This study was supported by a National Institutes of Health (NIH) Core Grant (5P30EY008098) and the following funds to X.W.: NIH Grant R01EY016099 and Research to Prevent Blindness Career Development Award. We thank Kira Lathrop for assistance in confocal imaging.

Day 1. Preparation
1. Set up fish crosses
Set up pairwise crosses of zebrafish in the evening. To prevent random mating before the desired time, use dividers to separate the female and male fish in the mating cassettes. GFP-expressing transgenic fish are used as a source of donor blastomeres. The pt104 transgenic line (Zou et al., 2008), used here, expresses GFP ubiquitously under the control of the EF1 promoter. GFP expression allows visualization of donor cells by live imaging. Other transgenic fish lines that express suitable fluorescent proteins may be used for specific needs.
2. Prepare wedge-shaped agarose wells for embryo holding
Pour melted 1% agarose, prepared in E3 egg water (Westerfield, 2007), into a Petri dish. Float a plastic mold that has wedge-shaped protrusions on the agarose solution and let the agarose solution completely solidify. Remove the mold carefully and immerse the agarose bed with E3 water.
3. Prepare transplantation needles
Use a vertical Needle puller (David KOPF Instruments) at power setting 11.5 to make needles by pulling glass capillary pipettes (World Precision Instrument). Good needle tips should taper with long shafts. Grind the needle tips at a 45-degree angle with a Micro Grinder (Marishige) until beveled openings of 40-45&mu;m in inner diameter are obtained (Fig. 1A). To clean up the needle tips, attach a hose to the blunt end of the needle, connect the hose to a syringe, and then suck up and release a 10% hydrofluoric acid solution until all visible debris is removed. Next, wash the tips with distilled water, followed by 100% ethanol. Air-dry the needles until all ethanol evaporates. It is important to ensure that the tips are completely dry: residual ethanol will ruin the needles during the high-temperature polishing step. To soften and polish the rough edges of the needle tips, bake the needle tips with the heat generated by the filament of a Micro Forger (Marishige). Do not let the needle tips touch the filament during polishing. The amount of electricity passing though the filament and the distance between the filament and the needle tips can be adjusted to attain desired heating conditions. The temperature should not be too high: this will melt and deform the needle tips. After the rough edges are smoothened, carefully touch the heated filament with the needle tip and then gently pull the filament away from the needle to generate a pointed end (Fig. 1B). A sharp, smooth, and clean needle is critical for obtaining intact donor blastomeres and for not damaging the host embryos.
4. Prepare an embryo-positioning probe
Pull a glass pipette over a gas flame to obtain a fine glass probe with a smoothened end. This probe will be used to position the embryos in the proper orientation prior to transplantation.
Day 2. Blastomere transplantation
1. Collect and dechorionate embryos
The next day, remove the dividers in the cross cassettes to allow fish to mate. Place the embryos in a Petri dish filled with E3 water. Then, dechorionate embryos by manually tearing away the chorions from the embryos with a pair of fine forceps. Use a glass pipette with bent shaft to carefully transfer the dechorionated embryos into agarose wells. Avoid any contact of the embryos with the air, which cause the embryos to burst. Let the embryos develop at 28.5 &deg;C in the agarose wells until 3 hours post fertilization, or hpf.
2. Set up a transplantation apparatus
While the embryos develop, set up a transplantation apparatus by connecting the transplantation needle to the mineral oil-filled micro-pumping system. Make sure that the system does not leak and no air bubbles are trapped in the system. Back fill the entire transplantation needle with mineral oil, except for the 0.5 - 1 &mu;l of space at the tip opening.
3. Transplantation of blastomeres
To transplant blastomeres, use a glass positioning probe to orientate the blastomere side of the 3-4-hpf embryos upwards. Then, gently insert the transplantation needle into a donor embryo and slowly suck up 5-20 blastomeres into the needle. Insert the blastomere-filled needle into a host embryo and release the blastomeres slowly. The site at which the blastomeres are released will influence where their progeny localize at later developmental stages. To analyze retinal and brain development, release cells at the animal pole. During transplantation, make sure that the needle tip does not contact air. To minimize mechanical damages, host embryos shall be left in the agarose wells at 28.5 &deg;C until the next day. Then, transfer the mosaic embryos to 24-well plates pre-coated with 0.5 ml of 1% agarose, with one embryo in each well. 1 ml of E3 egg water is added to each well. To prevent bacterial infections, add 10 &mu;l of 100X penicillin and streptomycin to the E3 egg water.
Day 3. Embedding and live imaging of mosaic embryos
1. Embedding embryos
Prepare 1.5% low melting point agarose (LMP) in E3 egg water and keep the solution in a 30 &deg;C water bath. Transfer a mosaic embryo to a FluoroDish tissue culture dish with a glass bottom of 0.17mm in thickness (World Precision Instruments). Remove excess E3 water to leave the embryo just covered by a minimal amount of E3. Add 30-50 &mu;l melted 1.5% LMP to the embryo. Quickly position the eye or the brain as close to the glass bottom as possible before the agarose solution solidifies. After the solution cools down, add additional agarose solution to further secure the embryos in place. Then, immerse the solidified agarose block with 2-3 ml E3 egg water.
2. Confocal Imaging
For confocal microscopy, a water-immersion inverted objective (Olympus, PlanApo, 40 X/0.90 WLSM, Japan) with large focus depth is used. A drop of water is added onto the objective and then place a FluoroDish tissue culture dish on the objective. To image development, perform time-lapse imaging by taking pictures of the embryos every 5 minutes. Depending on the nature of experiments, the time intervals between shots should be adjusted.
Representative results
Figure 2 illustrates a representative 24-hpf mosaic zebrafish. GFP expression highlights the donor cells in green. The retinal neuroepithelial cells span the entire thickness of the retinal wall.
The movie shows the movements of donor cells in the host brain at 24 hpf. Notice cells morphologies change over time.
<img src="/files/ftp_upload/1924/1924fig1.jpg" alt="Figure 1" /> 
 Figure 1. Shapes of the transplantation needle tips. A. The beveled opening of the tip of a needle was free of debris after hydrofluoric acid washes. B. A pointed end was generated at the needle tip.
<img src="/files/ftp_upload/1924/1924fig2.jpg" alt="Figure 2" /> 
 Figure 2.Visualization of GFP-expressing donor cells in a chimeric zebrafish embryos at 24 hpf. The left panel shows the GFP signals in the brain and retina. The middle panel is a bright field picture of the embryo. The right panel shows the merged image.
Movie 1. Live imaging of GFP positive donor cells demonstrated the extensive changes in the morphologies of brain neuroepithelial cells during development. The individual frames of the movie were collected every 5 minutes between 24 and 25 hpf. <a href="https://www.jove.com/files/ftp_upload/1924/1924_Wei_Movie1.wmv">Click here to see the movie</a>.

