Name: using flow cytometry to detect and quantitate altered blood formation in the developing zebrafish
Uploaded: 2021-04-29T14:00:00.000+00:00
Duration: 7 min 32 s
Description: Watch this Scientific Journal Video about Using Flow Cytometry to Detect and Quantitate Altered Blood Formation in the Developing Zebrafish at JoVE.com

Funding was provided by the National Institutes of Health (NIH: R15 DK114732-01 to D.L.S.), the National Science Foundation (NSF: MRI 1626406 to D.L.S.), and from the Honor&#39;s Program at California State University Chico (to K.F.R.). The authors thank Betsey Tamietti for laboratory management and Kathy Johns for administrative assistance.

The Institutional Animal Care and Use Committee (IACUC) advisory board at California State University, Chico, approved all methods described below.
NOTE: It is advised to treat embryos with 1-phenyl 2-thiourea (PTU; Table 1) at 24 hours postfertilization (hpf) to prevent pigmentation which negatively affects fluorescence discrimination by flow cytometry. All procedures listed below are acceptable for flow cytometry and fluorescence-activated cell sorting (FACS). However, if one&#39;s goal is to culture the hematopoietic cells after FACS, then adhere to sterile practices when processing samples. The protocol for bleaching and preparing embryo samples in a sterile manner has been described previously12.
1. Dechorionation of 48 hpf zebrafish embryos
<ol>
	<li>With embryos (at least 5, but no more than 200 embryos) in a 10 cm plastic Petri dish, tilt the dish so that the embryos sink to the bottom edge. 
	NOTE: It is easy to tilt the dish by removing its lid and placing the lid underneath one edge of the Petri dish.</li>
	<li>Remove and discard as much E3 medium (Table 1) as possible and add 500 &#181;L of dechorionation protease (10 mg/mL). Incubate at room temperature for 5 min.</li>
	<li>After 5 min, pick up the Petri dish (keeping all the embryos at the bottom edge of the plate), and gently tap the side of the Petri dish, allowing embryos to gently rub against the bottom of the plate to completely remove chorions.</li>
	<li>With a squeeze bottle, add ~20 mL of E3 medium to dilute protease. Allow embryos to settle and remove the E3 medium by decanting.</li>
	<li>Repeat step 1.4 3x to remove all traces of the dechorionation protease. 
	NOTE: For each individual sample to analyze, at least 5 embryos are required. This dechorionation procedure can accommodate up to 200 embryos per Petri dish. Samples can be grouped at this stage and separated later if desired. Alternatively, manual dechorionation can be performed by gently tearing the chorions with watchmaker&#39;s forceps. This is advantageous when there are small numbers of embryos. Usually zebrafish will hatch from their chorions by 72 hpf. Severely malformed embryos may not, however, and will require dechorionation (either manual or chemical) after that time.</li>
</ol>
2. Preparation of embryo samples for dissociation
<ol>
	<li>Using a P1000 pipette, place 5&#8722;10 embryos into a single 1.5 mL microcentrifuge tube.</li>
	<li>Remove E3 medium with a pipette and discard. 
	CAUTION: Pipette with care as embryos can easily be discarded during the following steps.</li>
	<li>Add 1 mL of 10 mM dithiothreitol (DTT) in E3 medium to remove the mucus coating surrounding the embryonic zebrafish11,12,13,14. Lay the microcentrifuge tube horizontally, and incubate at room temperature for 30 min.</li>
</ol>
3. Embryo dissociation
<ol>
	<li>Wash the embryos 3x with 1 mL of Dulbecco&#39;s phosphate buffered saline (DPBS) with Ca2+ and Mg2+ to remove traces of DTT. After the last wash add 500 &#181;L of DPBS with Ca2+ and Mg2+ and 5 &#181;L of 5 mg/mL (26 U/mL) dissociation protease. 
