We thank Kristen Kwan for the pCS2-H2A.F/Z-EGFP and pCS2-mCherry-CAAX vectors. We thank Chris Rodesch for tutoring us in live imaging in zebrafish. We thank Shawn Williams, Erik Malarkey and Brad Yoder for assistance in confocal imaging at UAB and the High Resolution Imaging Facility at UAB. The High Resolution Imaging Facility is supported by the UAB Comprehensive Cancer Center Support Grant (P30CA013148) and the Rheumatic Disease Core Center (P30 AR048311). J.M.P. is supported by the National Institute of Neurological Disease and Stroke (NIH R21 NS092105), and pilot grants from American Cancer Society (ACS IRG-60-001-53-IRG) and the UAB Comprehensive Cancer Center (P30CA013148). S.M.P. is supported by the Cell and Molecular Biology T32 Training Grant (5T32GM008111-28).

1. In Vitro Transcription
<ol>
	<li>Linearize pCS2-H2A.F/Z-EGFP and/or pCS2-mCherry-CAAX vectors by NotI restriction enzyme digest31. Using an RNA in vitro transcription kit, generate 5&#39; capped mRNA products from each template, according to manufacturer&#39;s protocol.</li>
	<li>Purify the capped mRNA using a purification kit. Follow manufacturer&#39;s instructions. Elute with RNase-free H2O.</li>
	<li>Determine the concentration of RNA by absorbance at 260 nm using a spectrophotometer. (OD260 x dilution x 40 &#181;g/ml).</li>
	<li>Dilute the RNA to 100 ng/&#181;l for each H2A.F/Z-EGFP and mCherry-CAAX with RNase-free H2O. If the RNA concentration is too low, the fluorescence will be diminished or absent. Brighter samples will diminish the concerns over phototoxicity and photobleaching. On the other hand, too much RNA can be toxic and/or cause off target effects. 
	Note: Store the remaining purified mRNA in -80 &#176;C freezer.</li>
</ol>
2. Zebrafish Breeding, Embryo Collection, and mRNA Injection 36-38
<ol>
	<li>Assemble breeding tanks with a barrier to separate the tank into two regions and fill each breeding tank with aquaculture system water used in the zebrafish facility.</li>
	<li>In order to prevent untimely breeding, place two male fish on one side of the barrier and two female fish on the other side the night before breeding.</li>
	<li>The next day, thaw the previously prepared mRNA mixture on ice. Replace the water in breeding tanks with fresh aquaculture system water and remove the barriers. Immediately after the barriers are pulled, warm an injection mold to 28.5 &#176;C and set up the equipment for microinjection. 
	Note: For information on injection molds, please refer to Gerlach36.</li>
	<li>Collect eggs every 10 - 15 min using a tea strainer and rinse the eggs into a clean 100 x 15 mm Petri dish with E3 Blue (5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2, 0.33 mM MgSO4, 10-5% Methylene blue). E3 Blue is used to prevent fungal growth and ensure proper development of larval fish. For more information on mating and embryo collection, refer to Gerlach36 and Porazinkski37.</li>
	<li>Embed one-cell staged embryos in a warmed injection mold and inject the RNA into the yolk in the desired amount of embryos (Figure 1A). Account for natural embryonic death and unfertilized embryos by performing embryonic injections on 15% more embryos than needed for the experiment. For additional details on microinjection of zebrafish embryos, please refer to Gerlach36 and Porazinkski37. 
	NOTE: For first time use of mRNA, perform a dose-curve analysis to determine the optimal dose for fluorescence and viability (defined as no gross developmental defects up to 5 dpf) prior to performing 5D imaging. 150 - 200 ng/&#181;l injected into embryos is often the optimal concentration, therefore it is a good starting point for the final concentration.</li>
	<li>Carefully extract the injected embryos from the mold using a modified 9&#34; glass Pasteur pipette. To modify the pipette, melt the end using a Bunsen burner until it forms a ball.</li>
