We thank F. Bouallague and the IBENS animal facility for excellent zebrafish care. Research reported in this publication was supported by the Fondation ARC pour la recherche sur le cancer, grants N&#176; SFI20111203770 and N&#176; PJA 20131200143.

Note: Figure 1 presents the outline of the protocol.
1. Preparation of the Needles for Injection and Transplantation
Note: Needles can be prepared at any time and stored. Keep them in a Petri dish, on a band of modeling clay. Seal the dish with parafilm to protect from dust.
<ol>
	<li>For injection needles, pull a glass capillary (outside diameter 1.0 mm, inside diameter 0.58 mm, without filament (see list of Materials)) with a micropipette puller (see list of Materials). Prefer short and thin needles (tapered part of about 5 mm) as they are more efficient for piercing the chorion. 
	Note: The injection needles are single-use.</li>
	<li>Needle for Cell Transplantation
	<ol>
		<li>Pull a glass capillary (outside diameter 1.0 mm, inside diameter 0.78 mm, without filament (see list of Materials)) with a micropipette puller.</li>
		<li>Using a microforge (see list of materials), cut the tip of the needle at the point where the inside diameter is 35 &#181;m (slightly more than the diameter of cells to be transplanted).</li>
		<li>Bevel the cut extremity of the capillary with a microgrinder (see list of Materials) with an angle of 45&#176;.</li>
		<li>Optionally, pull a barb on the end of the needle using a microforge. For this, touch the hot filament with the tip of the bevel and rapidly pull it away, creating a barb that can help penetrating the embryo.</li>
		<li>Mount the needle on a needle holder attached to a syringe. Using the syringe, rinse the inside of the needle three times with 2% hydrofluoric acid to eliminate glass residues, and then rinse three times with acetone. 
		Caution: hydrofluoric acid is toxic, manipulate under the hood, with gloves. 
		Note: The cell transplantation needles can be used several times.</li>
	</ol>
	</li>
</ol>
2. Preparation of the Dishes for Injection and Cell Transplantation
<ol>
	<li>Heat 50 ml of embryo medium11 containing 0.5 g of agarose in a microwave for 1 min at full power to get 1% agarose in embryo medium. Cool the agarose gel to about 60 &#176;C and pour the 50 ml in a 90 mm Petri dish. Place at the surface of the gel a specially designed mold (see list of material), either with lines for the injection dish or with wells for the cell transplantation dish.
	<ol>
		<li>Observe the mold float on the agarose, if the agarose is not too hot. After the agarose gel is set, remove the mold with forceps (Figure 2A-B). 
		Note: dishes can be stored at 4 &#176;C, up to a few weeks. Keep them in a closed wet box, to avoid drying.</li>
	</ol>
	</li>
	<li>Place the dish in a 28 &#176;C incubator 30 min before use.</li>
</ol>
3. Collection of Embryos and Injection
<ol>
	<li>Injection Solution Preparation 
	Note: Actin binding domain of the yeast actin binding protein 140 (ABP-140) such as Lifeact bound to mCherry (referred to as ABP140-mCherry) is used to visualize polymerized actin within the cell, in order to analyze the protrusive activity. Add another compound to test the function of a candidate gene (e.g. mRNA of dominant negative or constitutively active construct, or Morpholino oligonucleotides). To assess the functions of Sin1 as described in the results (Figure 3), we used Morpholino oligonucleotides. As a control, we used a morpholino identical to the sin1 morpholino but for 5 nucleotides distributed along the sequence so that this control morpholino doesn&#39;t match the target RNA.
	<ol>
		<li>Thaw sin1 and 5-mismatch control morpholinos (2 mM stock solutions), and ABP140-mCherry mRNAs on ice. Add 375 ng of mRNAs (75 ng/&#181;l final) and 0.75 &#181;l of either sin1 or 5-mismatch control morpholino (0.3 mM final) in Danieau buffer (58 mM NaCl, 0.7 mM KCl, 0.4 mM MgSO4, 0.6 mM Ca(NO3)2, 5.0 mM HEPES pH 7.6), to reach a total volume of 5 &#181;l.</li>
	</ol>
	</li>
	<li>Embryo Collection 
	Note: for this experimental set up both donor and host embryos are transgenic, expressing the GFP under the control of the goosecoid (gsc) promoter in order to visualize the prospective prechordal plate11.
	<ol>
		<li>Collect Tg(gsc:gfp) eggs. Place about 80 eggs in a 90 mm Petri dish filled with embryo medium and incubate at 28 &#176;C.</li>
	</ol>
	</li>
	<li>Inject Donor Embryos
	<ol>
		<li>Under a stereomicroscope, gently squeeze collected embryos with forceps into the lines of an injection dish filled with embryo medium. Using forceps, orient the embryos with the cells towards the top. Place the injection dish in the incubator at 28 &#176;C until the embryos have reached the 4-cell stage (1 hr post fertilization).</li>
		<li>Load an injection needle with 2 &#181;l of injection solution. Insert the needle in a needle holder (see list of materials) that is placed in a micro-manipulator (see list of materials) and connected with polytetrafluoroethylene (PTFE) tubing (see list of Materials) to an air transjector (see list of materials).</li>
		<li>Open the capillary by gently touching the tip with fine forceps. If necessary, cut the capillary open by pinching its extremity.</li>
		<li>Enter one of the four cells with the needle and inject the solution within the cell in order to fill 70% of the cell volume (2 nl). Inject 30 embryos. 
		Note: Getting the needle in the cell may be difficult, as the plasma membrane is soft. Aiming at the junction between the cell and the yolk facilitates needle penetration.</li>