	NOTE: Other isotonic buffered solutions may be used, but they must contain Ca2+ and Mg2+ for the dissociation enzyme to function properly.</li>
	<li>Incubate samples at 37 &#176;C on a horizontal orbital shaker at 185 rpm for 60 min. Triturate embryos with a P1000 pipette until samples are fully dissociated. 
	CAUTION: Take care not to overdigest the embryos; some solid tissue should be present, and it should not be completely homogenous. It is possible to overdigest, which will destroy the target blood cells to be observed on the flow cytometer (Supplemental Figure 1). 
	NOTE: This procedure works best for 48&#8722;72 hpf embryos. Later stages may require longer incubation with dissociation protease. Timing for different stages must be determined empirically. There are quicker alternative methods to dissociate embryonic tissue, but it is found empirically that this method, although it takes 60 min, is gentle and thorough enough to allow efficient isolation of live hematopoietic cells.</li>
</ol>
4. Preparation of dissociated embryos for flow cytometry
<ol>
	<li>Pipette the 5 dissociated embryos onto the top reservoir of a 5 mL polystyrene round bottom tube with a 35 &#181;mM cell strainer cap.</li>
	<li>Rinse cells by adding 4 mL of phosphate buffered saline (PBS) with no Ca2+ or Mg2+ to the strainer cap of the 5 mL polystyrene tube containing the filtered cells. 
	NOTE: Other isotonic buffered solutions may also be used, but they should lack Ca2+ or Mg2+ to dilute and render the dissociation enzyme ineffective.</li>
	<li>Centrifuge tubes at 4 &#176;C and 300 x g for 5 min to pellet the cells. With a pipette, remove and discard supernatant. Resuspend the cells in 500 &#181;L of PBS (with no Ca2+ or Mg2+).</li>
</ol>
5. Flow cytometry
<ol>
	<li>Gently vortex the cell suspension and add 1:1,000 red dead cell stain (Table of Materials). 
	NOTE: The red dead cell stain is a cell-permeable dye that allows exclusion of dead cells and debris from target cells and is an excellent choice of dye for dead cell discrimination, because it is excited by the 633 nm laser, which is different from the 488 nm laser used to excite common fluorophores such as green fluorescent protein (GFP) and DsRed. In this way, there is no spectral overlap from dead cells and transgenic blood cells. Propidium iodide (PI) at 1 mg/mL is an inexpensive alternative to the red dead cell stain, but make sure to reserve some samples stained only with PI as well as samples only with the fluorophore of interest to set the instrument compensation. Otherwise, the signals of PI and fluorophores such as GFP will overlap, resulting in false-positive results. If the flow cytometer also has a 405 nm laser, other dyes such as blue dad cell stain are also an excellent choice. 
	CAUTION: When analyzing blood cells, note that after 30 min certain cell types will begin to phagocytose the red dead cell stain. Keep cells protected from light and on ice until analysis and perform flow cytometry within 30 min of adding the dye.</li>
	<li>Flow cytometry analysis 
	NOTE: Every flow cytometer is a little different, but the following steps will assist in preparing samples for analysis on most machines. For users new to flow cytometry, seek the advice of someone that is an expert on the particular piece of equipment. The following instructions pertain to the BD FACSAria Fusion flow cytometer.
	<ol>
		<li>Turn on the cytometer and allow lasers to warm up for 20 min. Empty waste tank, fill sheath tank with appropriate solution (varies based on flow cytometer) and start the fluidics system. 
		NOTE: There is usually a particular startup procedure for each type of flow cytometer; follow that startup procedure carefully.</li>