	<li>Place injected embryos in a 100 x 15 mm Petri dish in E3 Blue and house in a 28.5 &#176;C incubator.</li>
	<li>Six hr post-injection, remove any dead or unfertilized embryos from the plates and add clean E3 Blue. House the embryos at 28.5 &#176;C.</li>
</ol>
3. Preparation and Embedding of Live Zebrafish Embryos for Imaging (Figure 1B)
<ol>
	<li>Two hr before imaging, screen the injected embryos for GFP using a fluorescent dissecting microscope. Place bright green GFP-expressing embryos in a new 100 x 15 mm Petri dish with E3 Blue.</li>
	<li>Boil a stock solution of 1% low melt agarose by adding 1 g of low melt agarose to 100 ml of E3 Blue. After using the agarose, cover the flask with aluminum foil. The stock solution remains useful for up to one month.</li>
	<li>Aliquot 3 ml of the melted agar into a 17 x 100 mm culture tube. Keep the agarose warm by placing the culture tube in a 42 &#176;C water bath until ready for use. Prepare a 15 mM Tricaine solution in deionized water to anesthetize the zebrafish embryos36. 
	NOTE: If imaging at earlier time points is desired, the concentration of agar can be decreased as low as 0.3%39.</li>
	<li>Bring the 15 mM Tricaine, screened embryos, low melt agarose, E3 Blue, and a 35 mm glass coverslip bottom culture dish to a dissection light microscope. Carefully remove the embryo&#39;s chorion with #5 tweezers. Do this for three embryos.</li>
	<li>Place the dechorionated embryos in a separate container to be anesthetized. The lid of the coverslip bottom dish is often used for this purpose. Using a transfer pipette, add three drops (approximately 150 &#181;l) of 15 mM Tricaine to the dish of 5 ml E3 blue (if using the lid of the coverslip bottom dish) or until embryos have been sufficiently anesthetized. In addition, add 3-4 drops (approximately 150 - 200 &#181;l) of 15 mM Tricaine solution to the 1% melted agarose tube.</li>
	<li>Using a p200 pipette with 1 cm of the pipette tip cut off; transfer the anesthetized embryos to the coverslip-bottomed dish. Remove any excess E3 Blue:Tricaine solution.</li>
	<li>Slowly add 5 - 10 &#181;l of low-melt agarose:Tricaine solution over the embryos, keeping each drop separate to ensure the embryos do not accidentally drift close to one another. 
	Warning: If the agarose is too warm, it will damage the embryo. A good temperature to maintain the agar at is 42 &#176;C.</li>
	<li>Using a 21 G 1 &#189; needle, gently orient the embryo in the agarose to the desired position. When using an inverted microscope for time-lapse imaging, orient the region of interest (ROI) as close to the coverslip as possible. 
	Note: For general purposes, the tail region offers ease of orientation and clarity due to the relatively thin tissue (Figure 1A). Other tissues, such as the epithelial layer surrounding the yolk and fin folds, can be used28,29. These tissues offer great clarity, however these regions are only a few cell layers thick. For the purpose of this protocol, it is beneficial to image the tail region to acquire as many cell divisions as possible.</li>
	<li>Allow a few min for partial solidification of the agar. Use the needle to break apart a small piece of agar to test its solidification. When a piece of agar can be pulled away from the drop, proceed to the next step.</li>
	<li>Cover the entire coverslip with low melt agar forming a dome over the embedded embryos. Allow the agar to solidify before moving the dish for confocal imaging (Figure 1A).</li>
	<li>During the agar solidification process (takes approximately 10 min), prepare 3 ml of E3 Blue solution with five drops of (approximately 250 &#181;l) 15 mM Tricaine to be placed over the embedded embryos during imaging.</li>
</ol>
4. 5D Confocal Imaging of Live Zebrafish Embryos40,41
