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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G., 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=Characterization+of+zebrafish+mutants+with+defects+in+embryonic+hematopoiesis.">Characterization of zebrafish mutants with defects in embryonic hematopoiesis.</a> Development. 123, 311-319 (1996).</li><li>Amsterdam, A., 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=A+large-scale+insertional+mutagenesis+screen+in+zebrafish.">A large-scale insertional mutagenesis screen in zebrafish.</a> Genes &amp; Development. 13, 2713-2724 (1999).</li><li>Gaiano, N., 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=Insertional+mutagenesis+and+rapid+cloning+of+essential+genes+in+zebrafish.">Insertional mutagenesis and rapid cloning of essential genes in zebrafish.</a> Nature. 383, 829-832 (1996).</li><li>Yeh, J. 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L., Traver, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cellular+dissection+of+zebrafish+hematopoiesis.">Cellular dissection of zebrafish hematopoiesis.</a> Methods in Cell Biology. 133, 11-53 (2016).</li><li>Berrun, A. C., Stachura, D. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+an+In+Vitro+Assay+to+Quantitate+Hematopoietic+Stem+and+Progenitor+Cells+(HSPCs)+in+Developing+Zebrafish+Embryos.">Development of an In Vitro Assay to Quantitate Hematopoietic Stem and Progenitor Cells (HSPCs) in Developing Zebrafish Embryos.</a> Journal of Visualized Experiments. (129), e56836 (2017).</li><li>Westerfield, M., Zon, L. I., Detrich, H. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Essential Zebrafish Methods: Cell and Developmental Biology. , (2009).</li><li>Drummond, I. A., Davidson, A. J., Detrich, H. W., Westerfield, M., Zon, L. 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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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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 Ca2+ and Mg2+</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&#160;</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 ...

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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

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

The zebrafish model is an emerging system for the study of neuromuscular disorders. In the study of neuromuscular diseases, the integrity of the muscle membrane is a critical disease determinant. To date, numerous neuromuscular conditions display degenerating muscle fibers with abnormal membrane integrity; this is most commonly observed in muscular dystrophies. Evans Blue Dye (EBD) is a vital, cell permeable dye that is rapidly taken into degenerating, damaged, or apoptotic cells; in contrast, it is not taken up by cells with an intact membrane. EBD injection is commonly employed to ascertain muscle integrity in mouse models of neuromuscular diseases. However, such EBD experiments require muscle dissection and/or sectioning prior to analysis. In contrast, EBD uptake in zebrafish is visualized in live, intact preparations. Here, we demonstrate a simple and straightforward methodology for performing EBD injections and analysis in live zebrafish. In addition, we demonstrate a co-injection strategy to increase efficacy of EBD analysis. Overall, this video article provides an outline to perform EBD injection and characterization in zebrafish models of neuromuscular disease.

In this study, we describe a straightforward method to perform Evans Blue Dye (EBD) analysis on zebrafish larvae. This technique is a powerful tool for the characterization of skeletal muscle integrity and delineation of zebrafish models of muscular dystrophy, and is a valuable method for the development of novel therapeutics.

The zebrafish model is an emerging system for the study of neuromuscular disorders.  In the study of neuromuscular diseases, the integrity of the muscle ...

Technique to Target Microinjection to the Developing Xenopus Kidney

The embryonic kidney of Xenopus&#160;laevis (frog), the pronephros, consists of a single nephron, and can be used as a model for kidney disease. Xenopus embryos are large, develop externally, and can be easily manipulated by microinjection or surgical procedures. In addition, fate maps have been established for early Xenopus embryos. Targeted microinjection into the individual blastomere that will eventually give rise to an organ or tissue of interest can be used to selectively overexpress or knock down gene expression within this restricted region, decreasing secondary effects in the rest of the developing embryo. In this protocol, we describe how to utilize established Xenopus fate maps to target the developing Xenopus kidney (the pronephros), through microinjection into specific blastomere of 4- and 8-cell embryos. Injection of lineage tracers allows verification of the specific targeting of the injection. After embryos have developed to stage 38 - 40, whole-mount immunostaining is used to visualize pronephric development, and the contribution by targeted cells to the pronephros can be assessed. The same technique can be adapted to target other tissue types in addition to the pronephros.

Technique to Target Microinjection to the Developing Xenopus Kidney

Here, we present a protocol to use fate maps and lineage tracers to target injections into individual blastomeres that give rise to the kidney of Xenopus laevis embryos.

The embryonic kidney of Xenopus&#160;laevis (frog), the pronephros, consists of a single nephron, and can be used as a model for kidney disease. Xenopus ...