		<li>In the analysis software, draw five plots (Figure 1A).
		<ol>
			<li>Have the first dot plot measure forward scatter (FSC) on the x axis and side scatter (SSC) on the y axis. Set FSC as linear and SSC as log. 
			NOTE: FSC is a measurement of cell size, and SSC is a measurement of granularity; this will allow the discrimination of cells from debris. Embryonic zebrafish blood cell populations do not separate by size and granularity in the same manner as adult zebrafish blood cells described previously15. Instead they will appear in the same population with all the other digested embryonic cells.</li>
			<li>Set the second plot to examine FSC height (H) versus FSC width (W). Set the third plot to measure SSC (H) versus SSC (W). 
			NOTE: This step is used to exclude doublets (cells that are stuck together when they pass through the laser).</li>
			<li>Set the fourth dot plot to measure the red dead cell stain on the x axis (depending on the cytometer this is usually equivalent to the filter used for allophycocyanin [APC]) and SSC on the y axis. This will allow the discrimination of live cells from dead cells. 
			NOTE: Filter sets of each cytometer can be different. Make sure to use the correct detection filter for the dead cell discrimination dye.</li>
			<li>The fifth plot measures the fluorophore of choice. Visualize this as a histogram of the fluorophore (Figure 1) or use a dot plot (Supplemental Figure 2). 
			NOTE: When examining more than one fluorophore in the same animal, it is recommended to use a dot plot to view both parameters at the same time. If there is any chance that the fluorophores have spectral overlap with each other (Supplemental Figure 2), a dot plot is also recommended. Use the Area (A) settings to calculate the area under the curve generated by the laser pulse of the cytometer for all parameters, because they are more accurate.</li>
		</ol>
		</li>
		<li>Load the sample and reduce the flow rate so that the sample does not run out rapidly. Adjust the FSC and SSC settings so that the bulk of the cell population can be clearly seen (Figure 1A).</li>
		<li>Draw a gate around the cell population and label it cells. Making sure that the second dot plot is gated on cells, exclude doublets by first gating on FSC and then on SSC singlets.</li>
		<li>On the fourth plot adjust the red dead cell stain settings so that there is clearly a negative population. Draw a gate around these cells, and label them live cells. On the fifth plot, gate on live cells and examine fluorophore negative and positive cells. Draw a gate around the positive cells. 
		NOTE: This hierarchical gating strategy examines single fluorophore+ cells that reside in the live cells gate, which also reside in the cell gate. Take care to set gates properly. These gates can be customized depending on the experiment.</li>
		<li>Run each sample, collecting at least 25,000 live cell events.</li>
		<li>Follow the shutdown procedure to turn off the fluidics. Refill sheath tank and empty the waste tank. 
		NOTE: It is often difficult to visualize fluorophore+ cells when the protein expression is low, or the cell population is rare. To ensure the parameters are set properly, fluorophore- sibling embryos should be prepared alongside to properly draw gates and adjust laser voltages (Supplemental Figure 2). If the target cell population is rare, collect more than 25,000 events. With 5&#8722;10 digested animals, one should be able to collect millions of events. The total number of fluorophore+ cells can also be obtained with these data. Simply count the total number of cells in the sample with a hemocytometer and multiply the percentage of fluorophore+ cells to obtain the absolute number of fluorophore+ cells per fish.</li>
	</ol>
	</li>
</ol>