NOTE: See Ariga40 and O&#39;Brien41 for details on how to perform 5D imaging using other confocal systems. For Z-interval, Z-stack, Z-depth, time interval, and 5D definitions see Figure 1C.
<ol>
	<li>Open the imaging software and set the microscope to 60X NA 1.4 objective lens. Apply immersion oil to the objective lens and place the culture dish in the slide holder on the microscope stage. Using the axis controller, center the embryo of interest above the objective lens and bring the objective lens upward to meet the culture dish.</li>
	<li>Click on the eye piece icon and switch to the GFP filter on the microscope. Focus on the ROI. Focusing on the tissue closest to the coverslip will offer the best imaging results.</li>
	<li>Remove the interlock. Select the GFP and mCherry channels (pre-set wavelengths in software) and set the line averaging option to normal.</li>
	<li>Use &#34;View/Acquisition Controls/A1 Scan Area&#34; command to open the A1 Scan Area tool.</li>
	<li>Begin scanning. Using the axis-controller, position the embryo so that the scan area is filled with as much of the zebrafish as possible. The laser power does not need to be optimal at this point. Lower the laser power to avoid unnecessary photobleaching.</li>
	<li>Use the &#34;View/Acquisition Controls/ND acquisition&#34; command to open the ND acquisition control panel.</li>
	<li>Begin scanning to set the Z-stack parameters. Set the Z-stack upper limit to where the cells are not in focus and lower limit to where cells are no longer visible. Allow for a 3 &#181;m space above the sample for growth and cell movement, which may expand into the imaging field. The Z-stack for the figures shown in this protocol covered a Z-depth of 40 &#181;m in the embryo tail.</li>
	<li>Set the Z-interval step size to 2 &#181;m. On average, a cell is 10 &#181;m in diameter; therefore 2 &#181;m will produce five intervals of imaging data to be analyzed for each cell. 
	NOTE: The depth of the image that can be acquired depends on the Z interval. Z resolution is sacrificed to gain overall depth in the Z dimension in order to image as many cells undergoing mitosis as possible (large Z-depth). The inverse is true, in that, by decreasing the step size, Z resolution is gained, while Z depth is sacrificed (small Z-depth).</li>
	<li>Adjust the image laser power, HV, and offset levels. For the experiments demonstrated in this protocol, use the following laser power, HV, and offset levels set at the corresponding levels, respectively, for the GFP channel; 2 - 5, 120 - 140, and -9 to -11. For the mCherry channel, use the following laser power, HV, and offset levels, respectively; 3 - 6, 120 - 140, and -3 to -8. Once the parameters are set, shut off the scan to prevent unnecessary laser exposure that may cause phototoxicity and photobleaching of the sample.</li>
	<li>Select the 2x line averaging icon. &#34;No averaging&#34; produces a grainy image while &#34;4x line averaging&#34; drastically increases the scan time. Use of 2x line averaging provides the best image quality and fastest scan time.</li>
	<li>Select the appropriate time interval and time duration necessary for the experiment. For wild-type divisions, two-minute time intervals for 2 hr is best for determining mitotic duration (used in Figures 1, 2, 3A, and 3B). Divisions that activate the spindle assembly checkpoint for longer than 30 min are more suitable for five minute intervals for four hr in order to preserve fluorescence as demonstrated in Figure 3C.</li>
	<li>Check the &#34;save to file&#34; box and name the file to automatically save the file as it is being acquired. Double check all parameters are set correctly and hit &#34;start run&#34;.</li>
	<li>After acquisition is complete, to view the file in a three dimensional format, click on the volume threshold icon.</li>
</ol>

