular masks</td>
		</tr>
		<tr>
			<td>6-well plastic dish (BD Falcon)</td>
			<td></td>
			<td></td>
			<td>Incubate embryos in agarose-coated wells until ready for mounting</td>
		</tr>
		<tr>
			<td>Embryo medium</td>
			<td></td>
			<td></td>
			<td>250 mg/l Instant Ocean salt, 1 mg/l methylene blue in reverse osmosis water adjusted to pH 7 with NaHCO3</td>
		</tr>
		<tr>
			<td>Protease from Streptomyces griseus (Sigma, P5147)</td>
			<td></td>
			<td></td>
			<td>Make a 5 mg/ml stock and use at 1 mg/ml to dechorionate embryos at the one-cell stage</td>
		</tr>
		<tr>
			<td>5 cm diameter glass petri dish</td>
			<td></td>
			<td></td>
			<td>For embryo dechorionation</td>
		</tr>
		<tr>
			<td>200 ml glass beaker</td>
			<td></td>
			<td></td>
			<td>For embryo dechorionation</td>
		</tr>
		<tr>
			<td>Microinjection apparatus</td>
			<td></td>
			<td></td>
			<td>For injection of mRNA and dye into embryos at the one-cell stage</td>
		</tr>
		<tr>
			<td>Stereomicroscope</td>
			<td></td>
			<td></td>
			<td>For injecting and mounting embryos</td>
		</tr>
		<tr>
			<td>1x Danieau&#39;s medium</td>
			<td></td>
			<td></td>
			<td>Dilute low melting point agarose and perform imaging in this medium; recipe: 0.2 mm filtered solution of 58 mM NaCl, 0.7 mM KCl, 0.4 mM MgSO4, 0.3 mM CaCl2, 5 mM HEPES pH 7.2</td>
		</tr>
		<tr>
			<td>UltraPure low melting point agarose (Invitrogen, 16520-100)</td>
			<td></td>
			<td></td>
			<td>For mounting embryos; use at a concentration of 1% in Danieau&#39;s medium: add 200 mg to 20 ml Danieau&#39;s medium, microwave until dissolved, then aliquot 1 ml into microcentrifuge tubes; aliquots can be stored at 4 &#176;C, re-melted at 70 &#176;C, and cooled to 40 - 42 &#176;C when ready to use</td>
		</tr>
		<tr>
			<td>Glass Pasteur pipette (Kimble Chase (via Fisher), 63A53WT)</td>
			<td></td>
			<td></td>
			<td>For mounting embryos; flame the tip to prevent jagged edges from injuring embryos</td>
		</tr>
		<tr>
			<td>Metal probe</td>
			<td></td>
			<td></td>
			<td>For positioning embryos during mounting</td>
		</tr>
		<tr>
			<td>Glass bottom dishes (MatTek, P35G-1.5-14-C)</td>
			<td></td>
			<td></td>
			<td>Use the appropriate cover glass thickness for your objective; part number listed here is for cover glass No. 1.5</td>
		</tr>
		<tr>
			<td>15 ml tube (BD Falcon) filled with ~5 ml embryo medium&#160;</td>
			<td></td>
			<td></td>
			<td>For rinsing residual agarose from the Pasteur pipette</td>
		</tr>
		<tr>
			<td>Inverted laser scanning confocal microscope</td>
			<td></td>
			<td></td>
			<td>A mercury arc lamp, 488 nm laser, 543 nm laser, and the appropriate filter sets are required</td>
		</tr>
		<tr>
			<td>Heated stage</td>
			<td></td>
			<td></td>
			<td>To maintain embryos at the optimal temperature of 28 &#176;C during the experiment</td>
		</tr>
		<tr>
			<td>Confocal software capable of time-lapse imaging</td>
			<td></td>
			<td></td>
			<td>Must be able to define multiple positions and automatically image them at defined intervals&#160;</td>
		</tr>
		<tr>
			<td>25x or 40x water objective</td>
			<td></td>
			<td></td>
			<td>Objective for imaging</td>
		</tr>
		<tr>
			<td>10x air objective</td>
			<td></td>
			<td></td>
			<td>Objective for photoconversion</td>
		</tr>
		<tr>
			<td>Immersion oil</td>
			<td></td>
			<td></td>
			<td>Immersion oil with the same refractive index as water</td>
		</tr>
	</tbody>
</table>

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.

FDAP has been used to determine the half-lives of extracellular signaling proteins in zebrafish embryos19. One of these proteins, Squint, induces expression of mesendodermal genes during embryogenesis34. Squint-Dendra2 activates expression of mesendodermal genes at levels similar to untagged Squint, as demonstrated by qRT-PCR and in situ hybridization assays19. Embryos were co-injected with Alexa488-dextran and mRNA encoding Squint-Dendra2 and subjected to the FDAP assay. A decre...

Watch this Scientific Journal Video about Measuring Protein Stability in Living Zebrafish Embryos Using Fluorescence Decay After Photoconversion FDAP at JoVE.com

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.

