This work was supported by NIH grant 5K01OD021419-02 and NC State University College of Veterinary Medicine.

1. Embryo Preparation
<ol>
	<li>Fix 48 h post-fertilization (hpf) chimeric zebrafish embryos generated by blastula-to-blastula transplant between AB wild-type zebrafish embryos in 4% paraformaldehyde (PFA) overnight with rocking at 4 &#176;C. Perform two 5 min washes with rocking at room temperature using 500 &#956;L of 1x phosphate buffered saline containing 0.1% of a non-ionic surfactant (1x PBSt; see the Table of Materials) 
	NOTE: Paraformaldehyde is toxic and a carcinogen and must be properly disposed of per institutional regulations. Work with paraformaldehyde should be performed in a chemical hood, and appropriate personal protective equipment (gloves, lab coat, and safety glasses) should always be used.</li>
	<li>Dehydrate embryos by washing sequentially with 500 &#956;L of 30%, 50%, and 70% methanol (MeOH) diluted with 1x PBSt for 10 min each at room temperature with rocking. Incubate embryos in 500 &#956;L of 100% MeOH at -20 &#176;C for at least 14 h. 
	NOTE: Methanol is toxic and must be properly disposed of per institutional regulations. Work with methanol should be performed in a chemical hood, and appropriate personal protective equipment (gloves, lab coat, and safety glasses) should always be used.</li>
	<li>Rehydrate embryos by washing sequentially with 500 &#956;L of 70%, 50%, and 30% MeOH diluted with 1x PBSt for 10 min each at room temperature with rocking. Perform two 5 min washes with 500 &#956;L of 1x PBSt at room temperature with rocking.</li>
	<li>Remove the 1x PBSt and incubate in 500 &#956;L of 30% sucrose diluted with deionized water at room temperature with rocking until the embryos sink to the bottom of the tube (approximately 1 h). Replace sucrose with 500 &#956;L of 15% fish gelatin/25% sucrose (15/25; see the Table of Materials) and incubate at room temperature with rocking overnight.</li>
	<li>Replace approximately half of the volume of 15/25 with optimal cutting temperature medium (OCT medium) and incubate at room temperature with rocking until the embryos sink to the bottom of the tube (approximately 1 h).</li>
	<li>Replace approximately half of the volume with OCT medium and incubate at room temperature with rocking for 1 h. Repeat this step one time.</li>
	<li>During incubation steps with 15/25 and OCT medium, invert or flick the tube as needed to combine these reagents.</li>
</ol>
2. Embryo Embedding and Preparation of Cryosections
<ol>
	<li>Transfer embryos to a plastic mold (see the Table of Materials) with forceps, minimizing any transfer of the 15/25-OCT mixture. Fill the mold approximately half full with OCT medium and gently mix the embryos in OCT medium.</li>
	<li>Prepare labeled plastic molds and transfer desired embryos (usually 1&#8210;3 embryos) into the empty, labeled plastic molds, minimizing carry-over of OCT medium. Once the desired embryos are in the plastic mold, gently fill with OCT medium to the top of the mold.</li>
	<li>Use forceps or 25 G needles to arrange embryos to the desired orientation, using a light stereomicroscope for visualization (Figure 1A,B).</li>
	<li>Freeze the prepared molds on dry ice in an insulated container with a metal platform. Place an ice bucket or foam cooler gently over the metal platform to create a cold chamber (Figure 1C,D)</li>
	<li>Prepare 10&#8210;12 &#956;m thick cryosections using a cryostat set at -20 &#176;C.
	<ol>
		<li>Set up one block at a time and use OCT medium to freeze the block onto the disc/chuck with the bottom of the block facing toward the blade (Figure 2A). Ensure that the block is completely frozen to the disc (the OCT medium will turn white) before beginning sectioning (Figure 2B).</li>
		<li>Place cryosections on charged glass slides (Figure 2C,D) and air-dry the slides overnight at room temperature.</li>
	</ol>
	</li>
</ol>
3. Immunofluorescence for pH3
<ol>
	<li>Perform three 5 min washes of the slides in 1x PBS in an appropriate container, such as a Coplin jar. If tissues are not well adhered to the slide, perform washes by laying slides on a flat surface and gently pipetting 500 &#956;L of 1x PBS onto the surface. Pour off 1x PBS between washes by gently tipping the slides.</li>
	<li>Outline the sections with a barrier pen or wax pencil to keep liquid on the slides (Figure 3).</li>
	<li>Lay the slides on a flat surface in a humid chamber and pipette 200 &#956;L of block buffer per section onto the slides. Incubate in block buffer for 2 h at room temperature or overnight at 4 &#176;C.</li>
	<li>Prepare the primary antibody dilution (rabbit anti-pH3 antibody, 1:200; see Table of Materials) in block buffer and mix well by pipetting. Gently tip the slides to drain off the block buffer, return to the humid chamber, and pipette 200 &#956;L of primary antibody solution per section onto the slides. For the secondary-only control, pipette 200 &#956;L of block buffer onto the appropriate sections.</li>
	<li>Incubate the slides at 4 &#176;C overnight in a humid chamber filled with deionized water. Seal the edges of the humid chamber to help retain moisture.</li>
	<li>Perform three 5 min washes of the slides in 1x PBS in an appropriate container, such as a Coplin jar. During wash steps, prepare the secondary antibody dilution (anti-rabbit fluorescent secondary antibody conjugate, 1:2,000; see Table of Materials) in block buffer and mix well by pipetting.</li>
	<li>Lay the slides on a flat surface in a humid chamber and pipette 200 &#956;L of secondary antibody solution per section onto the slides. Incubate the slides in secondary antibody solution(s) at room temperature, shielded from light, for 30 min.</li>
	<li>Perform three 5 min washes of the slides in 1x PBS in an appropriate container, such as a Coplin jar, shielded from light. During wash steps, prepare a nuclear staining solution (see the Table of Materials).</li>
	<li>Place the slides on a flat surface and pipette 200&#8210;500 &#956;L (depending on the size of sample) of the nuclear staining solution onto each section. Incubate the slides in the nuclear staining solution for 10 min, shielded from light.</li>