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D.L.S. is a scientific consultant and has received compensation from Finless Foods, Inc. and Xytogen Biotech, Inc. K.F.R. declares no competing interests.

Zebrafish are an excellent model system for studying primitive and definitive vertebrate hematopoiesis. Over the past few decades, multiple assays have been developed and refined, allowing zebrafish to become a quick and economical model for testing drugs, generating and testing genetic mutants, and overall allowing researchers to analyze molecular pathways essential for hematopoiesis. This protocol utilizes embryonic zebrafish which allows quick data collection, the use of less physical space than adult animals, and the...

Blood production (hematopoiesis) is an essential developmental process that first starts in the early embryo. This process begins by generating primitive red blood cells and macrophages directly from mesoderm, and later shifts towards the production of hematopoietic stem and progenitor cells (HSPCs). These stem cells, which are multipotent, generate all of the varieties of mature blood cells in the organism. Capable of self-renewal, the system is continually replenished through these HSPCs. While this process starts early in development, hematopoiesis continues for the life of the animal, providing the ability to transport oxygen to distant sites of the body, to stop bleeding after injury, and to protect the body from infection. The development of this complex system is controlled temporally and spatially during development and any perturbations in blood cell production can be catastrophic for the organism, resulting in anemia, thrombocytopenia, leukopenia, and leukemia.
A popular animal model used for hematopoietic research is the zebrafish (Danio rerio) because they have similar blood development when compared to humans. In fact, many of the genes and molecular pathways used during hematopoiesis are conserved throughout vertebrate evolution, allowing us to learn about human genes by studying zebrafish. Importantly, zebrafish embryos develop outside the body and within 7 days have generated most mature blood cell types, allowing for direct visualization of the hematopoietic system in a short amount of time. Zebrafish are also extremely fecund, which allows researchers to observe a larger number of samples in a short time frame, which is also important for generating reproducible data. The zebrafish&#39;s short generation time and external development provides for easier manipulation and observation during mutagenesis studies1,2,3,4,5 and drug screening6,7,8,9,10. This allows a panel of promising therapeutic compounds for human blood disorders to be quickly and efficiently tested.
Importantly, zebrafish are also genetically amenable, and the genome is sequenced and annotated. This tractability allows reverse genetics techniques such as morpholino- (MO-) mediated knockdown and CRISPR-mediated genetic ablation to be performed. Zebrafish have also proven their utility as a model to perform forward genetic screens; many essential genes and pathways involved in vertebrate blood formation have been discovered in this manner. Numerous methods of observing blood cells have also been developed in zebrafish. While traditional histological staining techniques exist, it is also possible to perform in situ hybridization for blood-specific transcripts. Importantly, numerous transgenic lines of fish also exist whereby fluorescent proteins are expressed by lineage-specific promoters, allowing the labelling of specific blood cells with fluorescent proteins11. This allows researchers to perform up-to-the-minute observation of blood cell genesis, expansion, and regulation in a living organism over time.
Overall, conservation of the hematopoietic system, the presence and easy development of transgenic lines, easy visualization, and short generation time has made the zebrafish an economical, fast, and adaptable model of hematopoiesis. To improve upon the toolbox of techniques available for zebrafish researchers, we developed this assay to robustly quantitate the number of blood cells in embryos. The method involves digesting transgenic animals and performing flow cytometry for fluorescent blood cells. In this way, blood cells from mutant animals, the effect of MO and CRISPR modification, and the effect of small molecules can be quantitatively analyzed in a quick and reproducible manner. These assays are user-friendly and economical ways to enumerate blood cells, allowing examination of their generation, proliferation, and maintenance over time.

<table><tbody><tr><td>1.5 ml MCF tube</td><td>FisherBrand</td><td>05-408-129</td><td></td></tr><tr><td>10 mm Polystyrene easygrip Petri dish</td><td>Corning Falcon</td><td>351008</td><td></td></tr><tr><td>5 ml Polystyrene round bottom tube with cell strainer cap</td><td>Corning Falcon</td><td>352235</td><td></td></tr><tr><td>BD FACSAria Fusion flow cytometer</td><td>BD Biosciences</td><td></td><td></td></tr><tr><td>Dithiolthreitol (DTT)</td><td>Sigma-Aldrich</td><td>646563</td><td></td></tr><tr><td>DPBS (10x) with Ca&lt;sup&gt;2+&lt;/sup&gt; and Mg&lt;sup&gt;2+&lt;/sup&gt;</td><td>Life Technologies</td><td>14080-055</td><td></td></tr><tr><td>FBS 500 mL</td><td>Gemini Bio-Products</td><td>100-108</td><td></td></tr><tr><td>HyClone PBS (1x)</td><td>GE Healthcare Life Sciences</td><td>sh30256.01</td><td></td></tr><tr><td>Librease</td><td>Roche Sigma-Aldrich</td><td>5401119001</td><td>dissociation protease</td></tr><tr><td>Pronase</td><td>Roche Sigma-Aldrich</td><td>11459643001</td><td>dechorionation protease</td></tr><tr><td>SYTOX Red Dead Cell Stain</td><td>Invitrogen&amp;nbsp;</td><td>S34859</td><td></td></tr></tbody></table>

using flow cytometry to detect and quantitate altered blood formation in the developing zebrafish