		<li>Place the embryos in the 28 &#176;C incubator until they have reached the sphere stage (4 hours post fertilization).</li>
	</ol>
	</li>
</ol>
4. Preparation of the Embryos for the Cell Transplantation
<ol>
	<li>Prepare 20 ml of embryo medium with 200 &#181;l penicillin/streptomycin (see list of materials) (Pen/Strep EM).</li>
	<li>Dechorionation of the embryos at the sphere stage (4 hr post fertilization) 
	Note: Dechorionated embryos are very fragile and will explode in case of contact with air or plastic. They thus need to be placed in agarose-coated dishes, and transferred with fire-polished glass Pasteur pipettes.
	<ol>
		<li>Using a plastic Pasteur pipette, transfer host embryos collected at step 3.2 and donor embryos injected at step 3.3 into two 35 mm Petri dishes coated with 1% agarose in embryo medium and filled with Pen/Strep EM. Keep host embryos and donor embryos in two separate dishes.</li>
		<li>Manually extract the embryos from their chorions with fine tweezers (see list of materials).</li>
		<li>Place the embryos in the 28 &#176;C incubator until they have reached the shield stage (6 hr post fertilization).</li>
	</ol>
	</li>
</ol>
5. Cell Transplantation
<ol>
	<li>Arrange the Embryos for the cell Transplantation.
	<ol>
		<li>Under a fluorescent stereomicroscope, select donor embryos expressing ABP140-mCherry (red) within the shield (green).</li>
		<li>Using a fire-polished Pasteur pipette, transfer the embryos in the wells of the cell transplantation dish filled with Pen/Strep EM. Place the host embryos in one row and the donor embryos in the neighboring row.</li>
		<li>Using an eyelash, carefully orient the embryos with the shield up. 
		Note: the position of the shield can be assessed by GFP expression or anatomically.</li>
	</ol>
	</li>
	<li>Transplantation System Set-up. 
	Note: The transplantation system is composed of a needle holder (see list of materials) connected with PTFE tubing (see list of Materials) and a tube connector (see list of Materials) to a Hamilton syringe equipped with a micro-drive (see list of Materials). The needle holder is mounted on a 3-way micromanipulator (see list of Materials).
	<ol>
		<li>Using a needle holder attached to a syringe, rinse the needle for cell transplantation with 70% ethanol. Dry by sucking up air into the needle. Do not dry by blowing, as this may result in dust getting stuck in the tapered part of the needle.</li>
		<li>Insert the needle in the needle holder connected to the Hamilton syringe, with the bevel upwards.</li>
	</ol>
	</li>
	<li>Shield-to-shield Cell Transplantation
	<ol>
		<li>Enter the shield of a donor embryo with the transplantation needle and draw up about 10-20 cells labeled with ABP140-mCherry. Carefully avoid drawing yolk.</li>
		<li>Transplant the cells into the shield of the host embryo. Repeat until all host embryos have been transplanted.</li>
		<li>Remove any damaged embryos with a fire-polished Pasteur pipette. Place the transplanted embryos in the incubator at 28 &#176;C and let them recover for about 30 min.</li>
		<li>Using a needle holder attached to a syringe, rinse the cell transplantation needle with water and place it back in a Petri dish on a band of modeling clay. Seal the dish with parafilm.</li>
	</ol>
	</li>
</ol>
6. (OPTIONAL) Single Cell Transplantation
Note: In the embryo, cells adhere to each other, so that it is difficult to draw only one cell in the transplantation needle. We developed a modified protocol to easily transplant single prechordal plate progenitor cells. The idea is to dissociate cells prior to transplantation. Because isolated prechordal plate progenitor cells tend to lose their identity, we genetically impose them a prospective prechordal plate identity, by activating the Nodal signaling pathway, in absence of the Sox32 transcription factor. We have verified that these induced cells behave like endogenous prechordal plate progenitor cells8. Below are the specific steps to perform single cell transplantations. Cell dissociation is achieved by placing embryos in Calcium-free Ringer11, dissecting an explant and mechanically stirring it.
<ol>
	<li>At step 2.1, prepare the transplantation dish with 1% agarose in Calcium-free Ringer instead of Embryo Medium.</li>
	<li>At step 3.1, in addition to ABP140-mCherry RNA and sin1 morpholino, add 3 ng of RNAs encoding the active form of the Nodal receptor Taram-A (0.6 ng/&#181;l final) and 1.25 &#181;l of morpholino against sox32 (stock solution at 2 mM, hence 0.5 mM final).</li>
	<li>After step 4, transfer three selected donor embryos in the dish for cell transplantation filled with Pen/Strep Calcium-free Ringer using a fire-polished Pasteur pipette. With fine tweezers, dissect an explant containing the injected cells (ABP140-mCherry labeled cells). Rapidly discard the rest of the embryos.</li>
	<li>With an eyelash, gently stir the explant until cells dissociate.</li>
	<li>Draw up a single isolated cell in the transplantation needle and transplant it into the shield of a host embryo.</li>
	<li>Using a fire-polished Pasteur pipette, transfer the embryos in an agarose-coated dish filled with Pen/Strep EM. Place the embryos in the incubator at 28 &#176;C for 30 min.</li>
</ol>
7. Embryo Mounting
<ol>
	<li>Imaging Chamber
	<ol>
		<li>Method 1: use an imaging chamber composed of a plastic slide (75*25*0.5 mm) pierced with 4 holes (5.5 mm diameter), with 3 mm high edges on one side (Figure 2C-D). Glue a glass coverslip on the back side, with cyanoacrylate glue (super glue). 
