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><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</td><td>Life Technologies</td><td>AM1340</td><td>To generate capped mRNA for injection into embryos</td></tr><tr><td>Alexa488-dextran conjugate, 3 kDa</td><td>Life Technologies</td><td>D34682</td><td>Co-inject with mRNA to create intracellular and extracellular masks</td></tr><tr><td>6-well plastic dish</td><td>BD Falcon</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&amp;nbsp;mg/L methylene blue in reverse osmosis water adjusted to pH 7 with NaHCO&lt;sub&gt;3&lt;/sub&gt;</td></tr><tr><td>Protease from &lt;em&gt;Streptomyces griseus&lt;/em&gt;</td><td>Sigma</td><td>P5147</td><td>Make a 5 mg/ml stock and use at 1&amp;nbsp;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&#039;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 MgSO&lt;sub&gt;4&lt;/sub&gt;, 0.3 mM CaCl&lt;sub&gt;2&lt;/sub&gt;, 5&amp;nbsp;mM HEPES pH 7.2</td></tr><tr><td>UltraPure low melting point agarose</td><td>Invitrogen</td><td>16520-100</td><td>For mounting embryos; use at a concentration of 1% in Danieau&#039;s medium: add 200 mg to 20 ml Danieau&#039;s medium, microwave until dissolved, then aliquot 1&amp;nbsp;ml into microcentrifuge tubes; aliquots can be stored at 4 &amp;deg;C, re-melted at 70&amp;nbsp;&amp;deg;C, and cooled to 40&amp;#8211;42 &amp;deg;C when ready to use</td></tr><tr><td>Glass Pasteur pipette</td><td>Kimble Chase (via Fisher)</td><td>63A53WT</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</td><td>MatTek</td><td>P35G-1.5-14-C</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 filled with ~5 ml embryo medium</td><td>BD Falcon</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&amp;nbsp;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 &amp;deg;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&amp;nbsp;</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

JoVE Journal

Developmental Biology

11.2K Views.  Harvard University.  Eiwitniveaus in cellen en weefsels vaak zijn gereguleerd door het evenwicht van eiwitproductie en klaring. Met behulp van fluorescentie Decay Na Photoconversion (FDAP), kan de klaringskinetiek van eiwitten experimenteel worden gemeten in vivo. 

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

Tracking Cells in GFP-transgenic Zebrafish Using the Photoconvertible PSmOrange System

The rapid development of transparent zebrafish embryos (Danio rerio) in combination with fluorescent labelings of cells and tissues allows visualizing developmental processes as they happen in the living animal. Cells of interest can be labeled by using a tissue specific promoter to drive the expression of a fluorescent protein (FP) for the generation of transgenic lines. Using fluorescent photoconvertible proteins for this purpose additionally allows to precisely follow defined structures within the expression domain. Illuminating the protein in the region of interest, changes its emission spectrum and highlights a particular cell or cell cluster leaving other transgenic cells in their original color. A major limitation is the lack of known promoters for a large number of tissues in the zebrafish. Conversely, gene- and enhancer trap screens have generated enormous transgenic resources discretely labeling literally all embryonic structures mostly with GFP or to a lesser extend red or yellow FPs. An approach to follow defined structures in such transgenic backgrounds would be to additionally introduce a ubiquitous photoconvertible protein, which could be converted in the cell(s) of interest. However, the photoconvertible proteins available involve a green and/or less frequently a red emission state1 and can therefore often not be used to track cells in the FP-background of existing transgenic lines. To circumvent this problem, we have established the PSmOrange system for the zebrafish2,3. Simple microinjection of synthetic mRNA encoding a nuclear form of this protein labels all cell nuclei with orange/red fluorescence. Upon targeted photoconversion of the protein, it switches its emission spectrum to far red. The quantum efficiency and stability of the protein makes PSmOrange a superb cell-tracking tool for zebrafish and possibly other teleost species.

We established the photoconvertible PSmOrange system as a powerful, straight-forward and cost inexpensive tool for in vivo cell tracking in GFP transgenic backgrounds. This protocol describes its application in the zebrafish model system.

The rapid development of transparent zebrafish embryos (Danio rerio) in combination with fluorescent labelings of cells and tissues allows visualizing ...

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 ...

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 ...