	<li>Drain off the nuclear staining solution and mount the slides with non-hardening fluorescent mounting media and a glass coverslip. Maintain the slides in the dark at 4 &#176;C until imaging is performed. 
	NOTE: The best results are obtained when the slides are imaged on the same day or the next day.</li>
	<li>Following IF, visualize and image the slides with a confocal fluorescence microscope and digital camera using 555 nm (red) and 645 nm (far-red) emission filters at 100x magnification (10x ocular magnification and 10x objective magnification). Keep the laser power consistent throughout imaging.</li>
	<li>After imaging, place the slides in individual containers of 1x PBS and store flat at 4 &#176;C overnight to prepare for coverslip removal. When placing the slides in 1x PBS, gently agitate the slides to assist in removal of the coverslip. 
	NOTE: Do not press down on the coverslip or attempt to remove the coverslip manually, as this can compromise quality of the sections. The coverslips must be removed horizontally with minimal forced movement.</li>
</ol>
4. Immunohistochemistry for Labeled Dextran
<ol>
	<li>After the coverslips have been removed, gently transfer the slides into new 1x PBS in an appropriate container, such as a Coplin jar. Remove the 1x PBS and perform three 5 min incubations of the slides in 1x Tris-buffered saline with 0.1% of a non-ionic surfactant (1x TBSt) at room temperature.</li>
	<li>Prepare 250 mL of 3% hydrogen peroxide (H2O2) diluted in deionized water in an appropriately sized container. Incubate the slides in 3% H2O2 solution at room temperature for 15 min. 
	NOTE: Hydrogen peroxide is corrosive and must be properly disposed of per institutional regulations. Appropriate personal protective equipment (gloves, lab coat, and safety glasses) should always be used.</li>
	<li>Quickly rinse the slides with deionized water. Perform three 5 min washes of the slides in 1x TBSt. Carefully dry the area around the tissue sections and place the slides flat in a humid chamber. Outline the tissue sections with a barrier pen or wax pencil to keep liquid on the sections if needed.</li>
	<li>Apply a ready-to-use 2.5% serum blocking solution provided with a secondary antibody labeling kit (see the Table of Materials) dropwise to each section. Incubate the slides in a humid chamber at room temperature for 20 min.</li>
	<li>Prepare the primary antibody diluted in antibody diluent (rabbit anti-labeled dextran, 1:7,500; see the Table of Materials) and mix by gentle pipetting. Gently tip the slides to drain off the block buffer, dry the area around the sections, and place the slides flat in a humid chamber.</li>
	<li>Do not wash before applying the primary antibody solution. Pipette 200 &#956;L of primary antibody solution per section onto the slides and incubate in a humid chamber at 4 &#176;C overnight. For the secondary-only control, pipette 200 &#956;L of antibody diluent onto the appropriate sections.</li>
	<li>Gently tip the slides to drain off the primary antibody solution and place in 1x TBSt in an appropriate container, such as a Coplin jar. Perform two 5 min washes of the slides in 1x TBSt. Dry the area around the sections and place flat in a humid chamber.</li>
	<li>Apply a ready-to-use background-reducing blocking reagent (see the Table of Materials) dropwise to each section. Incubate the slides in a humid chamber at room temperature for 20 min.</li>
	<li>Gently tip the slides to drain off the block buffer, carefully dry the area around the sections, and place the slides flat in a humid chamber. Do not wash before applying the secondary antibody solution.</li>
	<li>Apply a ready-to-use secondary antibody solution (anti-rabbit horseradish peroxidase (HRP) polymer secondary antibody; see the Table of Materials) to each section dropwise and incubate in a humid chamber at room temperature for 30 min.</li>
	<li>Gently tip the slides to drain off the secondary antibody solution and place in 1x TBSt in an appropriate container, such as a Coplin jar. Perform two 5 min washes of the slides in 1x TBSt.</li>
	<li>Prepare the HRP chromogenic substrate immediately before use according to the manufacturer&#8217;s protocol (see the Table of Materials).</li>
	<li>Dry the area around the sections, place the slides on a flat surface, and apply 200 &#956;L of the HRP chromogenic substrate to each section. Incubate at room temperature for 3 min, starting a timer when the substrate has been applied to the first slide (Figure 4).</li>
	<li>Drain off the substrate solution and rinse the slides briefly in a Coplin jar containing 1x PBS. Place the slides in deionized water in an appropriate container, such as a Coplin jar, and perform two 5 min washes in deionized water with gentle agitation.</li>
	<li>Counterstain the slides by placing them in hematoxylin staining solution for up to 30 s. 
	NOTE: Hematoxylin is toxic and must be disposed of per institutional regulations. Appropriate personal protective equipment (gloves, lab coat, and safety glasses) should always be used.</li>
	<li>Place the slides in deionized water in an appropriate container, such as a Coplin jar, and perform three 5 min washes in deionized water. Transfer the slides to a container containing Scott&#8217;s tap water (see the Table of Materials) and incubate for 1 min.</li>
	<li>Place the slides in deionized water in an appropriate container, such as a Coplin jar, and perform three 5 min washes in deionized water.</li>
	<li>Dehydrate the slides through a series of ethanol (EtOH) grades (diluted in deionized water) and xylene in a chemical hood. Perform one 2 min incubation in 70% EtOH, two 1 min incubations in 95% EtOH, two 1 min incubations in 100% EtOH, and three 1 min incubations in xylene. Remove the slides from xylene and place a coverslip while wet with a toluene-based mounting medium in a chemical hood. 
	NOTE: Xylene and toluene are toxic and flammable and must be properly disposed of per institutional regulations. Work with xylene and toluene should be performed in a chemical hood, and appropriate personal protective equipment (gloves, lab coat, and safety glasses) should always be used.</li>
	<li>Following IHC, visualize and image the slides with a compound light microscope and digital camera at 100x magnification (10x ocular magnification and 10x objective magnification). (Figure 5).</li>
</ol>