		Note: At the end of the experiment, place the chamber in water for one night to remove the coverslip and get rid of glue residues with sand paper.</li>
		<li>Method 2: use 35 mm MatTek glass bottom dishes (see list of Materials) with a 10 mm hole.</li>
	</ol>
	</li>
	<li>Embryo Mounting
	<ol>
		<li>Fill up a glass vial with 1 ml of hot 0.2 % agarose in embryo medium. Keep it at 42 &#176;C in a heat-block.</li>
		<li>Under the fluorescent stereomicroscope, select an embryo in which red transplanted cells are part of the prospective prechordal plate labeled in green (i.e. red transplanted cells are within the green prospective prechordal plate, and have started migrating towards the animal pole).</li>
		<li>Transfer a selected embryo into the agarose solution with a fire-polished Pasteur pipette. Discard the excess of embryo medium from the pipette. Draw the embryo in the pipette with agarose, and transfer the embryo in the imaging chamber, depositing a drop of agarose. Before agarose is set, orient the embryo with an eyelash. Place the prospective prechordal plate upwards for imaging with an upright microscope, or on the glass bottom for imaging with an inverted microscope. Wait until the agarose is set (around 1 min, depending on room temperature) before mounting another embryo in the next well of the chamber. 
		Note: an imaging chamber containing 4 wells allows mounting up to 4 embryos.</li>
		<li>When the agarose is set, fill the chamber with Pen/Strep EM to avoid drying.</li>
	</ol>
	</li>
</ol>
8. Live Imaging
<ol>
	<li>Use a 40X water-immersion long-range lens. Use appropriate filter sets to image GFP and mCherry (for GFP: excitation BP 470/40, beam splitter FT 510, and emission BP 540/50; for mCherry: excitation BP 578/21, beam splitter FT 596, and emission LP 641/75). Adjust exposure duration in order to optimize image dynamics.</li>
	<li>To image all cells in the plate, acquire z-stacks, starting a few &#181;m above the prospective prechordal plate (in the ectoderm) and ending a few &#181;m below the prospective prechordal plate (in the yolk). Use a z-step of 2 &#181;m. To optimize acquisition speed, switch colors between z-stacks rather than between frames. 
	Note: This may lead to slight differences between the green and red images, but is not a problem since no precise co-localization between the green and red signals is required. To reduce further imaging time and exposure to light, green may be acquired only once every 5 time points, which is sufficient to identify prechordal plate progenitor cells.</li>
	<li>Using such settings, image up to 4 embryos within the two-minute time interval which is necessary to analyze protrusion frequency and orientation. For analysis of protrusion lifetime, reduce time interval to 30 sec to get higher time resolution. Image from 60% epiboly to 80% epiboly.</li>
</ol>
9. Cell Dynamics Analysis
<ol>
	<li>Load 4D images in ImageJ 
	Note: Depending on the software used for acquisition, images are stored in different formats (one file per image, one file per z-stack, one file for the whole experiment). Some ImageJ Plugins are available online to open 4D stacks. Our data are stored as one file per z-stack. Because the dataset can be large, it may be convenient to open only part of it (some time points, or a subpart of the z-stack, or 8-bit converted images&#8230;). We use a custom-made ImageJ plugin to do so, which we would be happy to distribute on request.</li>
	<li>Analysis of the Orientation and Frequency of Cell Protrusions
	<ol>
		<li>Use GFP images to determine the main direction of prospective prechordal plate migration. Take this as a reference for cell protrusions angle measurements. Rotate the stack, so that this reference direction is parallel to one side of the image.</li>
		<li>Follow one cell. Select angle tool. For each frame and each protrusion, use the angle tool to measure the angle between the protrusion and the reference direction (Figure 3A). Use the Analyze/Measure command to store the value (or press M on keyboard). Save the results as a txt file. Repeat with the next cell.</li>
		<li>Import the data into R and plot angle distribution as histograms. Use the Kolmogorov-Smirnov test (ks.test in R) to compare different conditions. 
		Note: The angle tool provides values comprised between 0&#176; and 180&#176;, allowing the use of classical statistical tests and not circular tests.</li>
		<li>With this data set, calculate emission frequency as the number of protrusions per cell per minute. Use Student T-test (t.test in R) to compare different conditions.</li>
	</ol>
	</li>
	<li>Analysis of the Cell Protrusions Lifetime
	<ol>
		<li>For each cell and each protrusion, measure protrusion duration (number of frames during which the protrusion is present x time-interval between frames).</li>
		<li>Import the data into R. Use Student T-test (t.test in R) to compare different conditions. 
		Note: Other migration features can be interesting to analyze in order to characterize the cell migration. These include cell tracks, for speed, direction and persistence measurements, or cell morphology. Some commercial software applications are available to measure these features, but a large number of open-source software applications and ImageJ plugins are also available, as for instance ADAPT to analyze morphodynamics12, DiPer to look at cell trajectories and directional persistence13, CellTrack to track cell boundaries during the migration14.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

