Funding was provided by the National Institutes of Health (NIH: K01-DK087814-01A1 to D.L.S.), the California State University Program for Education &amp; Research in Biotechnology (CSUPERB: Molecular Control of the Vertebrate Hematopoietic Niche to D.L.S.) and from the Graduate Studies Office at California State University Chico (to A.C.B.).

The Institutional Animal Care and Use Committee (IACUC) advisory board at California State University, Chico, approved all methods described below.
1. Bleach 48 hpf Zebrafish Embryos
<ol>
	<li>With 15 cm (w) x 15 cm (l) x 7 cm (h) plastic containers create 3 wash stations: 1) with&#160;1 mL of 5% bleach in 1000 mL&#160;embryo medium (E3; see Table 1), 2) with sterile E3, and 3) with sterile E3. Ensure that each container has 500 mL of solution to submerge the embryos in. 
	NOTE: For each condition tested, at least 10 embryos will be needed. This washing procedure can accommodate up to 200 embryos.</li>
	<li>With a transfer pipette, place embryos into a tea strainer and submerge eggs in wash station #1 for 5 min. Remove the tea strainer (with embryos inside) and place into wash station #2 for 5 min; repeat for wash station #3 for a final 5 min.</li>
	<li>After the last wash, rinse embryos from tea strainer by turning it upside down over a clean 10 cm Petri dish and gently rinsing embryos off the tea strainer with sterile E3 and a transfer pipette.</li>
</ol>
2. Dechorionate Embryos
<ol>
	<li>With a transfer pipette, remove and discard as much E3 as possible from the Petri dish and add 500 &#956;L of dechorionation protease (10 mg/mL) to embryos. Incubate at room temperature for 5 min. Gently tap the side of the Petri dish to completely remove chorions.</li>
	<li>With a serological&#160;pipette, add 20&#160;mL of sterile E3 to dilute protease. Allow embryos to settle, and remove the E3 with a transfer pipette. Repeat this wash step 3 times to remove all traces of the dechorionation protease.</li>
</ol>
3. Preparing Embryo Samples
<ol>
	<li>Using a P1000 pipette, place 10 embryos into a single sterile 1.5 mL microcentrifuge tube.</li>
	<li>Remove E3 with a pipette and discard. 
	Caution: Pipette with care as embryos can easily be displaced and discarded during the following wash steps.</li>
</ol>
4. Preparing Embryos for Dissociation
<ol>
	<li>Transfer samples to laminar flow hood, and wash embryos by adding 1 mL sterile E3. Allow embryos to settle to bottom of tube and remove supernatant with a P1000 pipette. Repeat for a total of 3 washes.</li>
	<li>After last wash, remove E3 and discard. Add 1 mL of 10 mM dithiothreitol (DTT) in E3 to remove the mucus coating (and any spores, yeast, or bacteria that may be trapped in it) surrounding the embryonic zebrafish. Lay microcentrifuge tube horizontally, and incubate at room temperature in laminar flow hood for 25 min.</li>
</ol>
5. Embryo Dissociation
<ol>
	<li>Wash sample 3 times with 1 mL of sterile DPBS (with Ca2+ and Mg2+). After last wash add 500 &#956;L of DPBS (with Ca2+ and Mg2+) and add 5 &#956;L of 5 mg/mL (26 U/mL) dissociation protease. 
	Caution: This enzyme needs Ca2+ and Mg2+, so ensure that the DPBS contains these ions.</li>
	<li>Incubate samples at 37 &#176;C on a horizontal orbital shaker at 180 rpm for 60 min. Place samples in the laminar flow hood and triturate embryos with a P1000 until samples are fully dissociated. 
	Caution: Check embryos periodically to determine when they become dissociated. The embryo solution should have some amount of tissue present; it should not be completely homogenous. It is possible to over-digest, which will destroy the HSPCs (see Supplemental Figure 1).</li>
</ol>
6. Preparation of Dissociated Embryos
<ol>
	<li>Pipette the 10 dissociated embryos onto the top reservoir of a 5 mL polystyrene round bottom tube with a 35 &#956;m cell strainer cap.</li>
	<li>With a pipette, rinse the Petri tube with sterile PBS (with no Ca2+ or Mg2+) and transfer solution to 5 mL polystyrene tube containing filtered cells from step 6.1. Repeat until 4 mL of liquid is present in the 5 mL polystyrene tube. 
	NOTE: PBS, DPBS, or other isotonic buffered solutions may be used at this stage, as long as they do not contain Ca2+ or Mg2+</li>
	<li>Centrifuge tubes at 4 &#176;C and 300 x g for 5 min to pellet the homogenized cell sample. With a pipette, remove and discard supernatant from the round bottom tube. Take care to not disrupt the cells pelleted at the bottom of the tube. Resuspend the cells in 100 &#956;L of PBS.</li>
</ol>
7. Preparation of Methylcellulose
<ol>
	<li>Generate 1x complete methylcellulose (Table 1) stock solution and add 2.5 mL to sterile round-bottom 14 mL tubes with 3 mL syringes and 16 G needles. Use one tube for each condition tested.</li>
	<li>Add cytokines, small molecules, or other agents to be investigated to each sample.
	<ol>
		<li>For myeloid differentiation, add 1% Carp serum and 0.3 mg/mL recombinant zebrafish granulocyte colony stimulating factor (Gcsf)34 to methylcellulose medium. See Svoboda et al.37&#160;for full description on how to generate carp serum and recombinant cytokines.</li>
		<li>For erythroid differentiation, add 1% Carp serum and 0.1 mg/mL recombinant zebrafish erythropoietin (Epo)38&#160;to methylcellulose medium.</li>
		<li>To examine multilineage progenitors, add Epo and Gcsf. 
		Caution: Cytokines and additives should not total more than 10% of the total volume, as the medium will not be viscous enough to discern individual colonies.</li>
	</ol>
	</li>
</ol>
8. Addition of Dissociated Embryos to Methylcellulose
<ol>
	<li>Using a pipette, add 100 &#956;L of the dissociated 10 embryos to the surface of the prepared methylcellulose with recombinant cytokines and carp serum. Cap lid and vortex gently to fully homogenize sample. Each tube should now contain the dissociated tissue of 10 embryos.</li>
	<li>Using 3 mL syringes and 16 G needles, aliquot 1 mL of sample into 2 separate 35 mm Petri dishes. Make sure that the sample is fully dispersed throughout the Petri dish. Repeat for each sample.</li>
	<li>Place 35 mm Petri dishes with cells in methylcellulose into a 15 cm Petri dish. To each 15 cm dish, add one 35 mm Petri plate with 5 mL of sterile water to humidify the samples. Incubate at 32 &#176;C and 5% CO2 for 7 - 10 days.</li>
</ol>
9. Visualization of HSPC-derived Colony Forming Units (CFUs)
<ol>
	<li>After incubating 7 days, place samples onto an inverted microscope at 40 - 100X for visualization and enumeration. At this point, individual colonies can be isolated with a pipette and subjected to staining&#160;and/or gene analysis.</li>
</ol>