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 ...

measuring-protein-stability-living-zebrafish-embryos-using

Research

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Developmental Biology

11.2K vues.  Harvard University.  Les teneurs en protéines dans les cellules et les tissus sont souvent étroitement réglementées par l'équilibre de la production de protéines et de dégagement. En utilisant la fluorescence Decay Après Photoconversion (FDAP), la cinétique de dégagement de protéines peuvent être mesurées expérimentalement in vivo. 

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

Mechanical Vessel Injury in Zebrafish Embryos

Zebrafish (Danio rerio) embryos have proven to be a powerful model for studying a variety of developmental and disease processes. External development and optical transparency make these embryos especially amenable to microscopy, and numerous transgenic lines that label specific cell types with fluorescent proteins are available, making the zebrafish embryo an ideal system for visualizing the interaction of vascular, hematopoietic, and other cell types during injury and repair in vivo. Forward and reverse genetics in zebrafish are well developed, and pharmacological manipulation is possible. We describe a mechanical vascular injury model using micromanipulation techniques that exploits several of these features to study responses to vascular injury including hemostasis and blood vessel repair. Using a combination of video and timelapse microscopy, we demonstrate that this method of vascular injury results in measurable and reproducible responses during hemostasis and wound repair. This method provides a system for studying vascular injury and repair in detail in a whole animal model.

This article describes a method for creating a mechanical vessel injury in zebrafish embryos. This injury model provides a platform for studying hemostasis, injury-related inflammation, and wound healing in an organism ideally suited for real-time microscopy.

Zebrafish (Danio rerio) embryos have proven to be a powerful model for studying a variety of developmental and disease processes.  External development ...

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 ...

Analyzing In Vivo Cell Migration using Cell Transplantations and Time-lapse Imaging in Zebrafish Embryos

Cell migration is key to many physiological and pathological conditions, including cancer metastasis. The cellular and molecular bases of cell migration have been thoroughly analyzed in vitro. However, in vivo cell migration somehow differs from in vitro migration, and has proven more difficult to analyze, being less accessible to direct observation and manipulation. This protocol uses the migration of the prospective prechordal plate in the early zebrafish embryo as a model system to study the function of candidate genes in cell migration. Prechordal plate progenitors form a group of cells which, during gastrulation, undergoes a directed migration from the embryonic organizer to the animal pole of the embryo. The proposed protocol uses cell transplantation to create mosaic embryos. This offers the combined advantages of labeling isolated cells, which is key to good imaging, and of limiting gain/loss of function effects to the observed cells, hence ensuring cell-autonomous effects. We describe here how we assessed the function of the TORC2 component Sin1 in cell migration, but the protocol can be used to analyze the function of any candidate gene in controlling cell migration in vivo.

Analyzing In Vivo Cell Migration using Cell Transplantations and Time-lapse Imaging in Zebrafish Embryos

Combining cell transplantation, cytoskeletal labeling and loss/gain of function approaches, this protocol describes how the migrating zebrafish prospective prechordal plate can be used to analyze the function of a candidate gene in in vivo cell migration.

Cell migration is key to many physiological and pathological conditions, including cancer metastasis. The cellular and molecular bases of cell migration ...

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 ...

Biosensing Motor Neuron Membrane Potential in Live Zebrafish Embryos

The protocols described here are designed to allow researchers to study cell communication without altering the integrity of the environment in which the cells are located. Specifically, they have been developed to analyze the electrical activity of excitable cells, such as spinal neurons. In such a scenario, it is crucial to preserve the integrity of the spinal cell, but it is also important to preserve the anatomy and physiological shape of the systems involved. Indeed, the comprehension of the manner in which the nervous system-and other complex systems-works must be based on a systemic approach. For this reason, the live zebrafish embryo was chosen as a model system, and the spinal neuron membrane voltage changes were evaluated without interfering with the physiological conditions of the embryos. 
Here, an approach combining the employment of zebrafish embryos with a FRET-based biosensor is described. Zebrafish embryos are characterized by a very simplified nervous system and are particularly suited for imaging applications thanks to their transparency, allowing for the employment of fluorescence-based voltage indicators at the plasma membrane during zebrafish development. The synergy between these two components makes it possible to analyze the electrical activity of the cells in intact living organisms, without perturbing the physiological state. Finally, this non-invasive approach can co-exist with other analyses (e.g., spontaneous movement recordings, as shown here).

Protocols described here allow for the study of the electrical properties of excitable cells in the most non-invasive physiological conditions by employing zebrafish embryos in an in vivo system together with a fluorescence resonance energy transfer (FRET)-based genetically encoded voltage indicator (GEVI) selectively expressed in the cell type of interest.

The protocols described here are designed to allow researchers to study cell communication without altering the integrity of the environment in which the ...