The authors have nothing to disclose.

Use of this method allows one to infer nuclear envelope breakdown, formation of a metaphase plate by microtubule-kinetochore attachments, and segregation of sister chromatids to form two new cells in vivo and in a time-dependent manner. The ability to observe mitosis in zebrafish is advantageous over fixed samples and cell culture systems because the cells are being imaged in the natural physiology, the tissue is transparent which allows for fluorescent proteins to be used, they develop relatively fast, and time...

Mitosis is a critical cellular process essential for growth, differentiation, and regeneration in a living organism. Upon accurate preparation and replication of DNA in interphase, the cell is primed to divide. The first phase of mitosis, prophase, is initiated by activation of cyclin B/Cdk1. Prophase is characterized by condensation of chromatin material into chromosomes. Nuclear envelope breakdown occurs at the transition between prophase and prometaphase. In prometaphase, centrosomes, the nucleating center for spindle formation, begin to migrate to opposite poles while extending microtubules in search of kinetochore attachment. Upon attachment, conversions to end-on microtubule attachment and tension forces orient the chromosomes forming a metaphase plate1. If all chromosomes are attached correctly, the spindle assembly checkpoint is satisfied, cohesin rings holding the sister chromatids together are cleaved, and microtubules shorten to pull sister chromatids to opposite poles during anaphase2,3. The final phase, telophase, involves elongation of the cell and reformation of the nuclear envelope around the two new nuclei. Cytokinesis completes the division process by separating the cytoplasm of the two new daughter cells4-6. Alteration of key mitotic pathways (i.e., spindle assembly checkpoint, centrosome duplication, sister chromatid cohesion, etc.) can result in metaphase arrest, missegregation of chromosomes, and genomic instability7-10. Ultimately, defects in pathways controlling mitosis can cause developmental disorders and cancer, necessitating visualization of mitosis and its defects in a live, vertebrate, multi-cellular organism10-16.
Zebrafish embryos serve as a great model organism for live imaging due to the transparent tissue, ease of microinjection, and fast development. Using zebrafish, the overall goal of this manuscript is to describe a method of live 5D (dimensions X, Y, Z, time, and wavelength) imaging of mitosis17 (Figure 1C). The use of mutant zebrafish defective in different mitotic pathways demonstrate the consequence of such defects. For this protocol, Aurora B and Esco2 mutants were chosen to illustrate these defects. Aurora B is a kinase that is part of the chromosome passenger complex (CPC) involved in spindle formation and microtubule attachment. It is also required for cleavage furrow formation in cytokinesis18,19. In zebrafish, Aurora B deficiency leads to defects in furrow induction, cytokinesis, and chromosome segregation20. Esco2, on the other hand, is an acetyltransferase that is essential for sister chromatid cohesion21,22. It acetylates cohesin on the SMC3 portion of the ring thus stabilizing cohesin to ensure proper chromosome segregation at the metaphase-anaphase transition23. Loss of Esco2 in zebrafish leads to chromosome missegregation, premature sister chromatid separation, genomic instability, and p53-dependent and independent apoptosis24,25. Due to the availability, auroraBhi1045, and esco2hi2865 mutant zebrafish (hereafter referred to as aurBm/m and esco2m/m, respectively) will be used to illustrate this technique25-27.
Coupling confocal microscopy with fluorescent-tagged cell machinery has enabled researchers to visualize chromatin and cell membrane dynamics during mitosis25,28,29. Fluorescent-tagged histones have historically been used to visualize chromatin. Histones are nuclear proteins composed of four different pairs (H2A, H2B, H3, and H4) that are responsible for the nucleosome structure that composes chromosomes30. While H2B is arguably the most used histone for fluorescent proteins in mouse and cell culture, use of Histone 2A, Family Z (H2A.F/Z) has proved well for use in zebrafish31,32. Concanavalin A and casein kinase 1-gamma for example, localize to the cell membrane and have previously been shown effective in visualizing the cell membrane in sea urchins and drosophila33,34. Other studies have shown that the CAAX fluorescent-tagged protein labels the cell membrane and was successful in zebrafish31. CAAX is a motif that is recognized by post-translational modifying enzymes such as farnesyltransferases and geranylgeranyltransferases. Modifications by these enzymes cause proteins to become membrane-associated, thus labeling the cell membrane35.
Due to the prior use in zebrafish, this protocol chose to use H2A.F/Z and CAAX to label chromatin and the cell membrane. Application of this method will allow the researcher to monitor mitosis at the individual cell level to observe individual chromosome dynamics, as well as simultaneously monitor multiple cell divisions that may impact tissue differentiation and development. This article will focus on imaging the dynamics of chromosome segregation during mitosis at the individual cell level. Within this manuscript, the ability to observe several mitotic divisions, calculate division time, and decipher the mitotic phenotypes will be illustrated and discussed. By using these parameters, physiologically relevant data can be collected and applied to several disease states affected by mitotic defects.