The diversity of cell lineages that comprise mature blood in vertebrate animals arise from the differentiation of hematopoietic stem and progenitor cells (HSPCs). This is a critical process that occurs throughout the lifespan of organisms, and disruption of the molecular pathways involved during embryogenesis can have catastrophic long-term consequences. For a multitude of reasons, zebrafish (Danio rerio) has become a model organism to study hematopoiesis. Zebrafish embryos develop externally, and by 7 days postfertilization (dpf) have produced most of the subtypes of definitive blood cells that will persist for their lifetime. Assays to assess the number of hematopoietic cells have been developed, mainly utilizing specific histological stains, in situ hybridization techniques, and microscopy of transgenic animals that utilize blood cell-specific promoters driving the expression of fluorescent proteins. However, most staining assays and in situ hybridization techniques do not accurately quantitate the number of blood cells present; only large differences in cell numbers are easily visualized. Utilizing transgenic animals and analyzing individuals with fluorescent or confocal microscopy can be performed, but the quantitation of these assays relies on either counting manually or utilizing expensive imaging software, both of which can make errors. Development of additional methods to assess blood cell numbers would be economical, faster, and could even be automated to quickly assess the effect of CRISPR-mediated genetic modification, morpholino-mediated transcript reduction, and the effect of drug compounds that affect hematopoiesis on a large scale. This novel assay to quantitate blood cells is performed by dissociating whole zebrafish embryos and analyzing the amount of fluorescently labelled blood cells present. These assays should allow elucidation of molecular pathways responsible for blood cell generation, expansion, and regulation during embryogenesis, which will allow researchers to further discover novel factors altered during blood diseases, as well as pathways essential during the evolution of vertebrate hematopoiesis.

To enumerate red blood cells in embryonic zebrafish, lcr:GFP16 embryos were injected at the one-cell-stage with PBS, 7.0 ng/nL ism1 MO, or 7.0 ng/nL ism1 MO with 7.0 ng ism1 mRNA12. At 48 hpf they were digested and subjected to flow cytometry analysis. After analyzing the percentage of GFP+ red blood cells from each sample (each sample is 5 randomly selected embryos; Figure ...

Watch this Scientific Journal Video about Using Flow Cytometry to Detect and Quantitate Altered Blood Formation in the Developing Zebrafish at JoVE.com

Using Flow Cytometry to Detect and Quantitate Altered Blood Formation in the Developing Zebrafish

This assay is a simple method to quantitate hematopoietic cells in developing embryonic zebrafish. Blood cells from dissociated zebrafish are subjected to flow cytometry analysis. This allows the detection of blood defects in mutant animals and after genetic modification.

The diversity of cell lineages that comprise mature blood in vertebrate animals arise from the differentiation of hematopoietic stem and progenitor cells ...

using-flow-cytometry-to-detect-quantitate-altered-blood-formation

Universidad San Sebastian

Research

JoVE Journal

Developmental Biology

5.4K Views.  California State University, Chico. This assay is a simple method to quantitate hematopoietic cells in developing embryonic zebrafish. Blood cells from dissociated zebrafish are subjected to flow cytometry analysis. This allows the detection of blood defects in mutant animals and after genetic modification.

Video: Using Flow Cytometry to Detect and Quantitate Altered Blood Formation in the Developing Zebrafish

Isolation and Characterization of Single Cells from Zebrafish Embryos

The zebrafish (Danio rerio) is a powerful model organism to study vertebrate development. Though many aspects of zebrafish embryonic development have been described at the morphological level, little is known about the molecular basis of cellular changes that occur as the organism develops. With recent advancements in microfluidics and multiplexing technologies, it is now possible to characterize gene expression in single cells. This allows for investigation of heterogeneity between individual cells of specific cell populations to identify and classify cell subtypes, characterize intermediate states that occur during cell differentiation, and explore differential cellular responses to stimuli. This study describes a protocol to isolate viable, single cells from zebrafish embryos for high throughput multiplexing assays. This method may be rapidly applied to any zebrafish embryonic cell type with fluorescent markers. An extension of this method may also be used in combination with high throughput sequencing technologies to fully characterize the transcriptome of single cells. As proof of principle, the relative abundance of cardiac differentiation markers was assessed in isolated, single cells derived from nkx2.5 positive cardiac progenitors. By evaluation of gene expression at the single cell level and at a single time point, the data support a model in which cardiac progenitors coexist with differentiating progeny. The method and work flow described here is broadly applicable to the zebrafish research community, requiring only a labeled transgenic fish line and access to microfluidics technologies.

This protocol describes a method for isolating single cells from zebrafish embryos, enriching for cells of interest, capturing zebrafish cells in microfluidic based single cell multiplex systems, and assessing gene expression from single cells.