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The authors have nothing to disclose.

The zebrafish model system has become an efficient, effective, and inexpensive model for studying primitive and definitive vertebrate hematopoiesis. Generation of assays that are fast, inexpensive, and present few technical difficulties can be utilized for testing small molecules, analyzing mutant embryos, and elucidating molecular pathways important for HSPC biology. In vitro plating of HSPCs from adult zebrafish is an effective method to study mutagenesis, cytokines, and hematopoietic defects. This protocol builds upon those studies, but utilizes embryonic zebrafish. Applying these assays to embryonic zebrafish allows for quicker data collection, the use of less physical space, the use of less drugs (for drug screens), and permits testing mRNA overexpression and/or morpholino (MO) knockdown of specific genetic pathways. Importantly, it also allows the study of stages of hematopoiesis that only occur during early vertebrate development.
Due to the flexibility of this assay, it can easily be modified to meet many diverse needs. One modification of this protocol is the modulation of developmental timing; modifying the age of embryos utilized allows the quantitation of HSPC generation over time. This can also allow the study of distinct subsets of HSPCs such as erythromyeloid progenitors6, which arise before 36 hpf, versus bona fide HSCs3,4,5,8, which arise afterwards. No one has yet been able to isolate and identify more restricted zebrafish HSPC subsets in the developing embryo, but these assays allow these experiments. It is essential that when changing the developmental stages of the animals, the amount of cells added into the methylcellulose is also optimized; too few animals will yield no colonies, while too many will create an uncountable plate of HSPC colonies. Adding different cytokines is another modification that can be adapted to specific experiments. Adding Epo will allow the quantitation of erythroid colonies, while Gcsf will allow enumeration of myeloid colonies. Adding both of these cytokines will allow multi-lineage CFU generation. Importantly, these assays permit testing of putative cytokines and their effect on HSPC proliferation and differentiation. These assays can also be modified to perform large scale drug testing; embryos can be exposed to different concentrations of compounds at different time points during development to see if the chemicals have a positive or negative effect on blood production. A final way that these experiments can be modified is that embryos can be injected at the one-cell stage with specific mRNA or MOs, which will overexpress or knockdown specific genes, respectively. After developing for 24 - 48 h, these embryos can be subjected to these assays to determine if gain- or loss-of-function of specific genes play an essential role in HSPC formation, proliferation, and differentiation. It is also important to note that utilizing different transgenic strains of zebrafish will aid in the speed and ease of collecting and interpreting data. For instance, if one is interested in erythroid development, it is easy to perform these experiments in gata1:DsRed embryos40, which have the promoter of the erythroid-specific transgenic Gata1 transcription factor driving DsRed fluorescence. In this way, when analyzing erythroid colonies, fluorescence microscopy can be utilized. Likewise, these studies can be performed on animals with multiple transgenes present to look for multiple cell lineages at once. This technique would be useful for identifying multilineage HSPCs; although erythroid and myeloid colonies are easily distinguishable by morphology, it is difficult to identify colonies that contain bipotential progenitors without visualizing the presence of multiple lineage-specific transgene expression in the same colony.
Although these assays have many benefits, it is essential to set up adequate controls every time these experiments are performed; minor variations in technique could change the results and interpretations of the findings. For example, it is possible to get different numbers of HSPCs when you perform the experiments on different days if your technique is not precise; to deal with this possibility, one should always include an appropriate control. If an experiment is performed to test the effect of a MO on development, then control embryos (either uninjected or injected with a control MO) that are from the same clutch should also be processed alongside the experimental sample. If the experiment is designed to test the effect of a particular drug, then untreated samples should be included as a control. Finally, if testing the effects of a specific cytokine on development, then animals from the same clutch should also be processed and examined at the same time without having the cytokine added to the culture.
While this in vitro assay is straightforward to perform and the results are easy to interpret, some potential problems can arise during the procedure. If no colonies are seen after 7 days in culture, there are several issues that may have occurred. First, check to make sure that the cytokines are pure and at proper concentrations; depending on how the cytokines have been produced it may be necessary to alter concentrations and optimize their activity. Another issue that commonly arises is over-digesting the embryos, which destroys the HSPCs; take care at the digestion step, and alter the timing if no colonies are repeatedly seen. Finally, the mixing of methylcellulose and addition of carp serum could be the issue, so make sure that all ingredients were added and mixed correctly. In addition to technical issues, it is important to realize that colonies may need longer to develop under certain circumstances; often small colonies are much more visible after an additional 3 days in culture. Going past 10 days is not recommended, as the methylcellulose starts to dry out shortly thereafter. Another common issue is that sometimes colonies tend to collect around the edges of the 35 mm Petri dishes; scanning the plates thoroughly, especially around the edges, is essential. Also, it is important to remember that these colonies are developing in a thick, viscous material; when focusing the microscope, adjust the focus up and down to observe slightly different planes- the colonies will not be sitting on the bottom of the plastic dishes.
These in vitro assays have plenty of advantages, but they also have a few limitations. The largest issue is that these assays cannot prove definitively that a progenitor is an HSC- that can only be proven by long-term reconstitution of irradiated animals. Additionally, not all cytokines have been defined and investigated in zebrafish yet, which limits the types of HSPCs that can be investigated. For instance, no zebrafish lymphoid-supportive cytokines have been identified yet, preventing B and T cell production in these assays. Furthermore, it is unclear when culturing multipotent HSPCs if the addition of certain types (or amounts) of specific cytokines can skew the differentiation down certain hematopoietic pathways. Another potential caveat is that this assay cannot effectively discern if HSPCs are actually present in a sample, but for some reason are unable to differentiate (or are otherwise not viable). As with most in vitro assays, further experiments should always be performed to validate the findings. Barring these issues, much information can be gleaned from these clonal assays.
These assays are significant, because they allow quick and efficient screening of hematopoietic defects. Researchers can knock down certain genetic pathways with MOs, and plate digested animals to see if the gene plays a role in hematopoiesis. Additionally, mutant fish (previously generated or newly generated in mutagenesis screens) can be assessed quickly and easily. These assays also allow efficient drug screening assays on live, whole organisms to identify the modulation of HSPC biology while also being able to observe if the drugs have a negative effect on development or survival. These assays also allow quantitation of HSPCs in the developing zebrafish at different timepoints. While multiple studies have attempted to quantitate HSC numbers in adult7,27,30,41 and embryonic zebrafish3,4,42,43, no studies have quantitated the different amounts of lineage-restricted HSPCs in zebrafish. Additionally, these assays can be utilized to examine and modulate molecular pathways important in blood cell formation and differentiation, a process that remains enigmatic.