<ol>
	<li>Zou, J., Lathrop, K., Sun, M., Wei, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intact+RPE+maintained+by+Nok+is+essential+for+retinal+epithelial+polarity+and+cellular+patterning+in+zebrafish.">Intact RPE maintained by Nok is essential for retinal epithelial polarity and cellular patterning in zebrafish.</a> Journal of Neuroscience. 28 (50), 13684-13695 (2008).</li><li>Westerfield, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+guide+for+the+laboratory+use+of+zebrafish+(Danio+rerio).">A guide for the laboratory use of zebrafish (Danio rerio).</a> THE ZEBRAFISH BOOK. , (2007).</li><li>Ho, R. K., Kane, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell-autonomous+action+of+zebrafish+spt-1+mutation+in+specific+mesodermal+precursors.">Cell-autonomous action of zebrafish spt-1 mutation in specific mesodermal precursors.</a> Nature. 348, 728-730 (1990).</li><li>Catalano, A., Raymond, P., Goldman, D., Wei, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zebrafish+dou+yan+Mutation+Causes+Patterning+Defects+and+Extensive+Cell+Death+in+the+Retina.">Zebrafish dou yan Mutation Causes Patterning Defects and Extensive Cell Death in the Retina.</a> Developmental Dynamics. 236 (5), 1295-1236 (2007).</li></ol>

In this video, we demonstrate a method for blastomere transplantation to analyze the retinal and brain development in zebrafish. This protocol can also be used to analyze the development of other tissues and organs by simply releasing the donor blastomeres to corresponding locations in the host embryos. The labeling with GFP molecules yields a cleaner background than labeling with fluorescent dye-conjugated Dextran (Ho and Kane 1990). This is because the debris of Dextran dyes from dead donor cells lasts much longer than...