Fluorescent Protein Tagging in Zebrafish Larvae Using CRISPR/Cas9 Knock-In

Fluorescence Labeling to Visualize Low-Expressed Proteins in Zebrafish

CRISPR/Cas9-mediated knock-in (KI) technology allows for easier fluorescent-protein tagging in zebrafish (Danio rerio), a preferred model organism for in vivo imaging due to its transparency during the early developmental stage. Here, we provide a detailed protocol for performing high-efficiency fluorescence gene KI, rapid screening for KI founders, and low-abundance protein tracing in zebrafish larvae, which will lay a critical foundation for subsequent physio-pathological studies in zebrafish. The current protocol includes complete steps for the sgRNA design for the gene of interest, sgRNA in vitro transcription, Cas9 mRNA in vitro transcription, in vivo sgRNA screen for the one with the highest efficiency, donor plasmid design and construction, microinjection in zebrafish larvae, KI founder screen and zebrafish live imaging. Critical steps, troubleshooting tips, quality control methods, and advantages and applications of this protocol are included and discussed. This protocol assures quick and accurate results at a low cost and has been validated by multiple trials.

Here, we present a protocol to label proteins with a fluorescence protein tag in zebrafish larvae, a newly developed and modified in vivo system especially useful for visualizing low-abundance proteins in zebrafish.

CRISPR/Cas9-mediated knock-in (KI) technology allows for easier fluorescent-protein tagging in zebrafish (Danio rerio), a preferred model organism for in ...

Genetics

Two-Photon-Based Photoactivation in Live Zebrafish Embryos

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

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

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

Biology

A Simple and Effective Transplantation Device for Zebrafish Embryos

Classical embryological manipulations, such as removing cells and transplanting cells within or between embryos, are powerful techniques to study complex developmental processes. Zebrafish embryos are ideally suited for these manipulations since they are easily accessible, relatively large in size, and transparent. However, previously developed devices for cell removal and transplantation are cumbersome to use or expensive to purchase. In contrast, the transplantation device presented here is economical, easy to assemble, and simple to use. In this protocol, we first introduce the handling of the transplantation device as well as its assembly from commercially and widely available parts. We then present three applications for its use: generation of ectopic clones to study signal dispersal from localized sources, extirpation of cells to produce size-reduced embryos, and germline transplantation to generate maternal-zygotic mutants. Finally, we show that the tool can also be used for embryological manipulations in other species such as the Japanese rice fish medaka.

Embryological manipulations such as extirpation and transplantation of cells are important tools to study early development. This protocol describes a simple and effective transplantation device to perform these manipulations in zebrafish embryos.

Classical embryological manipulations, such as removing cells and transplanting cells within or between embryos, are powerful techniques to study complex ...

Single Cell Fate Mapping in Zebrafish

The ability to differentially label single cells has important implications in developmental biology. For instance, determining how hematopoietic, lymphatic, and blood vessel lineages arise in developing embryos requires fate mapping and lineage tracing of undifferentiated precursor cells. Recently, photoactivatable proteins which include: Eos1, 2, PAmCherry3, Kaede4-7, pKindling8, and KikGR9, 10 have received wide interest as cell tracing probes. The fluorescence spectrum of these photosensitive proteins can be easily converted with UV excitation, allowing a population of cells to be distinguished from adjacent ones. However, the photoefficiency of the activated protein may limit long-term cell tracking11. As an alternative to photoactivatable proteins, caged fluorescein-dextran has been widely used in embryo model systems7, 12-14. Traditionally, to uncage fluorescein-dextran, UV excitation from a fluorescence lamp house or a single photon UV laser has been used; however, such sources limit the spatial resolution of photoactivation. Here we report a protocol to fate map, lineage trace, and detect single labeled cells. Single cells in embryos injected with caged fluorescein-dextran are photoactivated with near-infrared laser pulses produced from a titanium sapphire femtosecond laser. This laser is customary in all two-photon confocal microscopes such as the LSM 510 META NLO microscope used in this paper. Since biological tissue is transparent to near-infrared irradiation15, the laser pulses can be focused deep within the embryo without uncaging cells above or below the selected focal plane. Therefore, non-linear two-photon absorption is induced only at the geometric focus to uncage fluorescein-dextran in a single cell. To detect the cell containing uncaged fluorescein-dextran, we describe a simple immunohistochemistry protocol16 to rapidly visualize the activated cell. The activation and detection protocol presented in this paper is versatile and can be applied to any model system. 
Note: The reagents used in this protocol can be found in the table appended at the end of the article.

A method is described to photoactivate single cells containing a caged fluorescent protein using two-photon absorption from a Ti:Sapphire femtosecond laser oscillator. To fate map the photoactivated cell, immunohistochemistry is used. This technique can be applied to any cell type.