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The authors have nothing to disclose.

We have presented a novel method for combined immunofluorescence and immunohistochemistry that represents an important step forward in performing colocalization experiments on cryosectioned zebrafish embryos. There is a critical lack of existing colocalization protocols that are optimized for use with small embryos and cryosectioned material17,18, both of which are otherwise commonly used in molecular studies14. Existing colocalization protocols mainly focus on simultaneous visualization of two fluorophores17,18. While these protocols can work well, their usefulness is limited by the availability of antibodies that (i) can be used in zebrafish and (ii) are compatible with IF.
We feel that use of cryosectioning for zebrafish embryos provides a significant advantage in terms of preserved tissue morphology. As IHC is more commonly performed on paraffin sections than cryosections7,8, further exploration of cryosection-based IHC methods for detection of protein expression is warranted. The use of whole mount IF is a common method for visualizing expression levels in zebrafish embryos, and is more commonly described in the literature than IF using cryosections1,2,3,4,19. However, whole mount protocols have limitations, including the lack of accurate localization of proteins expressed in deep tissues and the potential for surface level expression to obscure protein expression in deeper tissues. We have consistently observed excellent maintenance of cellular morphology following cryopreservation, sectioning, and sequential IF and IHC. For applications that require precise localization of protein expression at the single-cell level, there is a significant advantage to section-based procedures as we have described in this protocol. Resin embedding provides an alternative option for performing IF or IHC assays on sections from early-stage zebrafish embryos20. Resin sections provide superior preservation of tissue morphology compared to cryosections, and relatively thinner tissue sections can be prepared. However, manufacturers generally do not recommend the use of resin sections for immuno-based assays, as some components of the resins cannot be removed from the sections and may mask antibody binding sites21.
While the whole of the protocol is essential for accurate and successful analysis of expression patterns, a few specific steps are critical for experimental success. The first critical step is the handling of embryos during embedding in OCT13,14. It can be challenging to orient each embryo in the same transverse plane so that sections are cut from approximately the same area for all embryos. We have found that using small gauge needles to perform small, deliberate movements through the viscous OCT medium for embryo orientation is superior to using forceps, which are too large and displace too much OCT medium. A second critical step occurs during sectioning of frozen blocks, as it can be difficult to obtain high-quality sections that contain the desired tissue(s) without significant training and experience. We thus recommend use of test samples until the technique is mastered. A third critical step is coverslip removal, which has the potential to disturb or strip off tissue samples. We have found that a combination of gentle agitation to loosen the coverslip and slow removal of the slide from the PBS container is the most effective approach for removing coverslips. A gentle and steady hand is best; researchers may find that some trial and error is necessary to learn how to remove coverslips effectively.
Additional important steps in the protocol include antibody optimization (primary and secondary antibodies for both IF and IHC assays) and exposure of slides to the chromogenic substrate and counterstain. Thorough research regarding primary antibody specificity and target concentration is essential; generally, it is best to collect sufficient non-precious samples for at least two separate experiments for IF and IHC that will be performed to optimize primary antibody concentrations before beginning the IF/IHC combination. Including a known positive and negative control for each primary antibody is essential in each experiment to ensure that both IF and IHC assays are performing appropriately. Adjustments to secondary antibody concentration may also be necessary. Optimization of exposure of tissue sections to the chromogenic substrate and counterstain for the IHC assay is required, since manufacturer guidelines often describe a wide range of possible exposure times. Not all chromogens may be compatible with counterstains and endogenous tissue pigmentation, requiring careful planning with regard to chromogen selection.
There are numerous potential modifications that can be applied to the described protocol, and we have successfully performed this protocol with other combinations of antibodies that could not both be used for IF. As alluded to above, chromogenic substances have distinct compatibility with specific counterstains. We have found that these components are easily modifiable in the protocol. Additionally, the parameters for antibody incubation are quite flexible, and can be increased or decreased depending on desired staining intensity. The embedding process can also be modified. While we have focused on transverse sections, embryos can easily be oriented in any direction to address particular tissues of interest. Finally, there are multiple possible pause points in this protocol. Dehydrated embryos can be stored in 100% MeOH at -20 &#176;C for a few months; frozen blocks can be stored at -80 &#176;C for up to three months; and prepared slides can be stored at -80 &#176;C for up to nine months in a slide box.
Due to the relative complexity of this combined IF/IHC protocol, there are multiple possible sources of variation or error that may require troubleshooting, ranging from tissue quality to researcher experience to sample handling. Most of the critical steps described above are considered critical because they either require optimization at the individual user level or they require particular dexterity, skill, and experience. However, we have found that nonspecific background staining and low signal intensity are the most significant issues requiring optimization. As with any IF or IHC protocol, addressing these issues can be time consuming and labor intensive, and can be compounded with this method since the two techniques are combined. For this reason, we highly recommend optimizing IF and IHC separately before proceeding with the combined protocol.
While we predict that this method will be useful for a broad range of experiments, there are potential limitations. Successful performance of this procedure is dependent on maintaining intact tissue samples through two sequential experiments that include numerous wash steps. For this reason, any problems that arise during sectioning or tissue handling, including use of older slides that may have degraded in quality, will significantly limit researchers&#8217; ability to perform this protocol successfully. Although slides can potentially be stored at -80 &#176;C for up to nine months, tissue adherence to slides declines over time and we had the best success with slides that are used within a month of preparation or ideally, the next day. A second limitation is the small footprint of zebrafish embryos. While we have found this experiment to be useful in visualizing proteins that are relatively abundantly expressed in early-stage embryos, proteins that are expressed inconsistently or at low levels may be very difficult to capture in a section. Finally, since the protocol uses 10&#8210;12 &#181;m cryosections, it is best suited for evaluation of protein expression in larger structures, such as nuclei, rather than smaller sub-cellular components.
In summary, our combined IF/IHC protocol will be advantageous for a wide variety of studies in early-stage zebrafish embryos, and represents an important innovation in precise protein expression analysis in such specimens. Our method, shown successfully in Figure 5, will allow researchers to preserve the delicate morphology of early-stage zebrafish embryos in cryosections while working with the somewhat limited range of currently available primary antibodies that are validated for use in IF or IHC in this species. This protocol will likely be useful for studies in other fish and amphibian species (e.g., Medaka and Xenopus spp.), which are hampered by similar limitations to antibody availability, and may be applicable for traditional mammalian models as well.