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<td>5 &#190;" Glass pipette</td>
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<td>Borosilicate Glass Capillary</td>
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			E3 egg water: <ul> <li>5 mM NaCl</li> <li>0.17 mM KCl</li> <li>0.33 mM CaCl2</li> <li>0.33 mM MgSO4</li> <li>0.1% methylene blue</li> </ul>		</div>

transplantation of gfp-expressing blastomeres for live imaging of retinal and brain development in chimeric zebrafish embryos

Cells change extensively in their locations and property during embryogenesis. These changes are regulated by the interactions between the cells and their environment. Chimeric embryos, which are composed of cells of different genetic background, are great tools to study the cell-cell interactions mediated by genes of interest. The embryonic transparency of zebrafish at early developmental stages permits direct visualization of the morphogenesis of tissues and organs at the cellular level. Here, we demonstrate a protocol to generate chimeric retinas and brains in zebrafish embryos and to perform live imaging of the donor cells. The protocol covers the preparation of transplantation needles, the transplantation of GFP-expressing donor blastomeres to GFP-negative hosts, and the examination of donor cell behavior under live confocal microscopy. With slight modifications, this protocol can also be used to study the embryonic development of other tissues and organs in zebrafish. The advantages of using GFP to label donor cells are also discussed. 

Watch this Scientific Journal Video about Transplantation of GFP-expressing Blastomeres for Live Imaging of Retinal and Brain Development in Chimeric Zebrafish Embryos at JoVE.com

Transplantation of GFP-expressing Blastomeres for Live Imaging of Retinal and Brain Development in Chimeric Zebrafish Embryos

We demonstrate a protocol to generate chimeric zebrafish embryos for live imaging cellular behavior during embryogenesis.

Cells change extensively in their locations and property during embryogenesis. These changes are regulated by the interactions between the cells and their ...

transplantation-gfp-expressing-blastomeres-for-live-imaging-retinal

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10.6K Views.  University of Pittsburgh. We demonstrate a protocol to generate chimeric zebrafish embryos for live imaging cellular behavior during embryogenesis. 

Video: Transplantation of GFP-expressing Blastomeres for Live Imaging of Retinal and Brain Development in Chimeric Zebrafish Embryos

Two-photon axotomy and time-lapse confocal imaging in live zebrafish embryos

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo. Here we describe a method to mount zebrafish embryos for long-term imaging and demonstrate how to automate the capture of time-lapse images using a confocal microscope. We also describe a method to create controlled, precise damage to individual branches of peripheral sensory axons in zebrafish using the focused power of a femtosecond laser mounted on a two-photon microscope. The parameters for successful two-photon axotomy must be optimized for each microscope. We will demonstrate two-photon axotomy on both a custom built two-photon microscope and a Zeiss 510 confocal/two-photon to provide two examples.

Zebrafish trigeminal sensory neurons can be visualized in a transgenic line expressing GFP driven by a sensory neuron specific promoter 1. We have adapted this zebrafish trigeminal model to directly observe sensory axon regeneration in living zebrafish embryos. Embryos are anesthetized with tricaine and positioned within a drop of agarose as it solidifies. Immobilized embryos are sealed within an imaging chamber filled with phenylthiourea (PTU) Ringers. We have found that embryos can be continuously imaged in these chambers for 12-48 hours. A single confocal image is then captured to determine the desired site of axotomy. The region of interest is located on the two-photon microscope by imaging the sensory axons under low, non-damaging power. After zooming in on the desired site of axotomy, the power is increased and a single scan of that defined region is sufficient to sever the axon. Multiple location time-lapse imaging is then set up on a confocal microscope to directly observe axonal recovery from injury.

Here we describe a method for mounting zebrafish embryos for long-term imaging, two-photon imaging and tissue-damage techniques, and time-lapse confocal imaging.

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo.  ...

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the developing zebrafish brain (Gutzman et al, 2008). Such analysis affords a new understanding of the underlying cell biology required for development of the 3D structure of the vertebrate brain, and significantly increases our ability to study neural tube morphogenesis. The embryonic zebrafish brain is shaped beginning at 18 hours post fertilization (hpf) as the ventricles within the neuroepithelium inflate. By 24 hpf, the initial steps of neural tube morphogenesis are complete. Using the method described here, embryos at the one cell stage are injected with mRNA encoding membrane-targeted green fluorescent protein (memGFP). After injection and incubation, the embryo, now between 18 and 24 hpf, is mounted, inverted, in agarose and imaged by confocal microscopy. Notably, the zebrafish embryo is transparent making it an ideal system for fluorescent imaging. While our analyses have focused on the midbrain-hindbrain boundary and the hindbrain, this method could be extended for analysis of any region in the zebrafish to a depth of 
80-100 &mu;m.

