This work is supported by a grant from the MRC to M. F-S.

1. Development of the embryos
<ol>
 <li>When they are laid, eggs are clustered because of attachment filaments on the chorion. To let embryos develop normally, it is necessary to separate eggs. Tangle and cut attachment filaments by holding the attachment filaments with two forceps.</li>
 <li>After unclustering, eggs are separated from feces and algae and transferred to fresh embryo medium at a maximum density of 40 eggs per 6cm Petri dish.</li>
 <li>Medaka embryos develop slightly slower than zebrafish at 27&deg;C. The timing of hatching is different between the two species; medaka embryos hatch from the chorion in 7 days and immediately start to swim and eat, whereas zebrafish embryos hatch in 2 days but start to swim and eat in 5-6 days. Development of medaka is staged according to Iwamatsu's staging4. </li>
 <li>The development of medaka embryos can be conveniently adjusted to experimental plans by selecting the appropriate temperature. Development of medaka embryos can be stopped at 4&deg;C in early development. After stage 24 when heart beating starts, development can be slowed using a minimum temperature of 18&deg;C. </li>
 <li>The timing of appearance of organs/tissues is slightly different in medaka compared with zebrafish, i.e. in medaka somitogenesis occurs after the onset of brain development whereas in zebrafish somitegenesis precedes brain development.</li>
</ol>
2. Removing the chorion
The chorion of medaka consists of two protective layers with a hard inner layer and a soft outer surface. Thus, a two-step protease treatment employing pronase and hatching enzyme is necessary to remove this chorion.
Once dechorionated, embryos should be kept in 1X BSS. Semi-sterile conditions will enhance the successful culture of dechorionated embryos, especially when longer periods of observation are required. These include using sterile solutions (e.g. sterilized 1X BSS with antibiotics) and tools sterilized with 70% ethanol followed by rinsing with 1X BSS.
As dechorionated medaka embryos are softer and more fragile than dechorionated zebrafish embryos, extra care must be taken to ensure they do not contact air or bubbles in the pipette, as this will cause immediate collapse. To ensure minimal damage to embryos, a wide-mouthed heat polished glass pipette with pipette pump should be used for transferring embryos and a hair loop should be utilized to orientate embryos for observation. Non-adherent Petri dishes should be used to prevent embryos from attaching to surfaces.
<ol>
 <li>Prior to dechorionation it should be checked that eggs have been sufficiently separated and cleaned-up.</li>
 <li>Since both pronase and hatching enzymes are proteinases, they should be kept on ice and minimize exposure of embryos to these enzymes.</li>
 <li>Transfer eggs to sandpaper (p2000 grit size, waterproof) placed in the lid of a 9cm Petri dish. Remove excess medium but ensure a sufficient volume remains as to prevent drying of embryos.</li>
 <li>Gently roll embryos for around 45-60 seconds to remove some of the outer surface hairs and lightly score the surface of the chorion (Figure 1). Transfer embryos back to original Petri dish and examine. 
 
 When rolling embryos on sandpaper use the forefinger, applying minimal pressure and keeping finger parallel to surface of sand paper. Do not roll more than 5-7 embryos at once. This precaution will minimize the risk of crushing embryos beneath each other. 
 
 </li>
 <li>Replace egg medium in dish with 20mg/ml pronase and incubate embryos for 40-60 minutes at 27&deg;C. 
 
 Ensure embryos are sufficiently covered by pronase and a lid is present on the dish. Unused pronase should be kept on ice during this step to minimize self-digestion and can be reused until activity is lost (approximately 1-2 weeks). 
 
 </li>
 <li>Recover pronase for reuse and wash embryos 5X in embryo medium to remove traces of pronase as this will inactivate the hatching enzyme to be added.</li>
 <li>Remove embryo medium and cover embryos with hatching enzyme, ensuring embryos sit as a monolayer in the dish. 
 
 If eggs sit on top of one another in the dish, those at the bottom will be crushed as the chorion dissolves. Keep unused hatching enzyme on ice. 
 
 </li>
 <li>Incubate embryos at 27&deg;C and after 15 minutes periodically check the progress of hatching using a stereomicroscope. 
 