Identification Of Erythromyeloid Progenitors And Their Progeny In The Mouse Embryo By Flow Cytometry

Macrophages are professional phagocytes from the innate arm of the immune system. In steady-state, sessile macrophages are found in adult tissues where they act as front line sentinels of infection and tissue damage. While other immune cells are continuously renewed from hematopoietic stem and progenitor cells (HSPC) located in the bone marrow, a lineage of macrophages, known as resident macrophages, have been shown to be self-maintained in tissues without input from bone marrow HSPCs. This lineage is exemplified by microglia in the brain, Kupffer cells in the liver and Langerhans cells in the epidermis among others. The intestinal and colon lamina propria are the only adult tissues devoid of HSPC-independent resident macrophages. Recent investigations have identified that resident macrophages originate from the extra-embryonic yolk sac hematopoiesis from progenitor(s) distinct from fetal hematopoietic stem cells (HSC). Among yolk sac definitive hematopoiesis, erythromyeloid progenitors (EMP) give rise both to erythroid and myeloid cells, in particular resident macrophages. EMP are only generated within the yolk sac between E8.5 and E10.5 days of development and they migrate to the fetal liver as early as circulation is connected, where they expand and differentiate until at least E16.5. Their progeny includes erythrocytes, macrophages, neutrophils and mast cells but only EMP-derived macrophages persist until adulthood in tissues. The transient nature of EMP emergence and the temporal overlap with HSC generation renders the analysis of these progenitors difficult. We have established a tamoxifen-inducible fate mapping protocol based on expression of the macrophage cytokine receptor Csf1r promoter to characterize EMP and EMP-derived cells in vivo by flow cytometry.

While infiltrating macrophages are continuously recruited to adult tissues from circulating precursors, resident macrophages seed their tissue during development, where they are maintained without further input from progenitors. The progenitors for resident macrophages were recently identified. Here, we present methods for the genetic fate mapping of the resident macrophage progenitors.

Macrophages are professional phagocytes from the innate arm of the immune system. In steady-state, sessile macrophages are found in adult tissues where ...

Isolating and Analyzing Cells of the Pancreas Mesenchyme by Flow Cytometry

The pancreas is comprised of epithelial cells that are required for food digestion and blood glucose regulation. Cells of the pancreas microenvironment, including endothelial, neuronal, and mesenchymal cells were shown to regulate cell differentiation and proliferation in the embryonic pancreas. In the adult, the function and mass of insulin-producing cells were shown to depend on cells in their microenvironment, including pericyte, immune, endothelial, and neuronal cells. Lastly, changes in the pancreas microenvironment were shown to regulate pancreas tumorigenesis. However, the cues underlying these processes are not fully defined. Therefore, characterizing the different cell types that comprise the pancreas microenvironment and profiling their gene expression are crucial to delineate the tissue development and function under normal and diseased states. Here, we describe a method that allows for the isolation of mesenchymal cells from the pancreas of embryonic, neonatal, and adult mice. This method utilizes the enzymatic digestion of mouse pancreatic tissue and the subsequent fluorescence-activated cell sorting (FACS) or flow-cytometric analysis of labeled cells. Cells can be labeled by either immunostaining for surface markers or by the expression of fluorescent proteins. Cell isolation can facilitate the characterization of genes and proteins expressed in cells of the pancreas mesenchyme. This protocol was successful in isolating and culturing highly enriched mesenchymal cell populations from the embryonic, neonatal, and adult mouse pancreas.

Here, we describe a method for the isolation of cells in the pancreas microenvironment from embryonic, neonatal and adult mouse tissue, focusing on the isolation of mesenchymal cells. This method allows profiling of cell gene expression and protein secretion in order to elucidate the extrinsic signals that regulate pancreas development, function, and tumorigenesis.

The pancreas is comprised of epithelial cells that are required for food digestion and blood glucose regulation. Cells of the pancreas microenvironment, ...

Chondrogenic Pellet Formation from Cord Blood-derived Induced Pluripotent Stem Cells

Human articular cartilage lacks the ability to repair itself. Cartilage degeneration is thus treated not by curative but by conservative treatments. Currently, efforts are being made to regenerate damaged cartilage with ex vivo expanded chondrocytes or bone marrow-derived mesenchymal stem cells (BMSCs). However, the restricted viability and instability of these cells limit their application in cartilage reconstruction. Human induced pluripotent stem cells (hiPSCs) have received scientific attention as a new alternative for regenerative applications. With unlimited self-renewal ability and multipotency, hiPSCs have been highlighted as a new replacement cell source for cartilage repair. However, obtaining a high quantity of high-quality chondrogenic pellets is a major challenge to their clinical application. In this study, we used embryoid body (EB)-derived outgrowth cells for chondrogenic differentiation. Successful chondrogenesis was confirmed by PCR and staining with alcian blue, toluidine blue, and antibodies against collagen types I and II (COL1A1 and COL2A1, respectively). We provide a detailed method for the differentiation of cord blood mononuclear cell-derived iPSCs (CBMC-hiPSCs) into chondrogenic pellets.

Here, we propose a protocol for chondrogenic differentiation from cord blood mononuclear cell-derived human induced pluripotent stem cells.

Human articular cartilage lacks the ability to repair itself. Cartilage degeneration is thus treated not by curative but by conservative treatments. ...

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 ...