<table><tbody><tr><td>pCS2 vectors</td><td>Gift from K. Kwan</td><td></td><td>For plasmid of interest</td></tr><tr><td>NotI-HF restriction enzyme</td><td>New England BioLabs</td><td>R3189S</td><td>For restriction digest of plasmid</td></tr><tr><td>mMessage SP6 kit</td><td>Life Technologies</td><td>AM1340</td><td>For &lt;em&gt;in vitro&lt;/em&gt; transcription</td></tr><tr><td>RNeasy Mini kit</td><td>Qiagen</td><td>74104</td><td>For purifying mRNA</td></tr><tr><td>100 x 15 mm Petri dishes</td><td>Fisher Scientific</td><td>FB0875712</td><td>For housing embryos</td></tr><tr><td>microinjection mold</td><td>homemade</td><td></td><td>For holding embryos during microinjection</td></tr><tr><td>Agarose II</td><td>Amresco</td><td>0815-25G</td><td>For embedding embryos</td></tr><tr><td>Tricaine</td><td>Sigma-Aldrich</td><td>E10521-10G</td><td>For anesthetizing embryos</td></tr><tr><td>Sodium Chloride</td><td>Sigma-Aldrich</td><td>S9888</td><td>For embryo water (E3 Blue), dissolved in UltraPure H&lt;sub&gt;2&lt;/sub&gt;O</td></tr><tr><td>Potassium Chloride</td><td>Sigma-Aldrich</td><td>P3911</td><td>For embryo water (E3 Blue), dissolved in UltraPure H&lt;sub&gt;2&lt;/sub&gt;O</td></tr><tr><td>Calcium Chloride Dihydrate</td><td>Sigma-Aldrich</td><td>C8106</td><td>For embryo water (E3 Blue), dissolved in UltraPure H&lt;sub&gt;2&lt;/sub&gt;O</td></tr><tr><td>Magnesium Sulfate</td><td>Fisher Scientific</td><td>M7506</td><td>For embryo water (E3 Blue), dissolved in UltraPure H&lt;sub&gt;2&lt;/sub&gt;O</td></tr><tr><td>Methylene Blue Hydrate</td><td>Sigma-Aldrich</td><td>MB1</td><td>For embryo water (E3 Blue), dissolved in UltraPure H&lt;sub&gt;2&lt;/sub&gt;O</td></tr><tr><td>100 mm culture tube</td><td>Fisher Scientific</td><td>50-819-812</td><td>For melted agar</td></tr><tr><td>35 mm glass coverslip bottom culture dish&amp;nbsp;</td><td>MatTek Corp</td><td>P35G-0-20-C&amp;nbsp;</td><td>No. 0, 20 mm glass, For embedding embryos</td></tr><tr><td>#5 tweezers</td><td>Dumont</td><td>72701-D</td><td>For dechorionating embryos</td></tr><tr><td>21 G 1 1/2 gauge needle</td><td>Becton Dickinson</td><td>305167</td><td>For positioning embryos in agar</td></tr><tr><td>Dissecting microscope</td><td>Nikon AZ100</td><td></td><td>For screening and embedding embryos, any dissecting scope will do</td></tr><tr><td>Confocal microscope</td><td>Nikon A1+</td><td></td><td>For time-lapse imaging</td></tr><tr><td>Confocal software</td><td>NIS Elements AR 4.13.00</td><td></td><td>For image acquisition and processing</td></tr></tbody></table>

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.

Figure 2 demonstrates the ability to observe many cell divisions using a wide field view of an AB wild-type zebrafish tail. Over seven mitotic cells are imaged in a 14 min time frame (Movie 1). Within the two hr time-course, over 40 mitotic events were captured. On average, 50 dividing cells were observed in the AB and 30 dividing cells in aurBm/m embryos (Figure 2B). To account for the number of cells imaged,...