The zebrafish is an extremely robust model organism that is currently used across a wide variety of disciplines in biomedical research. In particular, the rapid external development and translucency of zebrafish embryos provide an excellent tool for in vivo studies. Herein, we describe a method for sequential immunofluorescence (IF) and immunohistochemical (IHC) analyses of cryosectioned zebrafish embryos. This novel procedure uses sequential application of two antibodies on a single slide, enabling accurate identification of colocalized proteins at the cellular level while conserving tissue sections. This protocol is particularly useful for studies with the zebrafish model, since a relatively small number of antibodies have been validated for use in IF and/or IHC applications in zebrafish compared to mouse models.
The observation of intercellular interactions is an essential element in many studies, and may provide key insights into molecular mechanisms functioning at the cellular level that underlie phenotypes at the organismal level. Additionally, protein expression can provide information about cellular function, especially when examining expression of multiple proteins simultaneously within the cell (co-localization). Although whole-mount IHC and IF are commonly used techniques for analyzing protein expression in zebrafish embryos1,2,3,4,5, whole mount procedures can be problematic for achieving precise colocalization data. In our experience, it can be difficult to differentiate between layers of tissue and visualize protein expression at the single-cell level in whole-mount specimens. Imaging software programs may generally be unable to distinguish between surface staining versus deeper staining. Non-surface level protein expressions can be obscured by more brightly expressed surface level expression, leading to inaccuracies in quantification. Additionally, most traditional zebrafish clearing methods are quite toxic6 and thus less desirable for use.
Antibody-based techniques, such as IF and IHC, are often used to detect protein expression in sectioned material, simplifying the identification of discrete cell populations that express a particular protein within complex tissues. IHC is commonly used for colocalization, most frequently by using two different antibodies conjugated in different host species and visualized with different colored chromogens4,7,8,9,10. However, using multiple chromogens can lead to nonspecific background staining or incompatibility of colors11,12.
We developed a novel protocol for detection of multiple proteins by sequential IF and IHC on cryosectioned early-stage zebrafish embryos. Cryosectioning is particularly well-suited to delicate tissues such as zebrafish embryos, and cryosections are superior to paraffin-embedded sections for fluorescence-based assays13,14. We chose to optimize combined IF and IHC rather than dual-color IF or IHC, to circumvent the problem of antibody incompatibility for a single assay type. These problems are particularly relevant for research involving zebrafish due to the limited number of commercially available antibodies that are validated for use in zebrafish. In fact, a study of four large companies showed that commercially available antibodies for use in mouse was about 112,000 versus about 5,300 for use in zebrafish15. Finally, we chose to develop a protocol that could be performed on a single cryosection, which is essential when working with small or limited tissue samples such as are obtained from zebrafish embryos.
This protocol was designed to assess the proliferative behavior of donor cells in 48 h post-fertilization chimeric zebrafish embryos that were generated by blastula-to-blastula transplant as described by Carmany-Rampey and Moens16. Donor embryos were injected at the one-cell stage with a fluorescently labeled dextran conjugate prior to transplant of donor cells into recipient embryos. We used immunofluorescence for Ser 10 phosphorylated Histone H3 (pH3) to detect proliferating cells followed by immunohistochemistry for the labeled dextran to detect donor cells in chimeric zebrafish embryos. Sequential detection of pH3 and the labeled dextran within a single cryosection enabled us to identify and quantify individual cells that expressed both markers.
This sequential IF/IHC protocol for cryosectioned zebrafish will provide a useful tool for zebrafish researchers who desire a colocalization protocol for protein expression. The problems that this protocol was designed to address, such as small tissue specimens and limited antibody availability, are not unique to the zebrafish model. This method may therefore be of use to any researcher desiring to perform sequential IF/IHC.
ETHICS STATEMENT:
All animal studies were approved by the Institutional Animal Care and Use Committee, North Carolina State University, Raleigh, North Carolina, USA.