 It will be seen that a number of lunar crater-like holes begin to appear in the inner layer of the chorion, which soon dissolves following this leaving the soft outer layer of the chorion easily removed manually. The entire process of hatching may take 15-60 minutes. 
 
 </li>
 <li>As soon as embryos come out from the chorion, transfer embryos to a Petri dish containing 1X BSS. 
 
 Ensure not to transfer hatching enzyme to the 1X BSS by touching the tip of the pipette onto the surface of the BSS allowing the embryos to gently roll out. 
 
 </li>
 <li>Once all embryos are transferred from hatching enzyme, make a final transfer to another fresh dish of 1X BSS. 
 
 This ensures embryos are not exposed to any remnants of hatching enzyme, as this will damage exposed embryos. Once in this final dish any embryos still possessing the outer layer of the chorion can be manually liberated using sterilized micro-forceps under the stereomicroscope. 
 
 If embryos need to be developed following dechorionation, penicillin/streptomycin should be added to the BSS to prevent bacterial growth. 
 
 </li>
</ol>
<img src="/files/ftp_upload/2055/2055fig1.jpg" alt="Figure 1" /> 
 Figure 1. Comparison of rolled and unrolled embryos during dechorionation. Observe how the rolled embryos in panel B lack the hairs seen on the unrolled embryos in panel A.
3. Mounting dechorionated embryos
Agarose embedding is useful for longer periods of imaging (e.g. time-lapse imaging) for live embryos as well as for detailed observations of fixed embryos. During gastrulation and early organogenesis (stage 14 to stage 28), medaka embryos exhibit waves of rhythmic contractile movements across the periderm, a tissue layer covering both the developing embryo and the yolk 5. Embryos are treated with 3.5mM 1-heptanol to stop contractile movements 6.
To stop movement of embryos after stage 28 (64hpf), embryos are anaesthetized by adding drops of Tricaine mesilate (TMS) before embedding. TMS is also added to the agarose (add only a minimal amount, this dose needs to be optimized, approximately a 1:25 dilution).
<ol>
 <li>Thaw a small amount of 3% low gelling temperature agarose (in 1X BSS) by heating to 37&deg;C and maintain at this temperature. </li>
 <li>The following steps need to be carried out swiftly to ensure that the agarose does not solidify before orientating the embryo. Using a wide-mouthed glass pipette transfer enough molten agarose to fill the cap depression of an Eppendorf tube.</li>
 <li>Transfer one dechorionated embryo to the cap depression minimizing carrying over BSS with the embryo. Immediately uptake the agarose and embryo from the cap depression and transfer to a Petri dish culture chamber. 
 
 A 3.5cm Petri dish with a glass window at the bottom is used. For imaging using an inverted microscope, embryos are orientated body facedown and placed close to the cover glass. For imaging using an upright microscope, the thickness of agarose is minimized. </li>
 <li>Transfer the 3.5cm Petri dish into a 14cm diameter Petri dish containing ice and water (water filled to roughly 1/3 total depth of large dish). Whilst holding the chamber down firmly on the bottom of the 14cm Petri dish, use a hair loop to gently orientate the embryo as desired in the molten agarose and hold the embryo until the agarose solidifies by gently raising and lowering the smaller dish to its original position in the ice-cold water.</li>
</ol>
4. Cell transplantation in medaka embryos
The goal of this procedure is to determine whether the gene of interest acts cell-autonomously (within a cell) or non cell-autonomously (between cells).
Donor cells can be labeled with a tracer dye such as rhodamine-dextran prior to transplantation (as outlined in the accompanying JoVE protocol 'Microinjection Of Medaka Embryos') or a transgenic strain with GFP expression may be utilized, allowing transplantation assessment. A combination of both labeling techniques is often useful to overcome auto-fluorescence that can be encountered. For time-lapse studies following transplantation, GFP expression is particularly useful.
Use a wide-mouthed glass pipette with pipette pump throughout this procedure and sterilize all tools (including slides) beforehand with 70% EtOH followed by thorough rinsing with sterile 1X BSS. Recipient embryos are usually developed to around stage 12 as this allows discrimination of the ventral and dorsal poles when carrying out transplantation.
<ol>
 <li>Commence dechorionating embryos 1.5 hours prior to transplantation and setup injection apparatus during enzyme incubations.</li>
 <li>Place a cavity microscope slide into a 9cm diameter Petri dish.</li>
 <li>Add a small amount of 3% methylcellulose to the centre of the cavity slide using a sterile pipette tip and spread thinly.</li>
 <li>Dry methylcellulose for approximately 1-2 minutes.</li>
 <li>Add 350&mu;l of sterile 1X BSS to fill slide depression.</li>
 <li>Transfer one donor embryo and up to four recipient embryos to the slide using a glass pipette. </li>
 <li>Orientate embryos using a hair loop so the blastoderm of the embryo is facing upwards. 
 