Watch this Scientific Journal Video about Observing Mitotic Division and Dynamics in a Live Zebrafish Embryo at JoVE.com

Observing Mitotic Division and Dynamics in a Live Zebrafish Embryo

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 ...

observing-mitotic-division-and-dynamics-in-a-live-zebrafish-embryo

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12.0K Views.  University of Alabama at Birmingham. 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.

Video: Observing Mitotic Division and Dynamics in a Live Zebrafish Embryo

Capturing Tissue Repair in Zebrafish Larvae with Time-lapse Brightfield Stereomicroscopy

The zebrafish larval tail fin is ideal for studying tissue regeneration due to the simple architecture of the larval fin-fold, which comprises of two layers of skin that enclose undifferentiated mesenchyme, and because the larval tail fin regenerates rapidly within 2-3 days. Using this system, we demonstrate a method for capturing the repair dynamics of the amputated tail fin with time-lapse video brightfield stereomicroscopy. We demonstrate that fin amputation triggers a contraction of the amputation wound and extrusion of cells around the wound margin, leading to their subsequent clearance. Fin regeneration proceeds from proximal to distal direction after a short delay. In addition, developmental growth of the larva can be observed during all stages. The presented method provides an opportunity for observing and analyzing whole tissue-scale behaviors such as fin development and growth in a simple microscope setting, which is easily adaptable to any stereomicroscope with time-lapse capabilities.

We present a protocol for capturing the dynamics of zebrafish larval tail fin regeneration on a whole-tissue scale using brightfield-based stereomicroscopy. This technique enables capturing the regeneration dynamics with single cell resolution. This methodology can be adapted to any stereomicroscope equipped with a CCD camera and time-lapse software.

The zebrafish larval tail fin is ideal for studying tissue regeneration due to the simple architecture of the larval fin-fold, which comprises of two ...

Profiling DNA Replication Timing Using Zebrafish as an In Vivo Model System

DNA replication timing is an important cellular characteristic, exhibiting significant relationships with chromatin structure, transcription, and DNA mutation rates. Changes in replication timing occur during development and in cancer, but the role replication timing plays in development and disease is not known. Zebrafish were recently established as an in vivo model system to study replication timing. Here is detailed the protocols for using the zebrafish to determine DNA replication timing. After sorting cells from embryos and adult zebrafish, high-resolution genome-wide DNA replication timing patterns can be constructed by determining changes in DNA copy number through analysis of next generation sequencing data. The zebrafish model system allows for evaluation of the replication timing changes that occur in vivo throughout development, and can also be used to assess changes in individual cell types, disease models, or mutant lines. These methods will enable studies investigating the mechanisms and determinants of replication timing establishment and maintenance during development, the role replication timing plays in mutations and tumorigenesis, and the effects of perturbing replication timing on development and disease.

Profiling DNA Replication Timing Using Zebrafish as an In Vivo Model System

Zebrafish were recently used as an in vivo model system to study DNA replication timing during development. Here is detailed the protocols for using zebrafish embryos to profile replication timing. This protocol can be easily adapted to study replication timing in mutants, individual cell types, disease models, and other species.

DNA replication timing is an important cellular characteristic, exhibiting significant relationships with chromatin structure, transcription, and DNA ...

Genetics

Ploidy Manipulation of Zebrafish Embryos with Heat Shock 2 Treatment

Manipulation of ploidy allows for useful transformations, such as diploids to tetraploids, or haploids to diploids. In the zebrafish Danio rerio, specifically the generation of homozygous gynogenetic diploids is useful in genetic analysis because it allows the direct production of homozygotes from a single heterozygous mother. This article describes a modified protocol for ploidy duplication based on a heat pulse during the first cell cycle, Heat Shock 2 (HS2). Through inhibition of centriole duplication, this method results in a precise cell division stall during the second cell cycle. The precise one-cycle division stall, coupled to unaffected DNA duplication, results in whole genome duplication. Protocols associated with this method include egg and sperm collection, UV treatment of sperm, in vitro fertilization and heat pulse to cause a one-cell cycle division delay and ploidy duplication. A modified version of this protocol could be applied to induce ploidy changes in other animal species.