<table><tbody><tr><td>15/25</td><td>N/A</td><td>N/A</td><td>15% fish gelatin/25% sucrose w/v, made in dH&lt;sub&gt;2&lt;/sub&gt;O</td></tr><tr><td>Alexa 555 secondary antibody</td><td>Invitrogen</td><td>A21429</td><td>used at 1:2,000 dilution in block buffer for IF</td></tr><tr><td>Antibody Diluent</td><td>Dako</td><td>S3022</td><td></td></tr><tr><td>Background Buster</td><td>Innovex</td><td>NB306</td><td></td></tr><tr><td>block buffer (IF)</td><td>N/A</td><td>N/A</td><td>5% normal goat serum, 0.1% Triton X-100 in 1x PBS</td></tr><tr><td>cellSens imaging software</td><td>Olympus</td><td>cellSens</td><td>imaging software used with compound light microscope</td></tr><tr><td>chimeric zebrafish embryos</td><td>N/A</td><td>N/A</td><td>AB wild-type chimera zebrafish embryos analyzed at 48 hpf&amp;nbsp;</td></tr><tr><td>Compound light microscope</td><td>Olympus</td><td>BX51</td><td>used with digital camera and cellSens imaging software for image capture after IHC</td></tr><tr><td>Confocal fluorescence microscope</td><td>Leica</td><td>DM 2500</td><td>used with digital camera and Leica Application Suite Advanced Fluorescence imaging software for image capture after IF</td></tr><tr><td>Disposable plastic molds, 15 mm x 15 mm x 5 mm</td><td>Ted Pella</td><td>27147-2</td><td></td></tr><tr><td>Digital camera, light microscope</td><td>Olympus</td><td>DP27</td><td></td></tr><tr><td>Digital camera, fluorescence microscope</td><td>Leica</td><td>TCS SPE</td><td></td></tr><tr><td>Ethanol</td><td>Koptec</td><td>V1001</td><td></td></tr><tr><td>Fish gelatin</td><td>Sigma</td><td>G7041</td><td></td></tr><tr><td>Glass slides</td><td>Fisher</td><td>12-550-15</td><td></td></tr><tr><td>Hydrogen peroxide, 30%</td><td>Fisher</td><td>H325-100</td><td></td></tr><tr><td>ImmPRESS HRP Polymer detection kit</td><td>Vector</td><td>MP-7401</td><td>anti-rabbit secondary antibody, HRP polymer</td></tr><tr><td>Leica Application Suite Advanced Fluorescence imaging software</td><td>Leica</td><td>LAS AF 2.3.6</td><td>imaging software used with confocal fluorescence microscope</td></tr><tr><td>Methanol</td><td>Fisher</td><td>A452-4</td><td></td></tr><tr><td>Modified Meyer&#039;s Hematoxylin</td><td>Thermo</td><td>72804</td><td></td></tr><tr><td>Normal Goat Serum</td><td>MP Biomedical</td><td>191356</td><td></td></tr><tr><td>OCT medium</td><td>Tissue-Tek/Fisher</td><td>4583/4585</td><td></td></tr><tr><td>anti-Oregon Green antibody</td><td>Molecular Probes/Life Technologies</td><td>A889</td><td>used at 1:7500 dilution for IHC</td></tr><tr><td>Oregon Green Dextran</td><td>Invitrogen</td><td>P7171</td><td>used at 2.5% in 0.2 M KCl</td></tr><tr><td>Paraformaldehyde, 4%</td><td>Acros Organics</td><td>416780030</td><td>made in lab in 1x PBS</td></tr><tr><td>Permount</td><td>Fisher</td><td>SP15</td><td></td></tr><tr><td>1x phosphate-buffered saline</td><td>made in lab</td><td>N/A</td><td>50 mL 10x PBS, up to 500 mL volume with dH&lt;sub&gt;2&lt;/sub&gt;O</td></tr><tr><td>10x phoshpate-buffered saline</td><td>made in lab</td><td>N/A</td><td>use Cold Spring Harbor protocol</td></tr><tr><td>1x PBSt</td><td>made in lab</td><td>N/A</td><td>50 mL 10x PBS, 1 mL 10% Tween 20, up to 500 mL volume with dH&lt;sub&gt;2&lt;/sub&gt;O</td></tr><tr><td>anti-Ser10 phosphorylated Histone H3 antibody</td><td>Santa Cruz</td><td>sc-8656-R</td><td>used at 1:500 dilution for IF</td></tr><tr><td>Scott&#039;s Tap Water</td><td>Electron Microscopy Sciences</td><td>26070-06</td><td>have used other brands successfully in addition to EMS</td></tr><tr><td>Sucrose</td><td>Amresco</td><td>335</td><td></td></tr><tr><td>Tween-20</td><td>Fisher</td><td>BP337</td><td></td></tr><tr><td>TO-PRO3</td><td>Invitrogen</td><td>T3605</td><td>used at 1:1,000 in 1x PBS for IF</td></tr><tr><td>1x Tris-buffered saline</td><td>made in lab</td><td>N/A</td><td>50 mL 10x TBS, up to 500 mL volume with dH&lt;sub&gt;2&lt;/sub&gt;O</td></tr><tr><td>10x Tris-buffered saline</td><td>made in lab</td><td>N/A</td><td>85 g NaCl, 61 g Tris base, up to 1 L volume with dH&lt;sub&gt;2&lt;/sub&gt;O</td></tr><tr><td>1x TBSt</td><td>made in lab</td><td>N/A</td><td>50 mL 10x TBS, 1 mL 10% Tween 20, up to 500 mL volume with dH&lt;sub&gt;2&lt;/sub&gt;O</td></tr><tr><td>Triton X-100</td><td>Acros Organics</td><td>327371000</td><td></td></tr><tr><td>Vectashield</td><td>Vector</td><td>H-1000</td><td>non-hardening mounting media</td></tr><tr><td>Vector NovaRed substrate</td><td>Vector</td><td>SK-4800</td><td>HRP substrate</td></tr><tr><td>Xylene</td><td>Fisher</td><td>X3P-1GAL</td><td></td></tr></tbody></table>

sequential immunofluorescence and immunohistochemistry on cryosectioned zebrafish embryos

Investigation of intercellular interactions often requires discrete labeling of specific cell populations and precise protein localization. The zebrafish embryo is an excellent tool for examining such interactions with an in vivo model. Whole-mount immunohistochemical and immunofluorescence assays are frequently applied in zebrafish embryos to assess protein expression. However, it can be difficult to achieve accurate mapping of co-localized proteins in three-dimensional space. In addition, some studies may require the use of two antibodies that are not compatible with the same technique (e.g., antibody 1 is only suitable for immunohistochemistry and antibody 2 is only suitable for immunofluorescence). The purpose of the method described herein is to perform sequential immunofluorescence and/or immunohistochemistry on individual cryosections derived from early-stage zebrafish embryos. Here we describe the use of sequential rounds of immunofluorescence, imaging, immunohistochemistry, imaging for a single cryosection in order to achieve precise identification of protein expression at the single-cell level. This methodology is suitable for any study in early-stage zebrafish embryos that requires accurate identification of multiple protein targets in individual cells.

We developed this protocol in order to identify and analyze protein expression in individual donor cells in 48 h post-fertilization chimeric zebrafish embryos that were generated by blastula-to-blastula transplant between AB wild-type zebrafish embryos. Successful analysis of protein expression at the one-cell level required preparation of cryosections with appropriately oriented embryos (Figure 1 and Figure 2) and careful, sequential application of IF and IHC techniques to these cryosections (Figure 3 and Figure 4). Use of specific antibodies to simultaneously detect proliferating cells (anti-pH3, Figure 5A,B) and donor cells (anti-labeled dextran, Figure 5D) can be used to identify and quantify donor cells that are actively proliferating (Figure 5E). It is essential to generate high-quality images after both IF (Figure 2A,B) and IHC (Figure 2D) in order to accurately identify individual cells. An image analysis program should be used to perform image overlay (Figure 5E). Depending on the image analysis program used, it may be possible to perform automated counts of labeled cells should quantification be desired.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59344/59344fig1.jpg" /> 
Figure 1: Preparation of frozen OCT blocks. (A) 50x microscopic view of embryos in OCT in a cryogenic mold prepared for freezing. Red arrow indicates head of zebrafish embryo position against the bottom surface of the mold; black arrow indicates tail pointing up towards the user. (B) Plastic molds with embryos and OCT placed on chilled metal platform. (C) Foam ice bucket placed over plastic molds to create freezing chamber. (D) OCT turns white from transparent once the block is fully frozen. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59344/59344fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59344/59344fig2.jpg" /> 
Figure 2: Cryosectioning of frozen OCT blocks. (A), Application of fresh OCT onto cryostat disc and placement of frozen block on OCT, rotated 180&#176; from the freezing orientation of the block. (B) Side view of block frozen to sectioning disc. (C) View of sectioning block with roll plate in position. (D) Pick-up of sections using a charged slide from the metal platform of the cryostat. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59344/59344fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/59344/59344fig3.jpg" /> 
Figure 3: View of prepared sections after placement on a slide. Arrow indicates hydrophobic barrier, and dotted line delineates the zone containing sections within barrier perimeter. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59344/59344fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/59344/59344fig4.jpg" /> 
Figure 4: Preparation for chromogenic substrate application during IHC. The slide is placed on a flat surface covered with plastic wrap to allow uniform application of the chromogenic substrate while containing any spillage of hazardous material. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59344/59344fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/59344/59344fig5.jpg" /> 
Figure 5: Representative IF and IHC labeling of cryosectioned 48 hpf chimeric zebrafish embryos generated by blastula-to-blastula transplant between AB wild-type zebrafish embryos. (A) IF to detect Ser 10 phosphorylated Histone H3 (anti-pH3, 1:200) expression in 48 hpf chimeric zebrafish embryo. Red = pH3-positive cells; blue = nuclei. Yellow arrows indicate examples of positive cells. (B) Subtraction of the blue nuclear stain in the digital image (ImageJ software) enhances visualization of pH3-positive cells for quantification and image overlay. (C) Negative control (secondary antibody only) for IF assay in cryosectioned 48 hpf chimeric zebrafish embryo. Scale bar represents 50 &#181;m (applicable to all panels). (D) IHC to detect labeled dextran (anti-labeled dextran, 1:7,500) in chimeric 48 hpf zebrafish embryo generated by blastula-to-blastula transplant between AB wild-type zebrafish embryos. Donor embryos were injected with a fluorescently labeled dextran conjugate at the one-cell stage prior to use for blastula-to-blastula transplant. Red indicates cells labeled with a dextran conjugate; blue indicates nuclei. Yellow arrows indicate examples of positive cells. (E) Overlay of panel B (IF for pH3) and panel C (IHC for labeled dextran) showing colocalization of pH3 expression and dextran labeling in individual cells. Yellow arrows indicate examples of double-positive cells. (F) Negative control (secondary antibody only) for IHC assay in cryosectioned 48 hpf chimeric zebrafish embryo. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59344/59344fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Sequential Immunofluorescence and Immunohistochemistry on Cryosectioned Zebrafish Embryos at JoVE.com