The zebrafish (Danio rerio) is a powerful model organism to study vertebrate development. Though many aspects of zebrafish embryonic development have been ...

Development of an In Vitro Assay to Quantitate Hematopoietic Stem and Progenitor Cells (HSPCs) in Developing Zebrafish Embryos

Hematopoiesis is an essential cellular process in which hematopoietic stem and progenitor cells (HSPCs) differentiate into the multitude of different cell lineages that comprise mature blood. Isolation and identification of these HSPCs is difficult because they are defined ex post facto; they can only be defined after their differentiation into specific cell lineages. Over the past few decades, the zebrafish (Danio rerio) has become a model organism to study hematopoiesis. Zebrafish embryos develop ex utero, and by 48 h post-fertilization (hpf) have generated definitive HSPCs. Assays to assess HSPC differentiation and proliferation capabilities have been developed, utilizing transplantation and subsequent reconstitution of the hematopoietic system in addition to visualizing specialized transgenic lines with confocal microscopy. However, these assays are cost prohibitive, technically difficult, and time consuming for many laboratories. Development of an in vitro model to assess HSPCs would be cost effective, quicker, and present fewer difficulties compared to previously described methods, allowing laboratories to quickly assess mutagenesis and drug screens that affect HSPC biology. This novel in vitro assay to assess HSPCs is performed by plating dissociated whole zebrafish embryos and adding exogenous factors that promote only HSPC differentiation and proliferation. Embryos are dissociated into single cells and plated with HSPC-supportive colony stimulating factors that cause them to generate colony forming units (CFUs) that arise from a single progenitor cell. These assays should allow more careful examination of the molecular pathways responsible for HSPC proliferation, differentiation, and regulation, which will allow researchers to understand the underpinnings of vertebrate hematopoiesis and its dysregulation during disease.

Development of an In Vitro Assay to Quantitate Hematopoietic Stem and Progenitor Cells HSPCs in Developing Zebrafish Embryos

Here, we present a simple method to quantitate hematopoietic stem and progenitor cells (HSPCs) in embryonic zebrafish. HSPCs from dissociated zebrafish are plated in methylcellulose with supportive factors, differentiating into mature blood. This allows the detection of blood defects and allows drug screening to be easily conducted.

Hematopoiesis is an essential cellular process in which hematopoietic stem and progenitor cells (HSPCs) differentiate into the multitude of different cell ...

Visualizing the Developing Brain in Living Zebrafish using Brainbow and Time-lapse Confocal Imaging

Development of the vertebrate nervous system requires a precise coordination of complex cellular behaviors and interactions. The use of high resolution in vivo imaging techniques can provide a clear window into these processes in the living organism. For example, dividing cells and their progeny can be followed in real time as the nervous system forms. In recent years, technical advances in multicolor techniques have expanded the types of questions that can be investigated. The multicolor Brainbow approach can be used to not only distinguish among like cells, but also to color-code multiple different clones of related cells that each derive from one progenitor cell. This allows for a multiplex lineage analysis of many different clones and their behaviors simultaneously during development. Here we describe a technique for using time-lapse confocal microscopy to visualize large numbers of multicolor Brainbow-labeled cells over real time within the developing zebrafish nervous system. This is particularly useful for following cellular interactions among like cells, which are difficult to label differentially using traditional promoter-driven colors. Our approach can be used for tracking lineage relationships among multiple different clones simultaneously. The large datasets generated using this technique provide rich information that can be compared quantitatively across genetic or pharmacological manipulations. Ultimately the results generated can help to answer systematic questions about how the nervous system develops.

In vivo imaging is a powerful tool that can be used to investigate the cellular mechanisms underlying nervous system development. Here we describe a technique for using time-lapse confocal microscopy to visualize large numbers of multicolor Brainbow-labeled cells in real time within the developing zebrafish nervous system.

Development of the vertebrate nervous system requires a precise coordination of complex cellular behaviors and interactions. The use of high resolution in ...

