This work was supported by the National Nature Sciences Foundation of China (Grant number: 81870239, 81741005, 81972999) and The Priority Academic Program Development of Jiangsu Higher Education Institutions.

Wild type zebrafish (AB) used in this study were obtained from the National Zebrafish Resource Center in Wuhan, China. All animal procedures outlined here have been reviewed and approved by the Animal Care Institution of The Ethics Committee of Soochow University.
1. PM2.5 sampling and organic compound extraction
NOTE: PM2.5 was collected in an urban area in Suzhou, China, August 1-7, 2015, as described previously5.
<ol>
	<li>Bake 47 mm quartz membrane filters in a 500 &#176;C muffle furnace for 2 h to remove the organic components.</li>
	<li>Place a filter in a PM2.5 sampler for 24 h of uninterrupted sampling.</li>
	<li>Remove the filter and dry for 24 h at room temperature.</li>
	<li>Quantify the filter with an analytical balance.</li>
	<li>Extract organic components from the filter by Soxhlet extraction using dichloromethane as a solvent19.</li>
	<li>Dry the EOM by a rotary evaporation in a 60 &#176;C water bath and nitrogen flow. Dissolve EOM in DMSO and store at -20 &#176;C.</li>
</ol>
2. Zebrafish embryo collection and treatment
<ol>
	<li>Maintain the zebrafish at 28.5 &#177; 0.5 &#176;C in a re-circulating aquaculture system with a 14 h light and 10 h dark photoperiod cycle.</li>
	<li>Place healthy adult zebrafish into a tank at a 2:1 male to female ratio.</li>
	<li>The next day, collect the embryos and wash them with system water (i.e., zebrafish breeding water).</li>
	<li>Select and randomly divide zebrafish embryos demonstrating a normal development (uniform size, full grains, and no egg coagulation) into 4 groups in individual glass Petri dishes with a diameter of 7 cm (about 50 embryos per dish).</li>
	<li>Treat the embryos with PM2.5 (5 mg/L) in the presence or absence of NAC at 0.25 &#956;M from 2 hpf until 72 hpf. Use DMSO as a vehicle control to a final concentration at 0.1% (v/v).</li>
</ol>
3. Morphological observation of zebrafish embryos and cardiac dissection
<ol>
	<li>At 72 hpf, transfer embryos to glass slides and observe under a stereo microscope. Record heart malformations, such as pericardial edema, altered looping, and decreased size.</li>
	<li>Calculate malformation rates (the percentage of embryos with heart defects out of the total living embryos) and analyze differences between groups using one-way ANOVA followed by Turkey&#39;s Multiple Comparison Test (p &#60; 0.05 = statistically significant).</li>
	<li>Anesthetize the embryos with 0.6 mg/mL MS-222 to immobilize them on glass slides.</li>
	<li>Record heart beats for 30 s and quantify heart rates using ImageJ software5,20.</li>
	<li>Dissect hearts from zebrafish embryos with a disposable syringe needle under a stereo microscope. Caution: Avoid destroying the heart shape.</li>
</ol>
4. Immunofluorescence Assay
<ol>
	<li>To use an immunofluorescence assay to detect PM2.5 induced DNA damage in the heart of zebrafish embryos, use a hydrophobic barrier pen to draw a circle on a clean glass side.</li>
	<li>Add 50 &#956;L of 4% paraformaldehyde to 1.25 mL of Phosphate Buffered Saline (PBS) to make a fixative solution.</li>
	<li>Place 3 dissected hearts into one hydrophobic barrier pen circle and incubate for 20 min at room temperature.</li>
	<li>Decant the solution under the microscope and dry the samples at room temperature for at least 5 min, so that the hearts completely attach to the glass slides.</li>
	<li>Wash the slides three times in PBS with 0.1% Tween 20 (PBST) for 5 min per wash.</li>
	<li>Add 50 &#956;L of bovine serum albumin (BSA) to 1000 &#956;L of PBST to obtain a 5% BSA solution and incubate the slides in a humid chamber for 1 h to block non-specific antibody binding.</li>
	<li>Decant the solution and wash the samples three times with PBST for 5 min per wash.</li>
	<li>Dilute 2 &#956;L of mouse monoclonal antibody against 8-OHdG and 2 &#956;L of rabbit polyclonal antibody against &#947;H2AX in 296 &#956;L of PBST to obtain a working primary antibody cocktail solution.</li>
	<li>Incubate the heart samples with 50 &#956;L of the primary antibody cocktail solution against 8-OHdG and &#947;H2AX in a humidified chamber for at least one hour at room temperature or overnight at 4 &#176;C (Overnight incubation can increase signal intensity).</li>
	<li>Decant the solution and wash the samples three times with PBST for 5 min per wash.</li>
	<li>Dilute 1 &#956;L of FITC-labeled goat anti-mouse secondary antibody and 1 &#956;L of cy3 goat anti-rabbit secondary antibody in 498 &#956;L of PBST to obtain a working secondary antibody cocktail solution and incubate the samples with the secondary antibodies (1:500 in PBST) for 1 h at room temperature in the dark.</li>
	<li>Decant the solution and wash the samples three times with PBS for 5 min per wash protected from light.</li>
	<li>Add 20 &#956;L of DAPI (4&#39;,6-diamidino-2-phenylindole) to the samples for nuclear staining for 30 min at room temperature.</li>
	<li>Apply a coverslip to slide and seal with nail polish to prevent drying and movement. Then image the samples under a fluorescence microscope and quantify the fluorescence signal of heart area using ImageJ software. Calculate the relative changes with the average of DMSO control samples. Determine the statistical significance of the data as in step 3.2.</li>
</ol>