 Embryos can be leant against each other to further increase stability. 
 
 </li>
 <li>Gently insert the micro-needle into donor blastoderm and slowly uptake 10-20 cells.</li>
 <li>Gently insert needle into required area of recipient embryo blastoderm and expel the cells slowly. Great care should be taken when inserting cells into the recipient blastoderm so as not to disrupt the cell-yolk sac boundary as this will result in death of the embryo.</li>
 <li>Pour sterilized 1X BSS into the Petri dish. 
 
 Carefully pour the BSS into dish as close to the side of the dish as possible so as not to disrupt the embryos. Never pour the BSS directly onto the embryos and add sufficient BSS such that the slide and embryos are immersed. 
 
 </li>
 <li>Add 100&mu;l of penicillin/streptomycin to the dish and cover. Carefully transfer dish to a 27&deg;C incubator to allow normal development.</li>
 <li>Embryos can be periodically observed as desired. When using transplantation to carry out gain-of-function and/or phenotype rescue experiments, some morphological changes observed might be due to effects of transplantation. Thus multiple transplantations are necessary. After 2-3 days, melanophores should be present on the yolk sac, head, eyes and trunk. If present on transplanted embryos then a successful chimera has most likely been produced (Figure 3).</li>
</ol>
If necessary, genotype donor embryo(s) by transferring to PCR tube(s) containing 25&mu;l of 20mg/ml proteinase K. Incubate at 55&deg;C for 4 hours followed by 10 minutes at 94&deg;C and carry out PCR.
5. Representative results
<img src="/files/ftp_upload/2055/2055fig2.jpg" alt="Figure 2" /> 
 Figure 2. Example position of an agarose-embedded embryo. Anterior is shown to the right and view is dorsal. Panel B: endothelial cells can be seen on either side of the embryo body and are labeled using GFP driven by the fli promoter.
<img src="/files/ftp_upload/2055/2055fig3.jpg" alt="Figure 3" /> 
 Figure 3. Images of post-transplantation embryos. Image A shows a donor and recipient embryo immediately post-transplantation. The donor embryo is shown on the right with a completely red blastoderm (arrow). The recipient embryo is shown on the left and is easily identified by the small mass of red transplanted cells visible in the blastoderm (arrowhead). Image B shows two recipient embryos approximately 7 hours post-transplantation (incubated at 27&deg;C). Notice the transplanted cells have migrated from the site of transplantation and are now dispersed across the blastoderm (arrows). Image C shows a recipient embryo at st.29 (approximately 74hpf). Note the pigmentation in the eye and the presence of melanophores in the trunk and brain region (arrows). The rhodamine-labelled cells can be seen to be colonising many of the embryonic structures indicating the successful production of a chimera.

<ol>
	<li>Furutani-Seiki, M., Sasado, T., Morinaga, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+systematic+genome-wide+screen+for+mutations+affecting+organogenesis+in+Medaka,+Oryzias+latipes.">A systematic genome-wide screen for mutations affecting organogenesis in Medaka, Oryzias latipes.</a> Mech Dev. 121, 647-658 (2004).</li><li>Furutani-Seiki, M., Wittbrodt, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Medaka+and+zebrafish,+an+evolutionary+twin+study.">Medaka and zebrafish, an evolutionary twin study.</a> Mech Dev. 121, 629-6237 (2004).</li><li>Hirose, Y., Varga, Z. M., Kondoh, H., Furutani-Seiki, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single+cell+lineage+and+regionalization+of+cell+populations+during+Medaka+neurulation.">Single cell lineage and regionalization of cell populations during Medaka neurulation.</a> Development. 131, 2553-2563 (2004).</li><li>Iwamatsu, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stages+of+normal+development+in+the+medaka+Oryzias+latipes.">Stages of normal development in the medaka Oryzias latipes.</a> Zool Sci. 11, 825-839 (1994).</li><li>Cope, J., Fluck, R., Nicklas, L., Plumhoff, L. A., Sincock, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+stellate+layer+and+rhythmic+contractions+of+the+Oryzias+latipes+embryo.">The stellate layer and rhythmic contractions of the Oryzias latipes embryo.</a> J Exp Zool. 254, 270-275 (1990).</li><li>Brend, T., Holley, S. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zebrafish+Whole+Mount+High-Resolution+Double+Fluorescent+In+Situ+Hybridization.">Zebrafish Whole Mount High-Resolution Double Fluorescent In Situ Hybridization.</a> J Vis Exp. , (2009).</li></ol>