Utilizing Larval Zebrafish for Efficient In Vivo Tumor Studies

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

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

Author Spotlight: Advancing Cancer Therapeutics Using Tumor Xenotransplantation in Zebrafish

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

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

Cancer Research

Single Cell Fate Mapping in Zebrafish

The ability to differentially label single cells has important implications in developmental biology. For instance, determining how hematopoietic, lymphatic, and blood vessel lineages arise in developing embryos requires fate mapping and lineage tracing of undifferentiated precursor cells. Recently, photoactivatable proteins which include: Eos1, 2, PAmCherry3, Kaede4-7, pKindling8, and KikGR9, 10 have received wide interest as cell tracing probes. The fluorescence spectrum of these photosensitive proteins can be easily converted with UV excitation, allowing a population of cells to be distinguished from adjacent ones. However, the photoefficiency of the activated protein may limit long-term cell tracking11. As an alternative to photoactivatable proteins, caged fluorescein-dextran has been widely used in embryo model systems7, 12-14. Traditionally, to uncage fluorescein-dextran, UV excitation from a fluorescence lamp house or a single photon UV laser has been used; however, such sources limit the spatial resolution of photoactivation. Here we report a protocol to fate map, lineage trace, and detect single labeled cells. Single cells in embryos injected with caged fluorescein-dextran are photoactivated with near-infrared laser pulses produced from a titanium sapphire femtosecond laser. This laser is customary in all two-photon confocal microscopes such as the LSM 510 META NLO microscope used in this paper. Since biological tissue is transparent to near-infrared irradiation15, the laser pulses can be focused deep within the embryo without uncaging cells above or below the selected focal plane. Therefore, non-linear two-photon absorption is induced only at the geometric focus to uncage fluorescein-dextran in a single cell. To detect the cell containing uncaged fluorescein-dextran, we describe a simple immunohistochemistry protocol16 to rapidly visualize the activated cell. The activation and detection protocol presented in this paper is versatile and can be applied to any model system. 
Note: The reagents used in this protocol can be found in the table appended at the end of the article.

A method is described to photoactivate single cells containing a caged fluorescent protein using two-photon absorption from a Ti:Sapphire femtosecond laser oscillator. To fate map the photoactivated cell, immunohistochemistry is used. This technique can be applied to any cell type.

The ability to differentially label single cells has important implications in developmental biology. For instance, determining how hematopoietic, ...

Biology

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 ...

Blood Collection for Biochemical Analysis in Adult Zebrafish

The zebrafish has been used as an animal model for studies of several human diseases. It can serve as a powerful preclinical platform for studies of molecular events and therapeutic strategies as well as for evaluating the physiological mechanisms of some pathologies1. 
There are relatively few publications related to adult zebrafish physiology of organs and systems2, which may lead researchers to infer that the basic techniques needed to allow the exploration of zebrafish systems are lacking3. Hematologic biochemical values of zebrafish were first reported in 2003 by Murtha and colleagues4 who employed a blood collection technique first described by Jagadeeswaran and colleagues in 1999. Briefly, blood was collected via a micropipette tip through a lateral incision, approximately 0.3 cm in length, in the region of the dorsal aorta5. Because of the minute dimensions involved, this is a high-precision technique requiring a highly skilled practitioner. The same technique was used by the same group in another publication in that same year6. In 2010, Eames and colleagues assessed whole blood glucose levels in zebrafish7. They gained access to the blood by performing decapitations with scissors and then inserting a heparinized microcapillary collection tube into the pectoral articulation. They mention difficulties with hemolysis that were solved with an appropriate storage temperature based on the work Kilpatrick et al.8. When attempting to use Jagadeeswaran's technique in our laboratory, we found that it was difficult to make the incision in precisely the right place as not to allow a significant amount of blood to be lost before collection could be started. 
Recently, Gupta et al.9 described how to dissect adult zebrafish organs, Kinkle et al.10 described how to perform intraperitoneal injections, and Pugach et al.11 described how to perform retro-orbital injections. However, more work is needed to more fully explore basic techniques for research in zebrafish.
The small size of zebrafish presents challenges for researchers using it as an experimental model. Furthermore, given this smallness of scale, it is important that simple techniques are developed to enable researchers to explore the advantages of the zebrafish model.