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R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Toxic+assessment+of+urban+atmospheric+particle-bound+PAHs:+relevance+of+composition+and+particle+size+in+Barcelona+(Spain).">Toxic assessment of urban atmospheric particle-bound PAHs: relevance of composition and particle size in Barcelona (Spain).</a> Environmental Pollution. 184, 555-562 (2014).</li><li>Zhang, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Crosstalk+between+AhR+and+wnt/beta-catenin+signal+pathways+in+the+cardiac+developmental+toxicity+of+PM2.5+in+zebrafish+embryos.">Crosstalk between AhR and wnt/beta-catenin signal pathways in the cardiac developmental toxicity of PM2.5 in zebrafish embryos.</a> Toxicology. 355-356, 31-38 (2016).</li><li>Duan, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multi-organ+toxicity+induced+by+fine+particulate+matter+PM2.5+in+zebrafish+(Danio+rerio)+model.">Multi-organ toxicity induced by fine particulate matter PM2.5 in zebrafish (Danio rerio) model.</a> Chemosphere. 180, 24-32 (2017).</li><li>Lorda-Diez, C. 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The authors have nothing to disclose.

Although zebrafish is an excellent vertebrate model for studying the cardiac developmental toxicity of environmental chemicals, due to the small size of the embryo heart, it is difficult to obtain enough protein for western blot analysis. Therefore, we present a sensitive immunofluorescence method for quantifying the protein expression levels of DNA damage biomarkers in the hearts of zebrafish embryos exposed to PM2.5.
During dissection, it is important to keep the integrity of the ...