Hematopoiesis is the process of making the multitude of mature blood cells required for an organism&#39;s survival. It is a key developmental process that involves the differentiation of hematopoietic stem cells (HSCs) into a variety of developmentally restricted cell types that comprise mature blood. These HSCs must self-renew so that the system is never exhausted and they must persist from early embryonic development until death. In vertebrates, constant differentiation and proliferation of hematopoietic stem and progenitor cells (HSPCs) are needed to adequately replenish the majority of blood cells that are post-mitotic and being recycled every day. HSCs generate mature blood cells by first differentiating into subsets of restricted progenitor cells; common lymphoid progenitors (CLPs)1, which eventually produce T, B, and NK cells, and common myeloid progenitors (CMPs)2 that generate granulocytes, erythrocytes, macrophages, and megakaryocytes. These progenitors are committed to generating specific cell lineages, and further differentiate into more developmentally restricted progenitor cells such as myeloid erythroid progenitors (MEPs) that generate erythrocytes and platelets, or granulocyte macrophage progenitors (GMPs) that generate basophils, eosinophils, neutrophils, and macrophages2. Identifying and isolating these progenitors allows the identification of important molecular pathways involved in hematopoietic differentiation and many hematopoietic diseases such as leukemia arise when these progenitor cells fail to properly differentiate.
Over the past few decades, the zebrafish (Danio rerio) model system has become a key research tool for embryonic and adult hematopoietic studies. Zebrafish are amenable to genetic analysis and are the phylogenetically lowest vertebrate model species that have a similar vasculature and hematopoietic system to humans. Zebrafish embryos develop ex utero, and within 48 hours post fertilization (hpf) generate HSPCs3,4,5,6,7,8. Zebrafish are also highly fecund, with females laying over 100 eggs in a single clutch, allowing for large sample sizes and experimental replication. Zebrafish embryos are optically transparent, allowing for microscopic visualization of the hematopoietic system. Several fluorescent transgenic lines of zebrafish marking HSCs such as runx1:EGFP fish9, cd41:EGFP fish10, and kdrl:mCherry; cmyb:GFP3 double-positive animals, allow for live, real-time visualization of HSC emergence and expansion in vivo3,4,7,8,9. The zebrafish&#39;s quick generation time and development ex utero has led to its use in mutagenesis studies11,12,13,14,15 and drug screening16,17,18,19,20 for compounds that hold therapeutic promise for human blood disorders. Overall, conservation of the hematopoietic system, the presence and easy development of transgenic lines, and quick regeneration time has made the zebrafish an inexpensive, quick, flexible, and ideal model for hematopoietic studies.
Numerous methods of isolating and testing HSCs have been developed in mammalian hematopoietic systems. Investigators can utilize a combination of cell surface receptors to mark HSCs21,22,23,24, as well as exploit the ability of HSCs to efflux dye25,26. After they are labeled, fluorescence-activated cell sorting (FACS) allows their physical separation. Proving that a cell is an HSC requires irradiating a host animal to destroy endogenous HSPCs, transplanting putative HSCs, and observing long-term, multi-lineage reconstitution of all mature blood cell types. These assays work well in mice, as there are numerous cell-surface antibodies against hematopoietic cells and inbred mouse strains that facilitate immune matching for transplantation. However, few zebrafish hematopoietic cell-surface antibodies have been generated27, hindering the identification and isolation of HSCs. The most common way to mark and isolate zebrafish HSCs is with transgenic animals, whereby a cell-specific promoter sequence is driving a fluorescent protein&#39;s expression. Studies have visualized and enumerated HSCs in the ventral wall of the dorsal aorta with microscopy utilizing this technique3,4,8,9. Other laboratories have generated clonal strains of zebrafish28,29 and have performed successful transplants in MHC-matched animals30. However, these techniques are cost prohibitive to many laboratories, are technically difficult, and are time consuming. To address these issues, laboratories have generated several in vitro assays to test for the presence, the proliferation rates, and the differentiation capacity of HSPCs31,32,33,34,35,36. These assays show proliferation and differentiation of HSPCs in vitro31,32,33,34,35,36, the rescue of hematopoietic defects36, and an efficient method for discovering and testing cytokines33,34,35. They have also been utilized to identify genes responsible for HSPC biology31,32. In this study, we take these assays a step further, allowing the quantitation of HSPCs in a developing zebrafish embryo. These assays can also be utilized to quantitate the number of HSPCs in mutant animals and animals treated with hematopoietic-disruptive drugs. In essence, these assays are fast, present few technical challenges, and are inexpensive ways to quantitate HSPC numbers, examine their proliferation, and investigate blocks in differentiation.