This paper presents a technique for collection of blood from the dorsal aorta of Zebrafish. It also provides instructions for obtaining serum for use in biochemical analyses, such as tests for determining cholesterol and triglyceride levels.

The zebrafish has been used as an animal model for studies of several human diseases. It can serve as a powerful preclinical platform for studies of ...

Imaging and 3D Reconstruction of Cerebrovascular Structures in Embryonic Zebrafish

Zebrafish are a powerful tool to study developmental biology and pathology in vivo. The small size and relative transparency of zebrafish embryos make them particularly useful for the visual examination of processes such as heart and vascular development. In several recent studies transgenic zebrafish that express EGFP in vascular endothelial cells were used to image and analyze complex vascular networks in the brain and retina, using confocal microscopy. Descriptions are provided to prepare, treat and image zebrafish embryos that express enhanced green fluorescent protein (EGFP), and then generate comprehensive 3D renderings of the cerebrovascular system. Protocols include the treatment of embryos, confocal imaging, and fixation protocols that preserve EGFP fluorescence. Further, useful tips on obtaining high-quality images of cerebrovascular structures, such as removal the eye without damaging nearby neural tissue are provided. Potential pitfalls with confocal imaging are discussed, along with the steps necessary to generate 3D reconstructions from confocal image stacks using freely available open source software.

Imaging of cerebrovascular development in larval zebrafish is described. Techniques to facilitate 3D imaging and modify cerebrovascular development using chemical treatments are also provided.

Zebrafish are a powerful tool to study developmental biology and pathology in vivo.  The small size and relative transparency of zebrafish embryos make ...

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 ...

Reverse Genetic Approach to Identify Regulators of Pigmentation using Zebrafish

Melanocytes are specialized neural crest-derived cells present in the epidermal skin. These cells synthesize melanin pigment that protects the genome from harmful ultraviolet radiations. Perturbations in melanocyte functioning lead to pigmentary disorders such as piebaldism, albinism, vitiligo, melasma, and melanoma. Zebrafish is an excellent model system to understand melanocyte functions. The presence of conspicuous pigmented melanocytes, ease of genetic manipulation, and availability of transgenic fluorescent lines facilitate the study of pigmentation. This study employs the use of wild-type and transgenic zebrafish lines that drive green fluorescent protein (GFP) expression under mitfa and tyrp1 promoters that mark various stages of melanocytes. 
Morpholino-based silencing of candidate genes is achieved to evaluate the phenotypic outcome on larval pigmentation and is applicable to screen for regulators of pigmentation. This protocol demonstrates the method from microinjection to imaging and fluorescence-activated cell sorting (FACS)-based dissection of phenotypes using two candidate genes, carbonic anhydrase 14 (Ca14) and a histone variant (H2afv), to comprehensively assess the pigmentation outcome. Further, this protocol demonstrates segregating candidate genes into melanocyte specifiers and differentiators that selectively alter melanocyte numbers and melanin content per cell, respectively.

Regulators of melanocyte functions govern visible differences in the pigmentation outcome. Deciphering the molecular function of the candidate pigmentation gene poses a challenge. Herein, we demonstrate the use of a zebrafish model system to identify candidates and classify them into regulators of melanin content and melanocyte number.

Melanocytes are specialized neural crest-derived cells present in the epidermal skin. These cells synthesize melanin pigment that protects the genome from ...

Using Flow Cytometry to Detect and Quantitate Altered Blood Formation in the Developing Zebrafish

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Isolation and Characterization of Single Cells from Zebrafish Embryos

Development of an In Vitro Assay to Quantitate Hematopoietic Stem and Progenitor Cells (HSPCs) in Developing Zebrafish Embryos

Visualizing the Developing Brain in Living Zebrafish using Brainbow and Time-lapse Confocal Imaging

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

Single Cell Fate Mapping in Zebrafish

Immunostaining of Dissected Zebrafish Embryonic Heart

Blood Collection for Biochemical Analysis in Adult Zebrafish

Imaging and 3D Reconstruction of Cerebrovascular Structures in Embryonic Zebrafish

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

Reverse Genetic Approach to Identify Regulators of Pigmentation using Zebrafish