Air pollution is now a serious environmental problem facing the world. Ambient fine particulate matter (PM2.5), which is one of the most important indicators of air quality, can carry a large number of harmful substances and enter the blood circulatory system, causing serious harm to human health1. Epidemiology studies have demonstrated that PM2.5 exposure can lead to an increased risk of congenital heart defects (CHDs)2,3. Evidence from animal experiments also showed that PM2.5 can cause abnormal cardiac development in zebrafish embryos and the offspring of mice but the molecular mechanisms of the cardiac developmental toxicity of PM2.5 is still largely unknown4,5,6.
DNA damage can cause cell cycle arrest and induce apoptosis, which may extensively destroy the potential of progenitor cells and, consequently, impair heart development7). It has been well documented that environmental pollutants, including PM2.5, have the potential to attack DNA through oxidative stress mechanisms8,9. Both human and zebrafish cardiac development are sensitive to oxidative stress10,11,12. 8-OHdG is an oxidative DNA damage marker, and &#947;H2AX signal is a marker of DNA double strand breaks. N-acetyl-L-cysteine (NAC), a synthetic precursor of intracellular cysteine and glutathione, is widely used as an anti-oxidative compound. In this study, we use NAC to investigate the role of oxidative stress in PM2.5- induced DNA damage13.
Zebrafish as a model vertebrate has been widely used to study cardiac development and human cardiovascular diseases because mechanisms of cardiac development are highly conserved among vertebrates14,15. The advantages of using zebrafish as a model include their small size, strong reproductive ability, and low feeding cost. Of particular interest to these studies, zebrafish embryos do not depend on the circulatory system during early development and can survive severe heart malformation14. Moreover, their transparency allows the entire body to be directly observed under a microscope. Thus, zebrafish embryos provide an outstanding opportunity for assessing the molecular mechanisms involved in the induction of cardiac developmental toxicity as a result of exposure to various environmental chemicals5,16,17. We have previously reported that PM2.5-induced oxidative stress leads to DNA damage and apoptosis, resulting in heart malformations in zebrafish18. In this study, we provide a detailed protocol for investigating PM2.5-induced DNA damage in the heart of zebrafish embryos.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>8-OHdG Antibody</td>
			<td>Santa Cruz Biotechnology, USA</td>
			<td>sc-66036</td>
			<td>Primary antibody</td>
		</tr>
		<tr>
			<td>Analytical balance</td>
			<td>Sartorius,China</td>
			<td>BSA124S</td>
			<td></td>
		</tr>
		<tr>
			<td>BSA</td>
			<td>Solarbio,Beijing,China</td>
			<td>SW3015</td>
			<td>For blocking</td>
		</tr>
		<tr>
			<td>DAPI</td>
			<td>Abcam, USA</td>
			<td>ab104139</td>
			<td>For nuclear counterstain.</td>
		</tr>
		<tr>
			<td>DMSO</td>
			<td>Solarbio,Beijing,China</td>
			<td>D8371</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescence microscope</td>
			<td>Olympus, Japan</td>
			<td>IX73</td>
			<td>For imaging fluorescence signals/</td>
		</tr>
		<tr>
			<td>Goat Anti-Rabbit IgG Cy3</td>
			<td>Carlsbad,USA</td>
			<td>CW0159</td>
			<td>Secondary antibody</td>
		</tr>
		<tr>
			<td>Goat Anti-Rabbit IgG FITC</td>
			<td>Carlsbad,USA</td>
			<td>RS0003</td>
			<td>Secondary antibody</td>
		</tr>
		<tr>
			<td>N-Acetyl-L-cysteine(NAC)</td>
			<td>Adamas-Beta, Shanghai, China</td>
			<td>616-91-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Orbital shaker</td>
			<td>QILINBEIER,China</td>
			<td>TS-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Sigma,China</td>
			<td>P6148</td>
			<td>Make 4% paraformaldehyde for fixation.</td>
		</tr>
		<tr>
			<td>Phosphate Buffered Saline</td>
			<td>HyClone,USA</td>
			<td>SH30256.01</td>
			<td>Prepare 0.1% Tween in PBS for washing.</td>
		</tr>
		<tr>
			<td>PM2.5 sampler</td>
			<td>TianHong,Wuhan, China</td>
			<td>TH-150C</td>
			<td>For 24-hr uninterrupted PM2.5 sampling.</td>
		</tr>
		<tr>
			<td>Re-circulating aquaculture system</td>
			<td>HaiSheng,Shanghai,China</td>
			<td></td>
			<td>The zebrafish was maintained in it.</td>
		</tr>
		<tr>
			<td>Soxhlet extractor</td>
			<td>ZhengQiao,Shanghai, China</td>
			<td>BSXT-02</td>
			<td>For organic components extraction.</td>
		</tr>
		<tr>
			<td>Stereomicroscope</td>
			<td>Nikon,Canada</td>
			<td>SMZ645</td>
			<td>For heart dissection from zebrafish embryos.</td>
		</tr>
		<tr>
			<td>Tricaine methanesulfonate (MS222)</td>
			<td>Sigma,China</td>
			<td>E10521</td>
			<td>To anesthetize zebrafish embryos</td>
		</tr>
		<tr>
			<td>Tween 20</td>
			<td>Sigma,China</td>
			<td>P1379</td>
			<td></td>
		</tr>
		<tr>
			<td>&#947;H2AX Antibody</td>
			<td>Abcam, USA</td>
			<td>ab26350</td>
			<td>Primary antibody</td>
		</tr>
	</tbody>
</table>

using immunofluorescence to detect pm2.5-induced dna damage in zebrafish embryo hearts