Sequential Immunofluorescence and Immunohistochemistry on Cryosectioned Zebrafish Embryos

This protocol demonstrates sequential immunofluorescence and immunohistochemistry on cryosections from early-stage zebrafish embryos enabling precise colocalization analyses in specific cell populations.

Investigation of intercellular interactions often requires discrete labeling of specific cell populations and precise protein localization. The zebrafish ...

sequential-immunofluorescence-immunohistochemistry-on-cryosectioned

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Visualizing Cells, Tissues, and Molecules

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13.9K Views.  NC State University College of Veterinary Medicine. Dit protocol toont een nieuwe procedure aan voor sequentiële immunofluorescentie en immunohistochemie op cryosections van vroege fase zebrafish embroys, die nauwkeurige colokalisatieanalyses in specifieke celpopulaties mogelijk maakt. Het belangrijkste voordeel van dit protocol is de flexibiliteit. Het combineert immunofluorescentie en immunohistochemie technieken met behulp van een enkele cryosection om weefselmorfologie, co-lokalisatie van expressie en antilichaam compatibiliteit te maxima...

Video: Sequential Immunofluorescence and Immunohistochemistry on Cryosectioned Zebrafish Embryos

Imaging Subcellular Structures in the Living Zebrafish Embryo

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such imaging experiments in higher vertebrates such as mammals generally requires surgical access to the system under study. The optical accessibility of embryonic and larval zebrafish allows such invasive procedures to be circumvented and permits imaging in the intact organism. Indeed the zebrafish is now a well-established model to visualize dynamic cellular behaviors using in vivo microscopy in a wide range of developmental contexts from proliferation to migration and differentiation. A more recent development is the increasing use of zebrafish to study subcellular events including mitochondrial trafficking and centrosome dynamics. The relative ease with which these subcellular structures can be genetically labeled by fluorescent proteins and the use of light microscopy techniques to image them is transforming the zebrafish into an in vivo model of cell biology. Here we describe methods to generate genetic constructs that fluorescently label organelles, highlighting mitochondria and centrosomes as specific examples. We use the bipartite Gal4-UAS system in multiple configurations to restrict expression to specific cell-types and provide protocols to generate transiently expressing and stable transgenic fish. Finally, we provide guidelines for choosing light microscopy methods that are most suitable for imaging subcellular dynamics.

Imaging the dynamic behavior of organelles and other subcellular structures in vivo can shed light on their function in physiological and disease conditions. Here, we present methods for genetically tagging two organelles, centrosomes and mitochondria, and imaging their dynamics in living zebrafish embryos using wide-field and confocal microscopy.

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such ...

Developmental Biology

Immunostaining Phospho-epitopes in Ciliated Organs of Whole Mount Zebrafish Embryos

The rapid proliferation of cells, the tissue-specific expression of genes and the emergence of signaling networks characterize early embryonic development of all vertebrates. The kinetics and location of signals - even within single cells - in the developing embryo complements the identification of important developmental genes. Immunostaining techniques are described that have been shown to define the kinetics of intracellular and whole animal signals in structures as small as primary cilia. The techniques for fixing, imaging and processing images using a laser-scanning confocal compound microscope can be completed in as few as 36 hr.
Zebrafish (Danio rerio) is a desirable organism for investigators who seek to conduct studies in a vertebrate species that is affordable and relevant to human disease. Genetic knockouts or knockdowns must be confirmed by the loss of the actual protein product. Such confirmation of protein loss can be achieved using the techniques described here. Clues into signaling pathways can also be deciphered by using antibodies that are reactive with proteins that have been post-translationally modified by phosphorylation. Preserving and optimizing the phosphorylated state of an epitope is therefore critical to this determination and is accomplished by this protocol.
This study describes techniques to fix embryos during the first 72 hr of development and co-localize a variety of relevant epitopes with cilia in the Kupffer&#39;s Vesicle (KV), the kidney and the inner ear. These techniques are straightforward, do not require dissection and can be completed in a relatively short period of time. Projecting confocal image stacks into a single image is a useful means of presenting these data.

Techniques are described to immunostain phospho-epitopes in whole zebrafish embryos and then conduct two-color fluorescent confocal localization in cellular structures as small as primary cilia. The techniques for fixing and imaging can define the location and kinetics of the appearance or activation of specific proteins.

The rapid proliferation of cells, the tissue-specific expression of genes and the emergence of signaling networks characterize early embryonic development ...