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			<td height="40" style="height:40px;width:341px;">10% Bovine Serum Albumin in Iscove&#39;s MDM</td>
			<td style="width:115px;">StemCell Technologies&#160;</td>
			<td style="width:119px;">9300</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:341px;">DPBS (10x) with Calcium2+ and Magnesium2+</td>
			<td style="width:115px;">Life Technologies</td>
			<td style="width:119px;">14080-055</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:341px;">HyClone PBS (1x)</td>
			<td style="width:115px;">GE Healthcare Life Sciences</td>
			<td style="width:119px;">sh30256.01</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:341px;">&#160;DMEM 500 mL</td>
			<td style="width:115px;">Corning CellGrow</td>
			<td style="width:119px;">10-017-CV</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:341px;">&#160;Ham&#39;s F12 500 mL</td>
			<td style="width:115px;">Corning CellGrow</td>
			<td style="width:119px;">10-080-CV</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:341px;">FBS 500 mL</td>
			<td style="width:115px;">Gemini Bio-Products</td>
			<td style="width:119px;">100-108</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:341px;">HEPES 100 mL (1 M)</td>
			<td style="width:115px;">Gibco life technologies</td>
			<td style="width:119px;">15-630-080</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:341px;">Penicillin/streptomycin (5000 U/mL 
			and 5000 mg/mL) with L-Glutamine (200 mM)</td>
			<td style="width:115px;">Corning Mediatech</td>
			<td style="width:119px;">30-009-CI</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:341px;">Gentamycin Sulfate 10 mL (50 mg/mL)</td>
			<td style="width:115px;">Corning Mediatech</td>
			<td style="width:119px;">30-005-CR</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:341px;">1.5 mL MCF tube</td>
			<td style="width:115px;">FisherBrand</td>
			<td style="width:119px;">05-408-129</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:341px;">3 mL 23gx1 injection needle with Luer lock</td>
			<td style="width:115px;">BD Safety Glide</td>
			<td style="width:119px;">305905</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:341px;">5 mL polystyrene round bottom tube with cell strainer cap</td>
			<td style="width:115px;">Corning Falcon</td>
			<td style="width:119px;">352235</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:341px;">Methocel MC</td>
			<td style="width:115px;">Sigma-Aldrich</td>
			<td style="width:119px;">64630</td>
			<td></td>
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		<tr height="40">
			<td height="40" style="height:40px;width:341px;">14 mL Polystyrene round bottom tube</td>
			<td style="width:115px;">Corning Falcon</td>
			<td style="width:119px;">352057</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:341px;">10 mm polystyrene easygrip Petri dish</td>
			<td style="width:115px;">Corning Falcon</td>
			<td style="width:119px;">351008</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:341px;">Librease TM</td>
			<td style="width:115px;">Roche Sigma-Aldrich</td>
			<td>5401119001</td>
			<td>dissociation protease</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:341px;">Pronase</td>
			<td style="width:115px;">Roche Sigma-Aldrich</td>
			<td style="width:119px;">11459643001</td>
			<td>dechorionation protease</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:341px;">15 cm Petri dish</td>
			<td style="width:115px;">Corning Falcon</td>
			<td style="width:119px;">351058</td>
			<td></td>
		</tr>
		<tr height="80">
			<td height="80" style="height:80px;width:341px;">AB</td>
			<td style="width:115px;">Zebrafish International Resource Center (ZIRC)</td>
			<td style="width:119px;">ZL1</td>
			<td>zebrafish strain used</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:341px;">Dithiolthreitol (DTT)</td>
			<td style="width:115px;">Sigma-Aldrich</td>
			<td>646563</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