This protocol presents an easy way to study the role of a candidate gene in cell migration in vivo, by combining the creation of chimeric embryos using cell transplantation with live imaging.
Creation of mosaic embryos
Studying the dynamics of a cell requires the visualization of its contour to analyze cytoplasmic extensions. This can be achieved by labeling isolated cells in an otherwise unlabeled&#160;&#8211; or differently labeled - environment, thus off...

In multicellular organisms, cell migration is essential both for the development of the embryo where it ensures the organization of cells into tissues and organs, and for adult life, where it takes part to tissue homeostasis (wound healing) and immunity. In addition to these physiological functions, cell migration is also involved in diverse pathological situations, including, in particular, cancer metastasis.
Cell migration has been analyzed in vitro for decades, providing an overall understanding of the molecular mechanisms ensuring cell movements on flat surfaces. In vivo however, cells are confronted by a more complex environment. It clearly appeared in the past years that migration within an organism may be influenced by external cues such as the extracellular matrix, neighboring cells or secreted chemokines guiding migration, and that the mechanisms driving cell migration may vary from what has been described in vitro1,2. The mechanisms ensuring in vivo cell migration have received less attention so far, mainly because of the increased technical difficulty, compared to in vitro studies. In vivo analysis of cell migration in particular requires direct optical access to migrating cells, techniques to label unique cells in order to see their dynamics and morphology, as well as gain or loss of function approaches to test the role of candidate genes. So far, only a few model systems harboring these characteristics have been used to dissect in vivo cell migration3.
We recently used the migration of the prospective prechordal plate in early zebrafish embryos as a new convenient model system to assess the function of candidate genes in controlling in vivo cell migration4,5. Prospective prechordal plate (also known as anterior mesendoderm) is a group of cells forming at the onset of gastrulation on the dorsal side of the embryo. During gastrulation this group collectively migrates towards the animal pole of the embryo6-8, to form the prechordal plate, a mesendodermal thickening, anterior to the notochord, and underlying the neural plate. The anterior part of the prechordal plate will give rise to the hatching gland, while its posterior part likely contributes to head mesoderm9. Thanks to the external development and optical clarity of the fish embryo, cell migration can be directly and easily observed in this structure.
Cell transplantation is a very potent technique that allows for the rapid and easy creation of mosaic embryos10. Expressing fluorescent cytoskeletal markers in transplanted cells results in the labeling of isolated cells, the morphology and dynamics of which can be easily observed. Combining this to loss or gain of function approaches permits the analysis of cell-autonomous functions of a candidate gene.
The presented protocol describes how we assessed the function of the TORC2 component Sin1 in controlling cell migration and actin dynamics in vivo5. But, as mentioned in the results and further discussed, it could be used to analyze the potential implication of any candidate gene in controlling cell migration in vivo.