Ambient fine particulate matter (PM2.5) exposure can lead to cardiac developmental toxicity but the underlying molecular mechanisms are still unclear. 8-hydroxy-2&#39;deoxygenase (8-OHdG) is a marker of oxidative DNA damage and &#947;H2AX is a sensitive marker for DNA double strand breaks. In this study, we aimed to detect PM2.5-induced 8-OHdG and &#947;H2AX changes in the heart of zebrafish embryos using an immunofluorescence assay. Zebrafish embryos were treated with extractable organic matters (EOM) from PM2.5 at 5 &#956;g/mL in the presence or absence of antioxidant N-acetyl-L-cysteine (NAC, 0.25 &#956;M) at 2 h post fertilization (hpf). DMSO was used as a vehicle control. At 72 hpf, hearts were dissected from embryos using a syringe needle and fixed and permeabilized. After being blocked, samples were probed with primary antibodies against 8-OHdG and &#947;H2AX. Samples were then washed and incubated with secondary antibodies. The resulting images were observed under fluorescence microscopy and quantified using ImageJ. The results show that EOM from PM2.5 significantly enhanced 8-OHdG and &#947;H2AX signals in the heart of zebrafish embryos. However, NAC, acting as a reactive oxygen species (ROS) scavenger, partially counteracted the EOM-induced DNA damage. Here, we present an immunofluorescence protocol for investigating the role of DNA damage in PM2.5-induced heart defects that can be applied to the detection of environmental chemical-induced protein expression changes in the hearts of zebrafish embryos.

This immunofluorescence assay is a sensitive and specific method for measuring protein expression changes in the hearts of zebrafish embryos exposed to environmental chemicals.
In this representative analysis, embryos exposed to PM2.5 in the absence or presence of the antioxidant NAC were evaluated for the presence the presence of heart malformations (Figure 1). As observed, EOM from PM<s...

Watch this Scientific Journal Video about Using Immunofluorescence to Detect PM2.5-induced DNA Damage in Zebrafish Embryo Hearts at JoVE.com

Using Immunofluorescence to Detect PM2.5-induced DNA Damage in Zebrafish Embryo Hearts

This protocol uses an immunofluorescence assay to detect PM2.5-induced DNA damage in the dissected hearts of zebrafish embryos.

Using Immunofluorescence to Detect PM2.5-induced DNA Damage in Zebrafish Embryo Hearts

Ambient fine particulate matter (PM2.5) exposure can lead to cardiac developmental toxicity but the underlying molecular mechanisms are still unclear. ...

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3.6K Views.  Soochow University. This protocol uses an immunofluorescence assay to detect PM2.5-induced DNA damage in the dissected hearts of zebrafish embryos.

Video: Using Immunofluorescence to Detect PM2.5-induced DNA Damage in Zebrafish Embryo Hearts

Analysis of Cardiomyocyte Development using Immunofluorescence in Embryonic Mouse Heart

During heart development, the generation of myocardial-specific structural and functional units including sarcomeres, contractile myofibrils, intercalated discs, and costameres requires the coordinated assembly of multiple components in time and space. Disruption in assembly of these components leads to developmental heart defects. Immunofluorescent staining techniques are used commonly in cultured cardiomyocytes to probe myofibril maturation, but this ex vivo approach is limited by the extent to which myocytes will fully differentiate in culture, lack of normal in vivo mechanical inputs, and absence of endocardial cues. Application of immunofluorescence techniques to the study of developing mouse heart is desirable but more technically challenging, and methods often lack sufficient sensitivity and resolution to visualize sarcomeres in the early stages of heart development. Here, we describe a robust and reproducible method to co-immunostain multiple proteins or to co-visualize a fluorescent protein with immunofluorescent staining in the embryonic mouse heart and use this method to analyze developing myofibrils, intercalated discs, and costameres. This method can be further applied to assess cardiomyocyte structural changes caused by mutations that lead to developmental heart defects.

Mutations that lead to congenital heart defects benefit from in vivo investigation of cardiac structure during development, but high-resolution structural studies in the mouse embryonic heart are technically challenging. Here we present a robust immunofluorescence and image analysis method to assess cardiomyocyte-specific structures in the developing mouse heart.