To assess HSPC numbers in embryonic zebrafish, 48 hpf embryos were digested, plated in methylcellulose with exogenous hematopoietic-supportive growth factors, and incubated for 7 days (Figure 1A). After 7 days, colony forming units (CFUs) were enumerated (Figure 1B) and imaged (Figure 1C). By controlling the different cytokines added, one can control the types of colonies produced: Epo is necessary for erythroid differentiation and Gcsf is necessary for myeloid differentiation. Carp serum is analogous to fetal bovine serum&#39;s use in mammalian tissue culture; it is a source of general growth factors. In Figure 1, Epo, Gcsf, and carp serum were added, allowing the enumeration of erythroid, myeloid, and mixed erythromyeloid colonies. These colonies can be discerned due to their shape and color; the erythroid colonies are tight, small, compact, and have a red color due to hemoglobinization. The myeloid colonies are larger and have more motile cells on their edges, giving them a ruffled appearance.
Successful plating of embryonic zebrafish treated with HSPC-supportive factors will yield CFUs after 7 - 10 days of incubation. Importantly, the quantity of CFUs directly correlates with the number of embryos plated (Figure 1B). Plating either 4 or 8 embryos per mL (which correlates to starting with 10 or 20 embryos at the beginning of the experiment) yields approximately 1,200 or 2,400 CFUs, respectively. Our protocol suggests using 10 embryos, as it is difficult and time consuming to count 2,400 CFUs when performing a larger experiment. Due to the methodology of digesting whole embryos, miscellaneous debris is present in the methylcellulose, but can easily be discerned from actual colonies with higher magnification under the microscope. Other cellular debris may also be present, but the only CFUs present will be HSPC-derived due to the addition of HSPC-supportive cytokines. If CFUs are small and not easily seen, the cultures can be incubated up to 10 days to encourage increased growth. However, increased incubation will not yield more CFUs; if CFUs are not present, this indicates a loss of cytokine activity, over digestion of embryos (and subsequent destruction of HSPCs; see Supplemental Figure 1), or improper plating of samples. It is always important to run proper controls every time this experiment is performed to ensure that slight variations in technique do not yield false differences in HSPC enumeration. For example, if an experiment is performed to analyze the number of HSPCs present in morphant embryos, a sample with normal siblings should be prepared alongside. If an experiment is performed to compare the effect of a drug on the development of HSPCs a sample with untreated siblings should be prepared and analyzed alongside.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56836/56836fig1.jpg" /> 
Figure 1: Enumerating HSPC-derived CFUs from 48 hpf zebrafish embryos. (A) Schematic overview of embryo plating on methylcellulose with HSPC-supportive cytokines for generation of hematopoietic CFUs. 48 hpf zebrafish embryos were dissociated and plated on 1% complete methylcellulose with erythropoietin (Epo), granulocyte colony stimulating factor (Gcsf), and carp serum, allowing erythromyeloid differentiation. (B) Enumeration of colony forming units (CFUs) from samples plated at concentrations of 4 or 8 embryos per mL (starting the experiment with 10 or 20 embryos, respectively). Enumeration was done at 100X on an inverted microscope. Bars indicate average numbers of CFUs, and error bars indicate standard deviation. (C) Colony morphology representative of embryo plating on methylcellulose. The colony&#39;s morphology indicates the types of cells that comprise the colony; images are representative erythroid and myeloid colonies. CFUs photographed at 400X. Scale bars = 50 &#956;m.&#160;&#160;<a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56836/56836fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Supplemental Figure 1" src="/files/ftp_upload/56836/56836supfig1.jpg" /> 
Supplemental Figure 1: Digestion of embryos with dissociation protease. (A) Images of 10 embryos before digestion (left; undigested), after adequate digestion (middle; digested), and after excess digestion (right; over digested). (B) Zoomed in images (7.3X) of digested and over digested tubes in (A). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56836/56836supfig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Development of an In Vitro Assay to Quantitate Hematopoietic Stem and Progenitor Cells HSPCs in Developing Zebrafish Embryos at JoVE.com