In this video, we demonstrate a method by which to analyze the developing vertebrate brain in live zebrafish embryos at single cell resolution by confocal microscopy. This includes the method by which we inject the single-cell zebrafish embryo and subsequently mount and image the developing brain.

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the ...

Generating Chimeric Zebrafish Embryos by Transplantation

One of the most powerful tools used to gain insight into complex developmental processes is the analysis of chimeric embryos. A chimera is defined as an organism that contains cells from more than one animal; mosaics are one type of chimera in which cells from more than one genotype are mixed, usually wild-type and mutant. In the zebrafish, chimeras can be readily made by transplantation of cells from a donor embryo into a host embryo at the appropriate embryonic stage. Labeled donor cells are generated by injection of a lineage marker, such as a fluorescent dye, into the one-cell stage embryo. Labeled donor cells are removed from donor embryos and introduced into unlabeled host embryos using an oil-controlled glass pipette mounted on either a compound or dissecting microscope. Donor cells can in some cases be targeted to a specific region or tissue of the developing blastula or gastrula stage host embryo by choosing a transplantation site in the host embryo based on well-established fate maps.

A step-by-step guide to generating targeted chimeric zebrafish embryos by transplantation at the blastula or gastrula stage.

One of the most powerful tools used to gain insight into complex developmental processes is the analysis of chimeric embryos. A chimera is defined as an ...

Live Imaging of Cell Motility and Actin Cytoskeleton of Individual Neurons and Neural Crest Cells in Zebrafish Embryos

The zebrafish is an ideal model for imaging cell behaviors during development in vivo. Zebrafish embryos are externally fertilized and thus easily accessible at all stages of development. Moreover, their optical clarity allows high resolution imaging of cell and molecular dynamics in the natural environment of the intact embryo. We are using a live imaging approach to analyze cell behaviors during neural crest cell migration and the outgrowth and guidance of neuronal axons.
Live imaging is particularly useful for understanding mechanisms that regulate cell motility processes. To visualize details of cell motility, such as protrusive activity and molecular dynamics, it is advantageous to label individual cells. In zebrafish, plasmid DNA injection yields a transient mosaic expression pattern and offers distinct benefits over other cell labeling methods. For example, transgenic lines often label entire cell populations and thus may obscure visualization of the fine protrusions (or changes in molecular distribution) in a single cell. In addition, injection of DNA at the one-cell stage is less invasive and more precise than dye injections at later stages.
Here we describe a method for labeling individual developing neurons or neural crest cells and imaging their behavior in vivo. We inject plasmid DNA into 1-cell stage embryos, which results in mosaic transgene expression. The vectors contain cell-specific promoters that drive expression of a gene of interest in a subset of sensory neurons or neural crest cells. We provide examples of cells labeled with membrane targeted GFP or with a biosensor probe that allows visualization of F-actin in living cells1.
Erica Andersen, Namrata Asuri, and Matthew Clay contributed equally to this work.

This protocol describes imaging of individual neurons or neural crest cells in living zebrafish embryos. This method is used to examine cellular behaviors and actin localization using fluorescence confocal time-lapse microscopy.

The zebrafish is an ideal model for imaging cell behaviors during development in vivo. Zebrafish embryos are externally fertilized and thus easily ...

Development of automated imaging and analysis for zebrafish chemical screens.

We demonstrate the application of image-based high-content screening (HCS) methodology to identify small molecules that can modulate the FGF/RAS/MAPK pathway in zebrafish embryos. The zebrafish embryo is an ideal system for in vivo high-content chemical screens. The 1-day old embryo is approximately 1mm in diameter and can be easily arrayed into 96-well plates, a standard format for high throughput screening. During the first day of development, embryos are transparent with most of the major organs present, thus enabling visualization of tissue formation during embryogenesis. The complete automation of zebrafish chemical screens is still a challenge, however, particularly in the development of automated image acquisition and analysis. We previously generated a transgenic reporter line that expresses green fluorescent protein (GFP) under the control of FGF activity and demonstrated their utility in chemical screens 1. To establish methodology for high throughput whole organism screens, we developed a system for automated imaging and analysis of zebrafish embryos at 24-48 hours post fertilization (hpf) in 96-well plates 2. In this video we highlight the procedures for arraying transgenic embryos into multiwell plates at 24hpf and the addition of a small molecule (BCI) that hyperactivates FGF signaling 3. The plates are incubated for 6 hours followed by the addition of tricaine to anesthetize larvae prior to automated imaging on a Molecular Devices ImageXpress Ultra laser scanning confocal HCS reader. Images are processed by Definiens Developer software using a Cognition Network Technology algorithm that we developed to detect and quantify expression of GFP in the heads of transgenic embryos. In this example we highlight the ability of the algorithm to measure dose-dependent effects of BCI on GFP reporter gene expression in treated embryos.