During heart development, the generation of myocardial-specific structural and functional units including sarcomeres, contractile myofibrils, intercalated ...

Microbead Implantation in the Zebrafish Embryo

The zebrafish has emerged as a valuable genetic model system for the study of developmental biology and disease. Zebrafish share a high degree of genomic conservation, as well as similarities in cellular, molecular, and physiological processes, with other vertebrates including humans. During early ontogeny, zebrafish embryos are optically transparent, allowing researchers to visualize the dynamics of organogenesis using a simple stereomicroscope. Microbead implantation is a method that enables tissue manipulation through the alteration of factors in local environments. This allows researchers to assay the effects of any number of signaling molecules of interest, such as secreted peptides, at specific spatial and temporal points within the developing embryo. Here, we detail a protocol for how to manipulate and implant beads during early zebrafish development.

The zebrafish is an excellent model system for genetic and developmental studies. Bead implantation is a valuable tissue manipulation technique that can be used to interrogate developmental mechanisms by introducing alterations in local cellular environments. This protocol describes how to perform microbead implantation in the zebrafish embryo.

The zebrafish has emerged as a valuable genetic model system for the study of developmental biology and disease. Zebrafish share a high degree of genomic ...

Optimization of the Wound Scratch Assay to Detect Changes in Murine Mesenchymal Stromal Cell Migration After Damage by Soluble Cigarette Smoke Extract

Cell migration is vital to many physiological and pathological processes including tissue development, repair, and regeneration, cancer metastasis, and inflammatory responses. Given the current interest in the role of mesenchymal stromal cells in mediating tissue repair, we are interested in quantifying the migratory capacity of these cells, and understanding how migratory capacity may be altered after damage. Optimization of a rigorously quantitative migration assay that is both easy to customize and cost-effective to perform is key to answering questions concerning alterations in cell migration in response to various stimuli. Current methods for quantifying cell migration, including scratch assays, trans-well migration assays (Boyden chambers), micropillar arrays, and cell exclusion zone assays, possess a range of limitations in reproducibility, customizability, quantification, and cost-effectiveness. Despite its prominent use, the scratch assay is confounded by issues with reproducibility related to damage of the cell microenvironment, impediments to cell migration, influence of neighboring senescent cells, and cell proliferation, as well as lack of rigorous quantification. The optimized scratch assay described here demonstrates robust outcomes, quantifiable and image-based analysis capabilities, cost-effectiveness, and adaptability to other applications.

The capacity to migrate is a key function of many different cell types, including mesenchymal stromal cells (MSCs). However, quantifying alterations in migratory capacity after damage is challenging. This protocol describes an easily adaptable migration assay that uses rigorous statistics to quantify changes in MSC migratory capacity after damage.

Cell migration is vital to many physiological and pathological processes including tissue development, repair, and regeneration, cancer metastasis, and ...

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 ...

Observing Mitotic Division and Dynamics in a Live Zebrafish Embryo

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin condensation, microtubule-kinetochore attachment, chromosome segregation, and cytokinesis in a small time frame. Errors in the delicate process can result in human disease, including birth defects and cancer. Traditional approaches investigating human mitotic disease states often rely on cell culture systems, which lack the natural physiology and developmental/tissue-specific context advantageous when studying human disease. This protocol overcomes many obstacles by providing a way to visualize, with high resolution, chromosome dynamics in a vertebrate system, the zebrafish. This protocol will detail an approach that can be used to obtain dynamic images of dividing cells, which include: in vitro transcription, zebrafish breeding/collecting, embryo embedding, and time-lapse imaging. Optimization and modifications of this protocol are also explored. Using H2A.F/Z-EGFP (labels chromatin) and mCherry-CAAX (labels cell membrane) mRNA-injected embryos, mitosis in AB wild-type, auroraBhi1045, and esco2hi2865 mutant zebrafish is visualized. High resolution live imaging in zebrafish allows one to observe multiple mitoses to statistically quantify mitotic defects and timing of mitotic progression. In addition, observation of qualitative aspects that define improper mitotic processes (i.e., congression defects, missegregation of chromosomes, etc.) and improper chromosomal outcomes (i.e., aneuploidy, polyploidy, micronuclei, etc.) are observed. This assay can be applied to the observation of tissue differentiation/development and is amenable to the use of mutant zebrafish and pharmacological agents. Visualization of how defects in mitosis lead to cancer and developmental disorders will greatly enhance understanding of the pathogenesis of disease.