Use of Hematopoietic Stem Cell Transplantation to Assess the Origin of Myelodysplastic Syndrome

Myelodysplastic syndromes (MDS) are a diverse group of hematopoietic stem cell disorders that are defined by ineffective hematopoiesis, peripheral blood cytopenias, dysplasia, and a propensity for transformation to acute leukemia. NUP98-HOXD13 (NHD13) transgenic mice recapitulate human MDS in terms of peripheral blood cytopenias, dysplasia, and transformation to acute leukemia. We previously demonstrated that MDS could be transferred from a genetically engineered mouse with MDS to wild-type recipients by transplanting MDS bone marrow nucleated cells (BMNC). To more clearly understand the MDS cell of origin, we have developed approaches to transplant specific, immunophenotypically defined hematopoietic subsets. In this article, we describe the process of isolating and transplanting specific populations of hematopoietic stem and progenitor cells. Following transplantation, we describe approaches to assess the efficiency of transplantation and persistence of the donor MDS cells.

We describe the use of hematopoietic stem cell transplantation (HSCT) to assess the malignant potential of genetically engineered hematopoietic cells. HSCT is useful for evaluating various malignant hematopoietic cells in vivo as well as generating a large cohort of mice with myelodysplastic syndromes (MDS) or leukemia to evaluate novel therapies.

Myelodysplastic syndromes (MDS) are a diverse group of hematopoietic stem cell disorders that are defined by ineffective hematopoiesis, peripheral blood ...

Real-time Live-cell Flow Cytometry to Investigate Calcium Influx, Pore Formation, and Phagocytosis by P2X7 Receptors in Adult Neural Progenitor Cells

Live-cell flow cytometry is increasingly used among cell biologists to quantify biological processes in a living cell culture. This protocol describes a method whereby live-cell flow cytometry is extended upon to analyze the multiple functions of P2X7 receptor activation in real-time. Using a time module installed on a flow cytometer, live-cell functionality can be assessed and plotted over a given time period to explore the kinetics of calcium influx, transmembrane pore formation, and phagocytosis. This simple method is advantageous as all three canonical functions of the P2X7 receptor can be assessed using one machine, and the gathered data plotted over time provides information on the entire live-cell population rather than single-cell recordings often obtained using technically challenging patch-clamp methods. Calcium influx experiments use a calcium indicator dye, while P2X7 pore formation assays rely on ethidium bromide being allowed to pass through the transmembrane pore formed upon high agonist concentrations. Yellow-green (YG) latex beads are utilized to measure phagocytosis. Specific agonists and antagonists are applied to investigate the effects of P2X7 receptor activity. Individually, these methods can be modified to provide quantitative data on any number of calcium channels and purinergic and scavenger receptors. Taken together, they highlight how the use of real-time live-cell flow cytometry is a rapid, cost-effective, reproducible, and quantifiable method to investigate P2X7 receptor function.

Providing single-cell sensitivity, real-time flow cytometry is uniquely suited to quantify multimodal receptor functions of live cultures. Using adult neural progenitor cells, the P2X7 receptor function was assessed via calcium influx detected by calcium indicator dye, transmembrane pore formation by ethidium bromide uptake, and phagocytosis using fluorescent latex beads.

Live-cell flow cytometry is increasingly used among cell biologists to quantify biological processes in a living cell culture. This protocol describes a ...

Whole-Mount In Situ Hybridization in Zebrafish Embryos and Tube Formation Assay in iPSC-ECs to Study the Role of Endoglin in Vascular Development

Vascular development is determined by the sequential expression of specific genes, which can be studied by performing in situ hybridization assays in zebrafish during different developmental stages. To investigate the role of endoglin(eng) in vessel formation during the development of hereditary hemorrhagic telangiectasia&#160;(HHT), morpholino-mediated targeted knockdown of eng in zebrafish are used to study its temporal expression and associated functions. Here, whole-mount in situ RNA hybridization (WISH) is employed for the analysis of eng and its downstream genes in zebrafish embryos. Also, tube formation assays are performed in HHT patient-derived induced pluripotent stem cell-differentiated&#160;endothelial cells (iPSC-ECs; with eng mutations). A specific signal amplifying system using the whole amount In Situ Hybridization &#8211; WISH provides higher resolution and lower background results compared to traditional methods. To obtain a better signal, the post-fixation time is adjusted to 30 min after probe hybridization. Because fluorescence staining is not sensitive in zebrafish embryos, it is replaced with diaminobezidine (DAB) staining here. In this protocol, HHT&#160;patient-derived iPSC lines containing an eng mutation are differentiated into endothelial cells. After coating a plate with basement membrane matrix for 30 min at 37 &#176;C, iPSC-ECs are seeded as a monolayer into wells and kept at 37 &#176;C for 3 h. Then, the tube length and number of branches are calculated using microscopic images. Thus, with this improved WISH protocol, it is shown that reduced eng expression affects endothelial progenitor formation in zebrafish embryos. This is further confirmed by tube formation assays using iPSC-ECs derived from a patient with HHT. These assays confirm the role for eng in early vascular development.

Presented here is a protocol for whole-mount in situ RNA hybridization analysis in zebrafish and tube formation assay in patient-derived induced pluripotent stem cell-derived endothelial cells to study the role of endoglin in vascular formation.

Vascular development is determined by the sequential expression of specific genes, which can be studied by performing in situ hybridization assays in ...

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 ...