A modified protocol for ploidy manipulation uses a heat shock to induce a one-cycle stall in cytokinesis in the early embryo. This protocol is demonstrated in the zebrafish but may be applicable to other species.

Manipulation of ploidy allows for useful transformations, such as diploids to tetraploids, or haploids to diploids. In the zebrafish Danio rerio, ...

A Versatile Mounting Method for Long Term Imaging of Zebrafish Development

Zebrafish embryos offer an ideal experimental system to study complex morphogenetic processes due to their ease of accessibility and optical transparency. In particular, posterior body elongation is an essential process in embryonic development by which multiple tissue deformations act together to direct the formation of a large part of the body axis. In order to observe this process by long-term time-lapse imaging it is necessary to utilize a mounting technique that allows sufficient support to maintain samples in the correct orientation during transfer to the microscope and acquisition. In addition, the mounting must also provide sufficient freedom of movement for the outgrowth of the posterior body region without affecting its normal development. Finally, there must be a certain degree in versatility of the mounting method to allow imaging on diverse imaging set-ups. Here, we present a mounting technique for imaging the development of posterior body elongation in the zebrafish D. rerio. This technique involves mounting embryos such that the head and yolk sac regions are almost entirely included in agarose, while leaving out the posterior body region to elongate and develop normally. We will show how this can be adapted for upright, inverted and vertical light-sheet microscopy set-ups. While this protocol focuses on mounting embryos for imaging for the posterior body, it could easily be adapted for the live imaging of multiple aspects of zebrafish development.

Here, we present a versatile mounting method that allows for the long-term time-lapse imaging of the posterior body development of live zebrafish embryos without perturbing normal development.

Zebrafish embryos offer an ideal experimental system to study complex morphogenetic processes due to their ease of accessibility and optical transparency. ...

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 ...

Biology

Visualization of Cellular Electrical Activity in Zebrafish Early Embryos and Tumors

Bioelectricity, endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane, plays important roles in signaling processes of excitable neuronal and muscular cells and many other biological processes, such as embryonic developmental patterning. However, there is a need for in vivo electrical activity monitoring in vertebrate embryogenesis. The advances of genetically encoded fluorescent voltage indicators (GEVIs) have made it possible to provide a solution for this challenge. Here, we describe how to create a transgenic voltage indicator zebrafish using the established voltage indicator, ASAP1 (Accelerated Sensor of Action Potentials 1), as an example. The Tol2 kit and a ubiquitous zebrafish promoter, ubi, were chosen in this study. We also explain&#160;the processes of Gateway site-specific cloning, Tol2 transposon-based zebrafish transgenesis, and the imaging process for early-stage fish embryos and fish tumors using regular epifluorescent microscopes. Using this fish line, we found that there are cellular electric voltage changes during zebrafish embryogenesis, and fish larval movement. Furthermore, it was observed that in a few zebrafish malignant peripheral nerve sheath tumors, the tumor cells were generally polarized compared to the surrounding normal tissues.

Here, we show the process of creating a cellular electric voltage reporter zebrafish line to visualize embryonic development, movement, and fish tumor cells in vivo.

Bioelectricity, endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane, plays important roles in signaling ...

Dissection and Lateral Mounting of Zebrafish Embryos: Analysis of Spinal Cord Development

The zebrafish spinal cord is an effective investigative model for nervous system research for several reasons. First, genetic, transgenic and gene knockdown approaches can be utilized to examine the molecular mechanisms underlying nervous system development. Second, large clutches of developmentally synchronized embryos provide large experimental sample sizes. Third, the optical clarity of the zebrafish embryo permits researchers to visualize progenitor, glial, and neuronal populations. Although zebrafish embryos are transparent, specimen thickness can impede effective microscopic visualization. One reason for this is the tandem development of the spinal cord and overlying somite tissue. Another reason is the large yolk ball, which is still present during periods of early neurogenesis. In this article, we demonstrate microdissection and removal of the yolk in fixed embryos, which allows microscopic visualization while preserving surrounding somite tissue. We also demonstrate semipermanent mounting of zebrafish embryos. This permits observation of neurodevelopment in the dorso-ventral and anterior-posterior axes, as it preserves the three-dimensionality of the tissue.