Mitosis is critical to every living organism and defects often lead to cancer and developmental disorders. Using this imaging protocol and zebrafish as a model system, researchers can visualize mitosis in a live vertebrate organism and the multitude of defects that arise when mitotic processes are defective.

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin ...

Analysis of Coronary Vessels in Cleared Embryonic Hearts

Whole mount visualization of the embryonic coronary plexus from which the capillary and arterial networks will form is rendered problematic using standard microscopy techniques, due to the scattering of imaging light by the thick heart tissue, as these vessels are localized deep within the walls of the developing heart. As optical clearing of tissues using organic solvents such as BABB (1 part benzyl alcohol to 2 parts benzyl benzoate) has been shown to greatly improve the optical penetration depth that can be achieved, we combined clearance of whole, PECAM1-immunostained hearts, with laser-scanning confocal microscopy, in order to obtain high-resolution images of vessels throughout the entire heart. BABB clearance of embryonic hearts takes place rapidly and also acts to preserve the fluorescent signal for several weeks; in addition, samples can be imaged multiple times without loss of signal. This straightforward method is also applicable to imaging other types of blood vessels in whole embryos.

We present a protocol for the analysis of coronary vessels in whole embryonic murine hearts up to E15.5, using standard immunological staining methods followed by optical clearance and confocal microscopy. This technique enables visualization of blood vessels throughout the entire heart without the need for time-consuming analysis of serial sections.

Whole mount visualization of the embryonic coronary plexus from which the capillary and arterial networks will form is rendered problematic using standard ...

Use of Immunolabeling to Analyze Stable, Dynamic, and Nascent Microtubules in the Zebrafish Embryo

Microtubules (MTs) are dynamic and fragile structures that are challenging to image in vivo, particularly in vertebrate embryos. Immunolabeling methods are described here to analyze distinct populations of MTs in the developing neural tube of the zebrafish embryo. While the focus is on neural tissue, this methodology is broadly applicable to other tissues. The procedures are optimized for early to mid-somitogenesis-stage embryos (1 somite to 12 somites), however they can be adapted to a range of other stages with relatively minor adjustments. The first protocol provides a method to assess the spatial distribution of stable and dynamic MTs and perform a quantitative analysis of these populations with image-processing software. This approach complements existing tools to image microtubule dynamics and distribution in real-time, using transgenic lines or transient expression of tagged constructs. Indeed, such tools are very useful, however they do not readily distinguish between dynamic and stable MTs. The ability to image and analyze these distinct microtubule populations has important implications for understanding mechanisms underlying cell polarization and morphogenesis. The second protocol outlines a technique to analyze nascent MTs specifically. This is accomplished by capturing the de novo growth properties of MTs over time, following microtubule depolymerization with the drug nocodazole and a recovery period after drug washout. This technique has not yet been applied to the study of MTs in zebrafish embryos, but is a valuable assay for investigating the in vivo function of proteins implicated in microtubule assembly.

Immunolabeling methods to analyze distinct populations of microtubules in the developing zebrafish brain are described here, which are broadly applicable to other tissues. The first protocol outlines an optimized method for immunolabeling stable and dynamic microtubules. The second protocol provides a method to image and quantify nascent microtubules specifically.

Microtubules (MTs) are dynamic and fragile structures that are challenging to image in vivo, particularly in vertebrate embryos. Immunolabeling methods ...

AFM and Microrheology in the Zebrafish Embryo Yolk Cell

Elucidating the factors that direct the spatio-temporal organization of evolving tissues is one of the primary purposes in the study of development. Various propositions claim to have been important contributions to the understanding of the mechanical properties of cells and tissues in their spatiotemporal organization in different developmental and morphogenetic processes. However, due to the lack of reliable and accessible tools to measure material properties and tensional parameters in vivo, validating these hypotheses has been difficult. Here we present methods employing atomic force microscopy (AFM) and particle tracking with the aim of quantifying the mechanical properties of the intact zebrafish embryo yolk cell during epiboly. Epiboly is an early conserved developmental process whose study is facilitated by the transparency of the embryo. These methods are simple to implement, reliable, and widely applicable since they overcome intrusive interventions that could affect tissue mechanics. A simple strategy was applied for the mounting of specimens, AFM recording, and nanoparticle injections and tracking. This approach makes these methods easily adaptable to other developmental times or organisms.