<table><tbody><tr><td>Glass capillaries (outside diameter 1.0 mm, inside diameter 0.58&amp;nbsp;mm)</td><td>Harvard Apparatus</td><td>300085</td><td>standard thickness</td></tr><tr><td>Glass capillaries (outside diameter 1.0 mm, inside diameter 0.78 mm)</td><td>Harvard Apparatus</td><td>300085</td><td>thin-walled</td></tr><tr><td>Penicillin-Streptomycin</td><td>Sigma-Aldrich</td><td>P4333</td><td>10&amp;nbsp;000&amp;nbsp;units penicillin and 10&amp;nbsp;mg streptomycin per ml</td></tr><tr><td>fine tweezers</td><td>Dumont Fine Science Tools</td><td>11254-20</td><td>5F</td></tr><tr><td>glass bottom dishes</td><td>MatTek</td><td>P35G-0-10-C</td><td></td></tr><tr><td>Air transjector</td><td>Eppendorf</td><td>5246</td><td></td></tr><tr><td>Micro-forge</td><td>Narishige</td><td>MF-900</td><td></td></tr><tr><td>Microgrinder</td><td>Narishige</td><td>EG-44</td><td></td></tr><tr><td>Micromanipulator (for injection)</td><td>Narishige</td><td>MN-151</td><td></td></tr><tr><td>Micromanipulator (for cell transplantation)</td><td>Leica</td><td>Leica Micromanipulator</td><td></td></tr><tr><td>Hammilton Syringe</td><td>Narishige</td><td>IM-9B</td><td></td></tr><tr><td>Micropipette puller</td><td>David Kopf Instruments</td><td>Model 720</td><td></td></tr><tr><td>Transplantation mold</td><td>Adapative Science Tools</td><td>PT-1</td><td></td></tr><tr><td>Needle holder</td><td>Narishige</td><td>HI-7</td><td></td></tr><tr><td>Tube connector</td><td>Narishige</td><td>CI-1</td><td></td></tr><tr><td>PTFE tubing</td><td>Narishige</td><td>CT-1</td><td></td></tr></tbody></table>

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.

The presented technique was used to analyze the role of Sin1, one of the core components of the Tor complex 2 (TORC2), in controlling in vivo cell migration. The use of cell transplantation permits labeling of isolated cells and analysis of cell-autonomous effects. Movie S1 shows the migration of transplanted prechordal plate progenitor cells. Actin labeling with ABP140 allows the easy visualization of actin-rich cytoplasmic protrusions. We measured their frequency and orientatio...

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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.

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 ...