Light Sheet-based Fluorescence Microscopy of Living or Fixed and Stained Tribolium castaneum Embryos

The red flour beetle Tribolium castaneum has become an important insect model organism in developmental genetics and evolutionary developmental biology. The observation of Tribolium embryos with light sheet-based fluorescence microscopy has multiple advantages over conventional widefield and confocal fluorescence microscopy. Due to the unique properties of a light sheet-based microscope, three dimensional images of living specimens can be recorded with high signal-to-noise ratios and significantly reduced photo-bleaching as well as photo-toxicity along multiple directions over periods that last several days. With more than four years of methodological development and a continuous increase of data, the time seems appropriate to establish standard operating procedures for the usage of light sheet technology in the Tribolium community as well as in the insect community at large. This protocol describes three mounting techniques suitable for different purposes, presents two novel custom-made transgenic Tribolium lines appropriate for long-term live imaging, suggests five fluorescent dyes to label intracellular structures of fixed embryos and provides information on data post-processing for the timely evaluation of the recorded data. Representative results concentrate on long-term live imaging, optical sectioning and the observation of the same embryo along multiple directions. The respective datasets are provided as a downloadable resource. Finally, the protocol discusses quality controls for live imaging assays, current limitations and the applicability of the outlined procedures to other insect species.
This protocol is primarily intended for developmental biologists who seek imaging solutions that outperform standard laboratory equipment. It promotes the continuous attempt to close the gap between the technically orientated laboratories/communities, which develop and refine microscopy methodologically, and the life science laboratories/communities, which require 'plug-and-play' solutions to technical challenges. Furthermore, it supports an axiomatic approach that moves the biological questions into the center of attention.

Light Sheet-based Fluorescence Microscopy of Living or Fixed and Stained Tribolium castaneum Embryos

Imaging the morphogenesis of insect embryos with light sheet-based fluorescence microscopy has become state of the art. This protocol outlines and compares three mounting techniques appropriate for Tribolium castaneum embryos, introduces two novel custom-made transgenic lines well suited for live imaging, discusses essential quality controls and indicates current experimental limitations.

The red flour beetle Tribolium castaneum has become an important insect model organism in developmental genetics and evolutionary developmental biology. ...

Induction of Hypoxia in Living Frog and Zebrafish Embryos

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and zebrafish embryos. Our system comprises a chamber featuring a simple setup that is nevertheless robust to induce and maintain a specific oxygen concentration and temperature in any experimental solution of choice. The presented system is very cost-effective but highly functional, it allows induction and sustainment of hypoxia for direct experiments in vivo and for various time periods up to 48 h.
To monitor and study the effects of hypoxia, we have employed two methods - measurement of levels of hypoxia-inducible factor 1alpha (HIF-1&#945;) in whole embryos or specific tissues and determination of retinal stem cell proliferation by 5-ethynyl-2&#8242;-deoxyuridine (EdU) incorporation into the DNA. HIF-1&#945; levels can serve as a general hypoxia marker in the whole embryo or tissue of choice, here embryonic retina. EdU incorporation into the proliferating cells of embryonic retina is a specific output of hypoxia induction. Thus, we have shown that hypoxic embryonic retinal progenitors decrease proliferation within 1 h of incubation under 5% oxygen of both frog and zebrafish embryos.
Once mastered, our setup can be employed for use with small aquatic model organisms, for direct in vivo experiments, any given time period and under normal, hypoxic or hyperoxic oxygen concentration or under any other given gas mixture.

We introduce a novel hypoxic chamber system for use with aquatic organisms such as frog and zebrafish embryos. Our system is simple, robust, cost-effective and allows the induction and sustainment of hypoxia in vivo and for up to 48 h. We present 2 reproducible methods to monitor the effectiveness of hypoxia.

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and ...

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 ...

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 ...

Whole Mount Immunohistochemistry in Zebrafish Embryos and Larvae

Immunohistochemistry is a widely used technique to explore protein expression and localization during both normal developmental and disease states. Although many immunohistochemistry protocols have been optimized for mammalian tissue and tissue sections, these protocols often require modification and optimization for non-mammalian model organisms. Zebrafish are increasingly used as a model system in basic, biomedical, and translational research to investigate the molecular, genetic, and cell biological mechanisms of developmental processes. Zebrafish offer many advantages as a model system but also require modified techniques for optimal protein detection. Here, we provide our protocol for whole-mount fluorescence immunohistochemistry in zebrafish embryos and larvae. This protocol additionally describes several different mounting strategies that can be employed and an overview of the advantages and disadvantages each strategy provides. We also describe modifications to this protocol to allow detection of chromogenic substrates in whole mount tissue and fluorescence detection in sectioned larval tissue. This protocol is broadly applicable to the study of many developmental stages and embryonic structures.

Here, we present a protocol for fluorescent antibody-mediated detection of proteins in whole preparations of zebrafish embryos and larvae.

Immunohistochemistry is a widely used technique to explore protein expression and localization during both normal developmental and disease states. ...