Chemistry

Identification of Orexin and Endocannabinoid Receptors in Adult Zebrafish Using Immunoperoxidase and Immunofluorescence Methods

Immunohistochemistry (IHC) is a highly sensitive and specific technique involved in the detection of target antigens in tissue sections with labeled antibodies. It is a multistep process in which the optimization of each step is crucial to obtain the optimum specific signal. Through IHC, the distribution and localization of specific biomarkers can be detected, revealing information on evolutionary conservation. Moreover, IHC allows for the understanding of expression and distribution changes of biomarkers in pathological conditions, such as obesity. IHC, mainly the immunofluorescence technique, can be used in adult zebrafish to detect the organization and distribution of phylogenetically conserved molecules, but a standard IHC protocol is not estasblished. Orexin and endocannabinoid are two highly conserved systems involved in the control of food intake and obesity pathology. Reported here are protocols used to obtain information about orexin peptide (OXA), orexin receptor (OX-2R), and cannabinoid receptor (CB1R) localization and distribution in the gut and brain of normal and diet-induced obese (DIO) adult zebrafish models. Also described are methods for immunoperoxidase and double immunofluorescence, as well as preparation of reagents, fixation, paraffin-embedding, and cryoprotection of zebrafish tissue and preparation for an endogenous activity-blocking step and background counterstaining. The complete set of parameters is obtained from previous IHC experiments, through which we have shown how immunofluorescence can help with the understanding of OXs, OX-2R, and CB1R distribution, localization, and conservation of expression in adult zebrafish tissues. The resulting images with highly specific signal intensity led to the confirmation that zebrafish are suitable animal models for immunohistochemical studies of distribution, localization, and evolutionary conservation of specific biomarkers in physiological and pathological conditions. The protocols presented here are recommended for IHC experiments in adult zebrafish.

Presented here are protocols for immunohistochemical characterization and localization of orexin peptide, orexin receptors, and endocannabinoid receptors in the gut and brains of normal and diet-induced obesity (DIO) adult zebrafish models using immunoperixidase and double immunofluorescence methods.

Immunohistochemistry (IHC) is a highly sensitive and specific technique involved in the detection of target antigens in tissue sections with labeled ...

Biochemistry

Visualizing Multiciliated Cells in the Zebrafish Through a Combined Protocol of Whole Mount Fluorescent In Situ Hybridization and Immunofluorescence

In recent years, the zebrafish embryo has emerged as a popular model to study developmental biology due to traits such as ex utero embryo development and optical transparency. In particular, the zebrafish embryo has become an important organism to study vertebrate kidney organogenesis as well as multiciliated cell (MCC) development. To visualize MCCs in the embryonic zebrafish kidney, we have developed a combined protocol of whole-mount fluorescent in situ hybridization (FISH) and whole mount immunofluorescence (IF) that enables high resolution imaging. This manuscript describes our technique for co-localizing RNA transcripts and protein as a tool to better understand the regulation of developmental programs through the expression of various lineage factors.

Visualizing Multiciliated Cells in the Zebrafish Through a Combined Protocol of Whole Mount Fluorescent In Situ Hybridization and Immunofluorescence

Cilia development is vital to proper organogenesis. This protocol describes an optimized method to label and visualize ciliated cells of the zebrafish.

In recent years, the zebrafish embryo has emerged as a popular model to study developmental biology due to traits such as ex utero embryo development and ...

Zebrafish Whole Mount High-Resolution Double Fluorescent In Situ Hybridization

Whole mount in situ hybridization is one of the most widely used techniques in developmental biology. Here, we present a high-resolution double fluorescent in situ hybridization protocol for analyzing the precise expression pattern of a single gene and for determining the overlap of the expression domains of two genes. The protocol is a modified version of the standard in situ hybridization using alkaline phosphatase and substrates such as NBT/BCIP and Fast Red 1,2. This protocol utilizes standard digoxygenin and fluorescein labeled probes along with tyramide signal amplification (TSA) 3. The commercially available TSA kits allow flexible experimental design as fluorescence emission from green to far-red can be used in combination with various nuclear stains, such as propidium iodide, or fluorescence immunohistochemistry for proteins. TSA produces a reactive fluorescent substrate that quickly covalently binds to moieties, typically tyrosine residues, in the immediate vicinity of the labeled antisense riboprobe. The resulting staining patterns are high resolution in that subcellular localization of the mRNA can be observed using laser scanning confocal microscopy 3,4. One can observe nascent transcripts at the chromosomal loci, distinguish nuclear and cytoplasmic staining and visualize other patterns such as cortical localization of mRNA. Studies in Drosophila indicate that roughly 70% of mRNAs exhibit specific patterns of subcellular localization that frequently correlate with the function of the encoded protein 5. When combined with computer-aided reconstruction of 3D confocal datasets, our protocol allows the detailed analysis of mRNA distribution with sub-cellular resolution in whole vertebrate embryos.

Zebrafish Whole Mount High-Resolution Double Fluorescent In Situ Hybridization

Whole mount in situ hybridization is one of the most widely used techniques in developmental biology. Here, we present a high-resolution double fluorescent in situ hybridization protocol for analyzing the precise expression pattern of a single gene and for determining the overlap of the expression domains of two genes. We include a propidium iodide nuclear counter-stain to highlight tissue organization.

Whole mount in situ hybridization is one of the most widely used techniques in developmental biology.  Here, we present a high-resolution double ...

Whole Mount Immunohistochemistry in Zebrafish Embryos and Larvae

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

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

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

Immunostaining of Dissected Zebrafish Embryonic Heart 

Zebrafish embryo becomes a popular in vivo vertebrate model for studying cardiac development and human heart diseases due to its advantageous embryology and genetics 1,2. About 100-200 embryos are readily available every week from a single pair of adult fish. The transparent embryos that develop ex utero make them ideal for assessing cardiac defects 3. The expression of any gene can be manipulated via morpholino technology or RNA injection 4. Moreover, forward genetic screens have already generated a list of mutants that affect different perspectives of cardiogenesis 5. 
Whole mount immunostaining is an important technique in this animal model to reveal the expression pattern of the targeted protein to a particular tissue 6. However, high resolution images that can reveal cellular or subcellular structures have been difficult, mainly due to the physical location of the heart and the poor penetration of the antibodies. 
 Here, we present a method to address these bottlenecks by dissecting heart first and then conducting the staining process on the surface of a microscope slide. To prevent the loss of small heart samples and to facilitate solution handling, we restricted the heart samples within a circle on the surface of the microscope slides drawn by an immEdge pen. After the staining, the fluorescence signals can be directly observed by a compound microscope. 
Our new method significantly improves the penetration for antibodies, since a heart from an embryonic fish only consists of few cell layers. High quality images from intact hearts can be obtained within a much reduced procession time for zebrafish embryos aged from day 2 to day 6. Our method can be potentially extended to stain other organs dissected from either zebrafish or other small animals.