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.

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 ...

development-an-vitro-assay-to-quantitate-hematopoietic-stem

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

7.9K Views.  California State University, Chico. 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.

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

Direct Induction of Human Neural Stem Cells from Peripheral Blood Hematopoietic Progenitor Cells

Human disease specific neuronal cultures are essential for generating in vitro models for human neurological diseases. However, the lack of access to primary human adult neural cultures raises unique challenges. Recent developments in induced pluripotent stem cells (iPSC) provides an alternative approach to derive neural cultures from skin fibroblasts through patient specific iPSC, but this process is labor intensive, requires special expertise and large amounts of resources, and can take several months. This prevents the wide application of this technology to the study of neurological diseases. To overcome some of these issues, we have developed a method to derive neural stem cells directly from human adult peripheral blood, bypassing the iPSC derivation process. Hematopoietic progenitor cells enriched from human adult peripheral blood were cultured in vitro and transfected with Sendai virus vectors containing transcriptional factors Sox2, Oct3/4, Klf4, and c-Myc. The transfection results in morphological changes in the cells which are further selected by using human neural progenitor medium containing basic fibroblast growth factor (bFGF) and vascular endothelial growth factor (VEGF). The resulting cells are characterized by the expression for neural stem cell markers, such as nestin and SOX2. These neural stem cells could be further differentiated to neurons, astroglia and oligodendrocytes in specified differentiation media. Using easily accessible human peripheral blood samples, this method could be used to derive neural stem cells for further differentiation to neural cells for in vitro modeling of neurological disorders and may advance studies related to the pathogenesis and treatment of those diseases.

A method was developed to directly derive human neural stem cells from hematopoietic progenitor cells enriched from peripheral blood cells.

Human disease specific neuronal cultures are essential for generating in vitro models for human neurological diseases. However, the lack of access to ...

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 ...

Generation of Parabiotic Zebrafish Embryos by Surgical Fusion of Developing Blastulae

Surgical parabiosis of two animals of different genetic backgrounds creates a unique scenario to study cell-intrinsic versus cell-extrinsic roles for candidate genes of interest, migratory behaviors of cells, and secreted signals in distinct genetic settings. Because parabiotic animals share a common circulation, any blood or blood-borne factor from one animal will be exchanged with its partner and vice versa. Thus, cells and molecular factors derived from one genetic background can be studied in the context of a second genetic background. Parabiosis of adult mice has been &#160;used extensively to research aging, cancer, diabetes, obesity, and brain development. More recently, parabiosis of zebrafish embryos has been used to study the developmental biology of hematopoiesis. In contrast to mice, the transparent nature of zebrafish embryos permits the direct visualization of cells in the parabiotic context, making it a uniquely powerful method for investigating fundamental cellular and molecular mechanisms. The utility of this technique, however, is limited by a steep learning curve for generating the parabiotic zebrafish embryos. This protocol provides a step-by-step method on how to surgically fuse the blastulae of two zebrafish embryos of different genetic backgrounds to investigate the role of candidate genes of interest. In addition, the parabiotic zebrafish embryos are tolerant to heat shock, making temporal control of gene expression possible. This method does not require a sophisticated set-up and has broad applications for studying cell migration, fate specification, and differentiation in vivo during embryonic development.