The ability to differentially label single cells has important implications in developmental biology. For instance, determining how hematopoietic, ...

An Efficient Protocol to Assess ERK Activity Modulation in Early Zebrafish Noonan Syndrome Models via Live FRET Microscopy and Immunofluorescence

RASopathies are genetic syndromes caused by ERK hyperactivation and resulting in multisystemic diseases that can also lead to cancer predisposition. Despite a broad genetic heterogeneity, germline gain-of-function mutations in key regulators of the RAS-MAPK pathway underlie the majority of the cases, and, thanks to advanced sequencing techniques, potentially pathogenic variants affecting the RAS-MAPK pathway continue to be identified. Functional validation of the pathogenicity of these variants, essential for accurate diagnosis, requires fast and reliable protocols, preferably in vivo. Given the scarcity of effective treatments in early childhood, such protocols, especially if scalable in cost-effective animal models, can be instrumental in offering a preclinical ground for drug repositioning/repurposing. 
Here we describe step-by-step the protocol for rapid generation of transient RASopathy models in zebrafish embryos and direct inspection of live disease-associated ERK activity changes occurring already during gastrulation through real-time multispectral F&#246;rster resonance energy transfer (FRET) imaging. The protocol uses a transgenic ERK reporter recently established and integrated with the hardware of commercial microscopes. We provide an example application for Noonan syndrome (NS) zebrafish models obtained by expression of the Shp2D61G. We describe a straightforward method that enables registration of ERK signal change in the NS fish model before and after pharmacological signal modulation by available low-dose MEK inhibitors. We detail how to generate, retrieve, and assess ratiometric FRET signals from multispectral acquisitions before and after treatment and how to cross-validate the results via classical immunofluorescence on whole embryos at early stages. We then describe how, via examining standard morphometric parameters, to query late changes in embryo shape, indicative of a resulting impairment of gastrulation, in the same embryos whose ERK activity is assessed by live FRET at 6 h post fertilization.

RASopathies are multisystem genetic syndromes caused by RAS-MAPK pathway hyperactivation. Potentially pathogenic variants awaiting validation emerge continuously while poor preclinical evidence limits therapy. Here, we describe our in vivo protocol to test and cross-validate RASopathy-associated ERK activation levels and its pharmacological modulation during embryogenesis by live FRET imaging in Teen-reporter zebrafish.

RASopathies are genetic syndromes caused by ERK hyperactivation and resulting in multisystemic diseases that can also lead to cancer predisposition. ...

Biochemistry

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 ...

Cell Tracking Using Photoconvertible Proteins During Zebrafish Development

Embryogenesis is a dynamic process that is best studied by using techniques that allow the documentation of developmental changes in vivo. The use of genetically-encoded fluorescent proteins has proven a valuable strategy for elucidating dynamic morphogenetic processes as they occur in the intact organism. During the past decade, the development of photoactivatable and photoconvertible fluorescent proteins has opened the possibility to investigate the fate of discrete subpopulations of tagged proteins1. Unlike photoactivatable proteins, photoconvertible fluorescent proteins (PCFPs) are readily tracked and imaged in their native emission state prior to photoconversion, making it easier to identify and select regions by optical inspection. PCFPs, such as Kaede2, KikGR3, Dendra4 and EosFP5, can be shifted from green to red upon exposure to UV or blue light due to a His-Tyr-Gly tripeptide sequence which forms a green chromophore that can be photoconverted to a red one by a light-catalyzed &beta;-elimination and subsequent extension of a &pi;-conjugated system3. PCFPs and their monomeric variants are useful tools for tracking cells6-10 and studying protein dynamics11-14, respectively. During recent years, PCFPs have been expressed in different animal model, such as zebrafish6, chicken7,8 and mouse9,10 for cell fate tracking. Here we report a protocol for cell-specific photoconversion of PCFPs in the living zebrafish embryo and further tracking of photoconverted proteins at later developmental stages. This methodology allows studying, in a tissue-specific manner, cell biological events underlying morphogenesis in the zebrafish animal model.

Here, we present a method for the photoactivated switch of photoconvertible fluorescent proteins (PCFPs) in the living zebrafish embryo and further tracking of photoconverted protein at specific time points during development. This methodology allows monitoring of cell biological events underlying different developmental processes in a live vertebrate organism.

Embryogenesis is a dynamic process that is best studied by using techniques that allow the documentation of developmental changes in vivo. The use of ...

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

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