Immunostaining of Dissected Zebrafish Embryonic Heart

A rapid way to conduct immunostaining of zebrafish embryonic heart is described. Compared to the whole mount immunostaining approach, this method dramatically increases the penetration of the antibodies, which allows obtaining high resolution images that reveal cellular/subcellular structures in the heart within a much reduced processing time.

Zebrafish embryo becomes a popular in vivo vertebrate model for studying cardiac development and human heart diseases due to its advantageous embryology ...

Analysis of Apoptosis in Zebrafish Embryos by Whole-mount Immunofluorescence to Detect Activated Caspase 3

Whole-mount immunofluorescence to detect activated Caspase 3 (Casp3 assay) is useful to identify cells undergoing either intrinsic or extrinsic apoptosis in zebrafish embryos. The whole-mount analysis provides spatial information in regard to tissue specificity of apoptosing cells, although sectioning and/or colabeling is ultimately required to pinpoint the exact cell types undergoing apoptosis. The whole-mount Casp3 assay is optimized for analysis of fixed embryos between the 4-cell stage and 32 hr-post-fertilization and is useful for a number of applications, including analysis of zebrafish mutants and morphants, overexpression of mutant and wild-type mRNAs, and exposure to chemicals. Compared to acridine orange staining, which can identify apoptotic cells in live embryos in a matter of hours, Casp3 and TUNEL assays take considerably longer to complete (2-4 days). However, because of the dynamic nature of apoptotic cell formation and clearance, analysis of fixed embryos ensures accurate comparison of apoptotic cells across multiple samples at specific time points. We have also found the Casp3 assay to be superior to analysis of apoptotic cells by the whole-mount TUNEL assay in regard to cost and reliability. Overall, the Casp3 assay represents a robust, highly reproducible assay in which to analyze apoptotic cells in early zebrafish embryos.

Certain genetic perturbations or exposure to toxins can disrupt normal developmental processes leading to death of specific cell types. The analysis of activated Caspase 3 by whole-mount immunofluorescence in zebrafish embryos reveals stage- and tissue-specific localization of cells specifically undergoing apoptosis.

Whole-mount immunofluorescence to detect activated Caspase 3 (Casp3 assay) is useful to identify cells undergoing either intrinsic or extrinsic apoptosis ...

Semi-automated Imaging of Tissue-specific Fluorescence in Zebrafish Embryos

Zebrafish embryos are a powerful tool for large-scale screening of small molecules. Transgenic zebrafish that express fluorescent reporter proteins are frequently used to identify chemicals that modulate gene expression. Chemical screens that assay fluorescence in live zebrafish often rely on expensive, specialized equipment for high content screening. We describe a procedure using a standard epifluorescence microscope with a motorized stage to automatically image zebrafish embryos and detect tissue-specific fluorescence. Using transgenic zebrafish that report estrogen receptor activity via expression of GFP, we developed a semi-automated procedure to screen for estrogen receptor ligands that activate the reporter in a tissue-specific manner. In this video we describe procedures for arraying zebrafish embryos at 24-48 hours post fertilization (hpf) in a 96-well plate and adding small molecules that bind estrogen receptors. At 72-96 hpf, images of each well from the entire plate are automatically collected and manually inspected for tissue-specific fluorescence. This protocol demonstrates the ability to detect estrogens that activate receptors in heart valves but not in liver.

Described here is a protocol for semi-automated imaging of tissue-specific fluorescence in zebrafish embryos.

Zebrafish embryos are a powerful tool for large-scale screening of small molecules. Transgenic zebrafish that express fluorescent reporter proteins are ...

Flat Mount Preparation for Observation and Analysis of Zebrafish Embryo Specimens Stained by Whole Mount In situ Hybridization

The zebrafish embryo is now commonly used for basic and biomedical research to investigate the genetic control of developmental processes and to model congenital abnormalities. During the first day of life, the zebrafish embryo progresses through many developmental stages including fertilization, cleavage, gastrulation, segmentation, and the organogenesis of structures such as the kidney, heart, and central nervous system. The anatomy of a young zebrafish embryo presents several challenges for the visualization and analysis of the tissues involved in many of these events because the embryo develops in association with a round yolk mass. Thus, for accurate analysis and imaging of experimental phenotypes in fixed embryonic specimens between the tailbud and 20 somite stage (10 and 19 hours post fertilization (hpf), respectively), such as those stained using whole mount in situ hybridization (WISH), it is often desirable to remove the embryo from the yolk ball and to position it flat on a glass slide. However, performing a flat mount procedure can be tedious. Therefore, successful and efficient flat mount preparation is greatly facilitated through the visual demonstration of the dissection technique, and also helped by using reagents that assist in optimal tissue handling. Here, we provide our WISH protocol for one or two-color detection of gene expression in the zebrafish embryo, and demonstrate how the flat mounting procedure can be performed on this example of a stained fixed specimen. This flat mounting protocol is broadly applicable to the study of many embryonic structures that emerge during early zebrafish development, and can be implemented in conjunction with other staining methods performed on fixed embryo samples.

Flat Mount Preparation for Observation and Analysis of Zebrafish Embryo Specimens Stained by Whole Mount In situ Hybridization

The zebrafish embryo is an excellent model for developmental biology research. During embryogenesis, zebrafish develop with a yolk mass, which presents three-dimensional challenges for sample observation and analysis. This protocol describes how to create two-dimensional flat mount preparations of whole mount in situ (WISH) stained zebrafish embryo specimens.

The zebrafish embryo is now commonly used for basic and biomedical research to investigate the genetic control of developmental processes and to model ...

Sequential Immunofluorescence and Immunohistochemistry on Cryosectioned Zebrafish Embryos

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