This protocol provides step-by-step instruction on how to generate parabiotic zebrafish embryos of different genetic backgrounds. When combined with the unparalleled imaging capabilities of the zebrafish embryo, this method provides a uniquely powerful means to investigate cell-autonomous versus non-cell-autonomous functions for candidate genes of interest.

Surgical parabiosis of two animals of different genetic backgrounds creates a unique scenario to study cell-intrinsic versus cell-extrinsic roles for ...

Directed Differentiation of Primitive and Definitive Hematopoietic Progenitors from Human Pluripotent Stem Cells

One of the major goals for regenerative medicine is the generation and maintenance of hematopoietic stem cells (HSCs) derived from human pluripotent stem cells (hPSCs). Until recently, efforts to differentiate hPSCs into HSCs have predominantly generated hematopoietic progenitors that lack HSC potential, and instead resemble yolk sac hematopoiesis.&#160;These resulting hematopoietic progenitors may have limited utility for in vitro disease modeling of various adult hematopoietic disorders, particularly those of the lymphoid lineages. However, we have recently described methods to generate erythro-myelo-lymphoid multilineage definitive hematopoietic progenitors from hPSCs using a stage-specific directed differentiation protocol, which we outline here. Through enzymatic dissociation of hPSCs on basement membrane matrix-coated plasticware, embryoid bodies (EBs) are formed. EBs are differentiated to mesoderm by recombinant BMP4, which is subsequently specified to the definitive hematopoietic program by the GSK3&#946; inhibitor, CHIR99021. Alternatively, primitive hematopoiesis is specified by the PORCN inhibitor, IWP2. Hematopoiesis is further driven through the addition of recombinant VEGF and supportive hematopoietic cytokines. The resulting hematopoietic progenitors generated using this method have the potential to be used for disease and developmental modeling, in vitro.

Here, we present human pluripotent stem cell (hPSC) culture protocols, used to differentiate hPSCs into CD34+ hematopoietic progenitors. This method uses stage-specific manipulation of canonical WNT signaling to specify cells exclusively to either the definitive or primitive hematopoietic program.

One of the major goals for regenerative medicine is the generation and maintenance of hematopoietic stem cells (HSCs) derived from human pluripotent stem ...

Induction of Hypoxia in Living Frog and Zebrafish Embryos

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and zebrafish embryos. Our system comprises a chamber featuring a simple setup that is nevertheless robust to induce and maintain a specific oxygen concentration and temperature in any experimental solution of choice. The presented system is very cost-effective but highly functional, it allows induction and sustainment of hypoxia for direct experiments in vivo and for various time periods up to 48 h.
To monitor and study the effects of hypoxia, we have employed two methods - measurement of levels of hypoxia-inducible factor 1alpha (HIF-1&#945;) in whole embryos or specific tissues and determination of retinal stem cell proliferation by 5-ethynyl-2&#8242;-deoxyuridine (EdU) incorporation into the DNA. HIF-1&#945; levels can serve as a general hypoxia marker in the whole embryo or tissue of choice, here embryonic retina. EdU incorporation into the proliferating cells of embryonic retina is a specific output of hypoxia induction. Thus, we have shown that hypoxic embryonic retinal progenitors decrease proliferation within 1 h of incubation under 5% oxygen of both frog and zebrafish embryos.
Once mastered, our setup can be employed for use with small aquatic model organisms, for direct in vivo experiments, any given time period and under normal, hypoxic or hyperoxic oxygen concentration or under any other given gas mixture.

We introduce a novel hypoxic chamber system for use with aquatic organisms such as frog and zebrafish embryos. Our system is simple, robust, cost-effective and allows the induction and sustainment of hypoxia in vivo and for up to 48 h. We present 2 reproducible methods to monitor the effectiveness of hypoxia.

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and ...

Single-cell Photoconversion in Living Intact Zebrafish

Animal and plant tissue is composed of distinct populations of cells. These cells interact over time to build and maintain the tissue and can cause disease when disrupted. Scientists have developed clever techniques to investigate characteristics and natural dynamics of these cells within intact tissue by expressing fluorescent proteins in subsets of cells. However, at times, experiments require more selected visualization of cells within the tissue, sometimes at the single-cell or population-of-cells manner. To achieve this and visualize single cells within a population of cells, scientists have utilized single-cell photoconversion of fluorescent proteins. To demonstrate this technique, we show here how to direct UV light to an Eos-expressing cell of interest in an intact, living zebrafish. We then image those photoconverted Eos+ cells 24 h later to determine how they changed in the tissue. We describe two techniques: single cell photoconversion and photoconversions of populations of cell. These techniques can be used to visualize cell-cell interactions, cell-fate and differentiation, and cell migrations, making it a technique that is applicable in numerous biological questions.