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Research

JoVE Journal

Developmental Biology

7.6K Views.  IBENS, Institut de Biologie de l'École Normale Supérieure. 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. 

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

Invasive Behavior of Human Breast Cancer Cells in Embryonic Zebrafish

In many cases, cancer patients do not die of a primary tumor, but rather because of metastasis. Although numerous rodent models are available for studying cancer metastasis in vivo, other efficient, reliable, low-cost models are needed to quickly access the potential effects of (epi)genetic changes or pharmacological compounds. As such, we illustrate and explain the feasibility of xenograft models using human breast cancer cells injected into zebrafish embryos to support this goal. Under the microscope, fluorescent proteins or chemically labeled human breast cancer cells are transplanted into transgenic zebrafish embryos, Tg (fli:EGFP), at the perivitelline space or duct of Cuvier (Doc) 48 h after fertilization. Shortly afterwards, the temporal-spatial process of cancer cell invasion, dissemination, and metastasis in the living fish body is visualized under a fluorescent microscope. The models using different injection sites, i.e., perivitelline space or Doc are complementary to one another, reflecting the early stage (intravasation step) and late stage (extravasation step) of the multistep metastatic cascade of events. Moreover, peritumoral and intratumoral angiogenesis can be observed with the injection into the perivitelline space. The entire experimental period is no more than 8 days. These two models combine cell labeling, micro-transplantation, and fluorescence imaging techniques, enabling the rapid evaluation of cancer metastasis in response to genetic and pharmacological manipulations.

Here, we describe xenograft zebrafish models using two different injection sites, i.e., perivitelline space and duct of Cuvier, to investigate the invasive behavior and to assess the intravasation and extravasation potential of human breast cancer cells, respectively.

In many cases, cancer patients do not die of a primary tumor, but rather because of metastasis. Although numerous rodent models are available for studying ...

Cancer Research

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 ...

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

The genetic and technical strengths have made the zebrafish vertebrate a key model organism in which the consequences of gene manipulations can be traced in vivo throughout the rapid developmental period. Multiple processes can be studied including cell proliferation, gene expression, cell migration and morphogenesis. Importantly, the generation of chimeras through transplantations can be easily performed, allowing mosaic labeling and tracking of individual cells under the influence of the host environment. For example, by combining functional gene manipulations of the host embryo (e.g., through morpholino microinjection) and live imaging, the effects of extrinsic, cell nonautonomous signals (provided by the genetically modified environment) on individual transplanted donor cells can be assessed. Here we demonstrate how this approach is used to compare the onset of fluorescent transgene expression as a proxy for the timing of cell fate determination in different genetic host environments.
In this article, we provide the protocol for microinjecting zebrafish embryos to mark donor cells and to cause gene knockdown in host embryos, a description of the transplantation technique used to generate chimeric embryos, and the protocol for preparing and running in vivo time-lapse confocal imaging of multiple embryos. In particular, performing multiposition imaging is crucial when comparing timing of events such as the onset of gene expression. This requires data collection from multiple control and experimental embryos processed simultaneously. Such an approach can easily be extended for studies of extrinsic influences in any organ or tissue of choice accessible to live imaging, provided that transplantations can be targeted easily according to established embryonic fate maps.

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

Live tracking of individual WT retinal progenitors in distinct genetic backgrounds allows for the assessment of the contribution of cell non-autonomous signaling during neurogenesis. Here, a combination of gene knockdown, chimera generation via embryo transplantation and in vivo time-lapse confocal imaging was utilized for this purpose.

The genetic and technical strengths have made the zebrafish vertebrate a key model organism in which the consequences of gene manipulations can be traced ...

Multi-Photon Time Lapse Imaging to Visualize Development in Real-time: Visualization of Migrating Neural Crest Cells in Zebrafish Embryos

Congenital eye and craniofacial anomalies reflect disruptions in the neural crest, a transient population of migratory stem cells that give rise to numerous cell types throughout the body. Understanding the biology of the neural crest has been limited, reflecting a lack of genetically tractable models that can be studied in vivo and in real-time. Zebrafish is a particularly important developmental model for studying migratory cell populations, such as the neural crest. To examine neural crest migration into the developing eye, a combination of the advanced optical techniques of laser scanning microscopy with long wavelength multi-photon fluorescence excitation was implemented to capture high-resolution, three-dimensional, real-time videos of the developing eye in transgenic zebrafish embryos, namely Tg(sox10:EGFP) and Tg(foxd3:GFP), as sox10 and foxd3 have been shown in numerous animal models to regulate early neural crest differentiation and likely represent markers for neural crest cells. Multi-photon time-lapse imaging was used to discern the behavior and migratory patterns of two neural crest cell populations contributing to early eye development. This protocol provides information for generating time-lapse videos during zebrafish neural crest migration, as an example, and can be further applied to visualize the early development of many structures in the zebrafish and other model organisms.