We report the development of a system for automated imaging and analysis of zebrafish transgenic embryos in multiwell plates. This demonstrates the ability to measure dose dependent effects of a small molecule, BCI, on Fibroblast Growth Factor reporter gene expression and provide technology for establishing high-throughput zebrafish chemical screens.

We demonstrate the application of image-based high-content screening (HCS) methodology to identify small molecules that can modulate the FGF/RAS/MAPK ...

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 ...

Dechorionation of Medaka Embryos and Cell Transplantation for the Generation of Chimeras

Medaka is a small egg-laying freshwater fish that allows both genetic and embryological analyses and is one of the three vertebrate model organisms in which genome-wide phenotype-driven mutant screens were carried out 1. Divergence of functional overlap of related genes between medaka and zebrafish allows identification of novel phenotypes that are unidentifiable in a single species 2, thus medaka and zebrafish are complementary for genetic dissection of the vertebrate genome functions. Manipulation of medaka embryos, such as dechorionation, mounting embryos for imaging and cell transplantation, are key procedures to work on both medaka and zebrafish in a laboratory. Cell transplantation examines cell autonomy of medaka mutations. Chimeras are generated by transplanting labeled cells from donor embryos into unlabeled recipient embryos. Donor cells can be transplanted to specific areas of the recipient embryos based on the fate maps 3 so that clones from transplanted cells can be integrated in the tissue of interest during development. Due to the hard chorion and soft embryos, manipulation of medaka embryos is more involved than in zebrafish. In this video, we show detailed procedures to manipulate medaka embryos.

Due to the hard chorion and soft embryos, manipulation of medaka embryos is more involved than in zebrafish. This video shows step-by-step procedures for how to manipulate medaka embryos, including dechorionation, mounting in agarose for imaging and cell transplantation for the production of chimeras. These procedures are essential to use medaka and zebrafish in a laboratory to take full advantage of their complementary features for the genetic dissection of vertebrate genome functions.

Medaka is a small egg-laying freshwater fish that allows both genetic and embryological analyses and is one of the three vertebrate model organisms in ...

Imaging Glycans in Zebrafish Embryos by Metabolic Labeling and Bioorthogonal Click Chemistry

Imaging glycans in vivo has recently been enabled using a bioorthogonal chemical reporter strategy by treating cells or organisms with azide- or alkyne-tagged monosaccharides1, 2. The modified monosaccharides, processed by the glycan biosynthetic machinery, are incorporated into cell surface glycoconjugates. The bioorthogonal azide or alkyne tags then allow covalent conjugation with fluorescent probes for visualization, or with affinity probes for enrichment and glycoproteomic analysis. This protocol describes the procedures typically used for noninvasive imaging of fucosylated glycans in zebrafish embryos, including: 1) microinjection of one-cell stage embryos with GDP-5-alkynylfucose (GDP-FucAl), 2) labeling fucosylated glycans in the enveloping layer of zebrafish embryos with azide-conjugated fluorophores via biocompatible Cu(I)-catalyzed azide-alkyne cycloaddition (CuAAC), and 3) imaging by confocal microscopy3. The method described here can be readily extended to visualize other classes of glycans, e.g. glycans containing sialic acid4 and N-acetylgalactosamine5, 6, in developing zebrafish and in other living organisms.

A click-chemistry based method that allows for the rapid, noninvasive, and robust labeling of alkyne-tagged glycans in zebrafish embryos is described. Fucosylated glycans in the enveloping layer of zebrafish embryos in the late gastrulation stage were imaged in this study.

Imaging glycans in vivo has recently been enabled using a bioorthogonal chemical reporter strategy by treating cells or organisms with azide- or ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

Analysis of Oxidative Stress in Zebrafish Embryos

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

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