Here, we present a protocol to show how cell photoconversion is achieved through UV exposure to specific areas expressing the fluorescent protein, Eos, in living animals.

Animal and plant tissue is composed of distinct populations of cells. These cells interact over time to build and maintain the tissue and can cause ...

Computational Analysis of the Caenorhabditis elegans Germline to Study the Distribution of Nuclei, Proteins, and the Cytoskeleton

The Caenorhabditis elegans (C. elegans) germline is used to study several biologically important processes including stem cell development, apoptosis, and chromosome dynamics. While the germline is an excellent model, the analysis is often two dimensional due to the time and labor required for three-dimensional analysis. Major readouts in such studies are the number/position of nuclei and protein distribution within the germline. Here, we present a method to perform automated analysis of the germline using confocal microscopy and computational approaches to determine the number and position of nuclei in each region of the germline. Our method also analyzes germline protein distribution that enables the three-dimensional examination of protein expression in different genetic backgrounds. Further, our study shows variations in cytoskeletal architecture in distinct regions of the germline that may accommodate specific spatial developmental requirements. Finally, our method enables automated counting of the sperm in the spermatheca of each germline. Taken together, our method enables rapid and reproducible phenotypic analysis of the C. elegans germline.

Computational Analysis of the Caenorhabditis elegans Germline to Study the Distribution of Nuclei, Proteins, and the Cytoskeleton

We present an automated method for three-dimensional reconstruction of the Caenorhabditis elegans germline. Our method determines the number and position of each nucleus within the germline and analyses germline protein distribution and cytoskeletal structure.

The Caenorhabditis elegans (C. elegans) germline is used to study several biologically important processes including stem cell development, apoptosis, and ...

Assessment of Oxidative Damage in the Primary Mouse Ocular Surface Cells/Stem Cells in Response to Ultraviolet-C (UV-C) Damage

The ocular surface is subjected to regular wear and tear due to various environmental factors. Exposure to UV-C radiation constitutes an occupational health hazard. Here, we demonstrate the exposure of primary stem cells from the mouse ocular surface to UV-C radiation. Reactive oxygen species (ROS) formation is the readout of the extent of oxidative stress/damage. In an experimental in vitro setting, it is also essential to assess the percentage of dead cells generated due to oxidative stress. In this article, we will demonstrate the 2&#39;,7&#39;-Dichlorofluoresceindiacetate (DCFDA) staining of UV-C exposed mouse primary ocular surface stem cells and their quantification based on the fluorescent images of DCFDA staining. DCFDA staining directly corresponds to ROS generation. We also demonstrate the quantification of dead and live cells by simultaneous staining with propidium iodide (PI) and Hoechst 3332 respectively and the percentage of DCFDA (ROS positive) and PI positive cells.

Assessment of Oxidative Damage in the Primary Mouse Ocular Surface Cells/Stem Cells in Response to Ultraviolet-C UV-C Damage

This protocol demonstrates the simultaneous detection of reactive oxygen species (ROS), live cells, and dead cells in live primary cultures from mouse ocular surface cells. 2&#39;,7&#39;-Dichlorofluoresceindiacetate, propidium iodide, and Hoechst staining are used to assess the ROS, dead cells, and live cells, respectively, followed by imaging and analysis.

The ocular surface is subjected to regular wear and tear due to various environmental factors. Exposure to UV-C radiation constitutes an occupational ...

Analysis of Hematopoietic Stem Progenitor Cell Metabolism

Hematopoietic stem progenitor cells (HSPCs) have distinct metabolic plasticity, which allows them to transition from their quiescent state to a differentiation state to sustain demands of the blood formation. However, it has been difficult to analyze the metabolic status (mitochondrial respiration and glycolysis) of HSPCs due to their limited numbers and lack of optimized protocols for non-adherent, fragile HSPCs. Here, we provide a set of clear, step-by-step instructions to measure metabolic respiration (oxygen consumption rate; OCR) and glycolysis (extracellular acidification rate; ECAR) of murine bone marrow-LineagenegSca1+c-Kit+ (LSK) HSPCs. This protocol provides a higher amount of LSK HSPCs from murine bone marrow, improves the viability of HSPCs during incubation, facilitates extracellular flux analyses of non-adherent HSPCs, and provides optimized injection protocols (concentration and time) for drugs targeting oxidative phosphorylation and glycolytic pathways. This method enables the prediction of the metabolic status and the health of HSPCs during blood development and diseases.

Hematopoietic stem progenitor cells (HSPCs) transition from a quiescent state to a differentiation state due to their metabolic plasticity during blood formation. Here, we present an optimized method for measuring mitochondrial respiration and glycolysis of HSPCs.

Hematopoietic stem progenitor cells (HSPCs) have distinct metabolic plasticity, which allows them to transition from their quiescent state to 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 ...