The lack of tools to measure material properties and tensional parameters in vivo prevents validating their roles during development. We employed atomic force microscopy (AFM) and nanoparticle-tracking to quantify mechanical features on the intact zebrafish embryo yolk cell during epiboly. These methods are reliable and widely applicable avoiding intrusive interventions.

Elucidating the factors that direct the spatio-temporal organization of evolving tissues is one of the primary purposes in the study of development. ...

Following Endocardial Tissue Movements via Cell Photoconversion in the Zebrafish Embryo

During embryogenesis, cells undergo dynamic changes in cell behavior, and deciphering the cellular logic behind these changes is a fundamental goal in the field of developmental biology. The discovery and development of photoconvertible proteins have greatly aided our understanding of these dynamic changes by providing a method to optically highlight cells and tissues. However, while photoconversion, time-lapse microscopy, and subsequent image analysis have proven to be very successful in uncovering cellular dynamics in organs such as the brain or the eye, this approach is generally not used in the developing heart due to challenges posed by the rapid movement of the heart during the cardiac cycle. This protocol consists of two parts. The first part describes a method for photoconverting and subsequently tracking endocardial cells (EdCs) during zebrafish atrioventricular canal (AVC) and atrioventricular heart valve development. The method involves temporally stopping the heart with a drug in order for accurate photoconversion to take place. Hearts are allowed to resume beating upon removal of the drug and embryonic development continues normally until the heart is stopped again for high-resolution imaging of photoconverted EdCs at a later developmental time point. The second part of the protocol describes an image analysis method to quantify the length of a photoconverted or non-photoconverted region in the AVC in young embryos by mapping the fluorescent signal from the three-dimensional structure onto a two-dimensional map. Together, the two parts of the protocol allows one to examine the origin and behavior of cells that make up the zebrafish AVC and atrioventricular heart valve, and can potentially be applied for studying mutants, morphants, or embryos that have been treated with reagents that disrupt AVC and/or valve development.

This protocol describes a method for the photoconversion of Kaede fluorescent protein in endocardial cells of the living zebrafish embryo that enables the tracking of endocardial cells during atrioventricular canal and atrioventricular heart valve development.

During embryogenesis, cells undergo dynamic changes in cell behavior, and deciphering the cellular logic behind these changes is a fundamental goal in the ...

Visualizing the Node and Notochordal Plate In Gastrulating Mouse Embryos Using Scanning Electron Microscopy and Whole Mount Immunofluorescence

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is the formation of the transient organizers, the node and notochordal plate, from cells that have passed through the primitive streak. The proper formation of these signaling centers is essential for the development of the body plan and techniques to visualize them are of high interest to mouse developmental biologists. The node and notochordal plate lie on the ventral surface of gastrulating mouse embryos around embryonic day (E) 7.5 of development. The node is a cup-shaped structure whose cells possess a single slender cilium each. The proper subcellular localization and rotation of the cilia in the node pit determines left-right asymmetry. The notochordal plate cells also possess single cilia albeit shorter than those of the node cells. The notochordal plate forms the notochord which acts as an important signaling organizer for somitogenesis and neural patterning. Because the cells of the node and notochordal plate are transiently present on the surface and possess cilia, they can be visualized using scanning electron microscopy (SEM). Among other techniques used to visualize these structures at the cellular level is whole mount immunofluorescence (WMIF) using the antibodies against the proteins that are highly expressed in the node and notochordal plate. In this report, we describe our optimized protocols to perform SEM and WMIF of the node and notochordal plate in developing mouse embryos to help in the assessment of tissue shape and cellular organization in wild-type and gastrulation mutant embryos.

The node and notochordal plate are transient signaling organizers in developing mouse embryos that can be visualized using several techniques. Here, we describe in detail how to perform two of the techniques to study their structure and morphogenesis: 1) scanning electron microscopy (SEM); and 2) whole mount immunofluorescence (WMIF).

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is ...

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Using Immunofluorescence to Detect PM_2.5-induced DNA Damage in Zebrafish Embryo Hearts