A combination of the advanced optical techniques of laser scanning microscopy with long wavelength multi-photon fluorescence excitation was implemented to capture high-resolution, three-dimensional, real-time imaging of neural crest migration in Tg(sox10:EGFP) and Tg(foxd3:GFP) zebrafish embryos.

Congenital eye and craniofacial anomalies reflect disruptions in the neural crest, a transient population of migratory stem cells that give rise to ...

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 ...

Biology

Utilizing Larval Zebrafish for Efficient In Vivo Tumor Studies

A Simple, Rapid, and Effective Method for Tumor Xenotransplantation Analysis in Transparent Zebrafish Embryos

In vivo studies of tumor behavior are a staple of cancer research; however, the use of mice presents significant challenges in cost and time. Here, we present larval zebrafish as a transplant model that has numerous advantages over murine models, including ease of handling, low expense, and short experimental duration. Moreover, the absence of an adaptive immune system during larval stages obviates the need to generate and use immunodeficient strains. While established protocols for xenotransplantation in zebrafish embryos exist, we present here an improved method involving embryo staging for faster transfer, survival analysis, and the use of flow cytometry to assess disease burden. Embryos are staged to facilitate rapid cell injection into the yolk of the larvae and cell marking to monitor the consistency of the injected cell bolus. After injection, embryo survival analysis is assessed up to 7 days post injection (dpi). Finally, disease burden is also assessed by marking transferred cells with a fluorescent protein and analysis by flow cytometry. Flow cytometry is enabled by a standardized method of preparing cell suspensions from zebrafish embryos, which could also be used in establishing the primary culture of zebrafish cells. In summary, the procedure described here allows a more rapid assessment of the behavior of tumor cells in vivo with larger numbers of animals per study arm and in a more cost-effective manner.

Author Spotlight: Advancing Cancer Therapeutics Using Tumor Xenotransplantation in Zebrafish

We describe a protocol for xenotransplantation into the yolk of transparent zebrafish embryos that is optimized by a simple, rapid staging method. Post-injection analyses include survival and assessing the disease burden of xenotransplanted cells by flow cytometry.

In vivo studies of tumor behavior are a staple of cancer research; however, the use of mice presents significant challenges in cost and time. Here, we ...

Live Imaging of Cell Extrusion from the Epidermis of Developing Zebrafish

Homeostatic maintenance of epithelial tissues requires the continual removal of damaged cells without disrupting barrier function. Our studies have found that dying cells send signals to their live neighbors to form and contract a ring of actin and myosin that ejects it out from the epithelial sheet while closing any gaps that might have resulted from its exit, a process termed cell extrusion1. The optical clarity of developing zebrafish provides an excellent system to visualize extrusion in living epithelia. Here we describe a method to induce and image extrusion in the larval zebrafish epidermis. To visualize extrusion, we inject a red fluorescent protein labeled probe for F-actin into one-cell stage transgenic zebrafish embryos expressing green fluorescent protein in the epidermis and induce apoptosis by addition of G418 to larvae. We then use time-lapse imaging on a spinning disc confocal microscope to observe actin dynamics and epithelial cell behaviors during the process of apoptotic cell extrusion. This approach allows us to investigate the extrusion process in live epithelia and will provide an avenue to study disease states caused by the failure to eliminate apoptotic cells.

Dying cells are extruded from epithelial tissues by concerted contraction of neighboring cells without disrupting barrier function. The optical clarity of developing zebrafish provides an excellent system to visualize extrusion in living epithelia. Here we describe methods to induce and image extrusion in the larval zebrafish epidermis at cellular resolution.

Homeostatic maintenance of epithelial tissues requires the continual removal of damaged cells without disrupting barrier function.  Our studies have found ...