Developmental processes such as proliferation, patterning, differentiation, and axon guidance can be readily modeled in the zebrafish spinal cord. In this article, we describe a mounting procedure for zebrafish embryos, which optimizes visualization of these events.

The zebrafish spinal cord is an effective investigative model for nervous system research for several reasons. First, genetic, transgenic and gene ...

Neuroscience

Refined Multiview Light Sheet Microscopy Protocol for Zebrafish Embryos

Light Sheet Microscopy Imaging and Mounting Strategies for Early Zebrafish Embryos

Light sheet microscopy has become the methodology of choice for live imaging of zebrafish embryos over long time scales with minimal phototoxicity. In particular, a multiview system, which allows sample rotation, enables imaging of entire embryos from different angles. However, in most imaging sessions with a multiview system, sample mounting is a troublesome process as samples are usually prepared in a polymer tube. To aid in this process, this protocol describes basic mounting strategies for imaging early zebrafish development between the 70% epiboly and early somite stages. Specifically, the study provides statistics on the various positions the embryos default to at the 70% epiboly and bud stages within the chorion. Furthermore, it discusses the optimum number of angles and the interval between angles required for imaging whole zebrafish embryos at the early stages of development so that cellular-scale information can be extracted by fusing the different views. Finally, since the embryo covers the entire field of view of the camera, which is required to obtain a cellular-scale resolution, this protocol details the process of using bead information from above or below the embryo for the registration of the different views.

Author Spotlight: Strategies for Mounting Zebrafish Embryos for High-Resolution Multiview Light-Sheet Microscopy &#8212; Techniques for Imaging and Image Reconstruction

A sample preparation strategy for imaging early zebrafish embryos within an intact chorion using a light-sheet microscope is described. It analyzes the different orientations that embryos acquire within the chorion at the 70% epiboly and bud stages and details imaging strategies for obtaining cellular-scale resolution throughout the embryo using the light-sheet system.

Light sheet microscopy has become the methodology of choice for live imaging of zebrafish embryos over long time scales with minimal phototoxicity. In ...

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.

Microglia are highly dynamic cells and their migration and colonization of the brain parenchyma is a crucial step for proper brain development and ...

Immunology and Infection

Zebrafish Reproduction and Development

The zebrafish (Danio rerio) has become a popular model for studying genetics and developmental biology. The transparency of these animals at early developmental stages permits the direct visualization of tissue morphogenesis at the cellular level. Furthermore, zebrafish are amenable to genetic manipulation, allowing researchers to determine the effect of gene expression on the development of a vertebrate with a high degree of genetic similarity to humans.
This video provides a brief overview of the major phases of zebrafish development, with particular focus on the first 24 hours post fertilization (hpf). The discussion begins with a zygote consisting of a single cell, or blastomere, atop a large ball of yolk. Cleavage of the blastomere is then shown to produce an embryo containing thousands of cells within a matter of hours. Next, the dramatic cellular movements known as epiboly and gastrulation are explained, revealing how they contribute to reshaping a mass of cells into a moving embryo with a beating heart in just 1 day. The presentation follows embryo development through the hatching phase, when they become swimming, feeding larvae. Important considerations for caring for larvae are incorporated, including a brief review of how fish are raised to adulthood in a dedicated facility known as the nursery. Finally, the video concludes with some common techniques utilized for studying embryo development, demonstrating how zebrafish are used to help us better understand human development and disease.

Observing Mitotic Division and Dynamics in a Live Zebrafish Embryo

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Dissection and Lateral Mounting of Zebrafish Embryos: Analysis of Spinal Cord Development

Light Sheet Microscopy Imaging and Mounting Strategies for Early Zebrafish Embryos

Live Imaging and Characterization of Microglia Dynamics in the Zebrafish Embryo

Zebrafish Reproduction and Development