Robust Zebrafish Cell Transplantation for Regeneration and Development Research

High-resolution Cell Transplantation in Embryonic and Larval Zebrafish

Development and regeneration occur by a process of genetically encoded spatiotemporally dynamic cellular interactions. The use of cell transplantation between animals to track cell fate and to induce mismatches in the genetic, spatial, or temporal properties of donor and host cells is a powerful means of examining the nature of these interactions. Organisms such as chick and amphibians have made crucial contributions to our understanding of development and regeneration, respectively, in large part because of their amenability to transplantation. The power of these models, however, has been limited by low genetic tractability. Likewise, the major genetic model organisms have lower amenability to transplantation. 
The zebrafish is a major genetic model for development and regeneration, and while cell transplantation is common in zebrafish, it is generally limited to the transfer of undifferentiated cells at the early blastula and gastrula stages of development. In this article, we present a simple and robust method that extends the zebrafish transplantation window to any embryonic or larval stage between at least 1 and 7 days post fertilization. The precision of this approach allows for the transplantation of as little as one cell with near-perfect spatial and temporal resolution in both donor and host animals. While we highlight here the transplantation of embryonic and larval neurons for the study of nerve development and regeneration, respectively, this approach is applicable to a wide range of progenitor and differentiated cell types and research questions.

Author Spotlight: Mapping Cellular Connectivity in the Zebrafish Nervous System During Development and Regeneration

Here we present a protocol to transplant cells with high spatial and temporal resolution in zebrafish embryos and larvae at any stage between at least 1 and 7 days post fertilization.

Development and regeneration occur by a process of genetically encoded spatiotemporally dynamic cellular interactions. The use of cell transplantation ...

Analyzing Craniofacial Morphogenesis in Zebrafish Using 4D Confocal Microscopy

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical clarity and amenability to genetic manipulation, the zebrafish embryo has become a popular model organism with which to perform time-lapse analysis of morphogenesis in living embryos. Confocal imaging of a live zebrafish embryo requires that a tissue of interest is persistently labeled with a fluorescent marker, such as a transgene or injected dye. The process demands that the embryo is anesthetized and held in place in such a way that healthy development proceeds normally. Parameters for imaging must be set to account for three-dimensional growth and to balance the demands of resolving individual cells while getting quick snapshots of development. Our results demonstrate the ability to perform long-term in vivo imaging of fluorescence-labeled zebrafish embryos and to detect varied tissue behaviors in the cranial neural crest that cause craniofacial abnormalities. Developmental delays caused by anesthesia and mounting are minimal, and embryos are unharmed by the process. Time-lapse imaged embryos can be returned to liquid medium and subsequently imaged or fixed at later points in development. With an increasing abundance of transgenic zebrafish lines and well-characterized fate mapping and transplantation techniques, imaging any desired tissue is possible. As such, time-lapse in vivo imaging combines powerfully with zebrafish genetic methods, including analyses of mutant and microinjected embryos.

Time-lapse confocal imaging is a powerful technique useful for characterizing embryonic development. Here, we describe the methodology and characterize craniofacial morphogenesis in wild-type, as well as pdgfra, smad5, and smo mutant embryos.

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical ...

Elucidating Microglia Behavior in Zebrafish Optic Tectum Development

Live Imaging and Characterization of Microglia Dynamics in the Zebrafish Embryo

Microglia are highly dynamic cells and their migration and colonization of the brain parenchyma is a crucial step for proper brain development and function. Externally developing zebrafish embryos possess optical transparency, which along with well-characterized transgenic reporter lines that fluorescently label microglia, make zebrafish an ideal vertebrate model for such studies. In this paper, we take advantage of the unique features of the zebrafish model to visualize the dynamics of microglia cells in vivo and under physiological conditions. We use confocal microscopy to record a timelapse of microglia cells in the optic tectum of the zebrafish embryo and then, extract tracking data using the IMARIS 10.0 software to obtain the cells' migration path, mean speed, and distribution in the optic tectum at different developmental stages. This protocol can be a useful tool to elucidate the physiological significance of microglia behavior in various contexts, contributing to a deeper characterization of these highly motile cells.

Author Spotlight: Using Zebrafish to Explore Microglia Migration During Brain Development

We demonstrate a method that takes advantage of fast scanning confocal microscopy to perform live imaging of microglia cells in the developing zebrafish optic tectum, allowing for the analysis of the dynamics of these cells in vivo.

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

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