No conflicts of interest declared.

In this video we demonstrate how to dechorionate and carry out cell transplanataion on medaka embryos. This is a powerful technique to produce chimeric embryos for the study of gene/protein function during development, as well as for elucidating the autonomy of the gene/protein in question. Medaka are a useful vertebrate model system for this technique due to well established fate maps and their transparency lending them well to in vivo real-time imaging. Transplantation can be combined with microinjection (as outlined in the accompanying JoVE protocol 'Microinjection Of Medaka Embryos') so that the behaviour of transplanted cells can be easily observed.

<table><tbody><tr><td>Embryo medium</td><td>Reagent</td><td>Made in-house</td><td>N/A</td><td>200ml 50X stock solution, 1ml 1%methyleneblue in H&lt;sub&gt;2&lt;/sub&gt;O/10 liter RO water (50X stock solution: NaCl 14.7g, KCl 0.6g, CaCl&lt;sub&gt;2&lt;/sub&gt; 2H&lt;sub&gt;2&lt;/sub&gt;O 2.4g, MgSO&lt;sub&gt;4&lt;/sub&gt; 7H&lt;sub&gt;2&lt;/sub&gt;O 4.0g/1 litre RO water.)</td></tr><tr><td>Micro-dissecting forceps</td><td>Tools</td><td>55 INOX A.DUMONT&amp;amp;FILS</td><td>N/A</td><td>Very sharp fine tips for removal of chorion.</td></tr><tr><td>Fine waterproof sandpaper - p2000 grit size</td><td>Tools</td><td>Hermes Abrasives Ltd.</td><td>N/A</td><td>Very fine grit size for rolling/cleaning embryos with chorions.</td></tr><tr><td>3% methylcellulose</td><td>Reagent</td><td>Sigma</td><td>M 0512</td><td>High viscosity for holding embryos in place during transplantation.</td></tr><tr><td>1X BSS</td><td>Reagent</td><td>Made in-house</td><td>N/A</td><td>20XBSS: 130g NaCl, 8g KCl, 4g MgSO&lt;sub&gt;4&lt;/sub&gt;.7H&lt;sub&gt;2&lt;/sub&gt;O, 4g CaCl&lt;sub&gt;2&lt;/sub&gt;.2H&lt;sub&gt;2&lt;/sub&gt;O, and 10mg Phenol Red in 1L MilliQ water and autoclave; 500mM Hepes in MilliQ water autoclaved. Add 25ml 20XBSS and 15ml 500ml Hepes pH7.0, fill up to 500ml and filter sterilize before use.</td></tr><tr><td>Penicillin-streptomycin</td><td>Reagent</td><td>Gibco</td><td>15140-122</td><td>Added to BSS when incubating dechorionated embryos.</td></tr><tr><td>Pronase</td><td>Reagent</td><td>Calbiochem</td><td>53702</td><td>For dechorionation.</td></tr><tr><td>Hatching enzyme</td><td>Reagent</td><td>Made in-house</td><td>N/A</td><td>For dechorionation.</td></tr><tr><td>Tricaine mesilate (TMS)</td><td>Reagent</td><td>Sigma</td><td>A-5040</td><td>400mg tricaine in 97.9ml distilled water and 2.1ml 1M Tris, pH adjusted to 7; 4.2ml of this solution in 100ml clean tank water.</td></tr><tr><td>3% ultra-low gelling temp. agarose</td><td>Reagent</td><td>Sigma</td><td>A-2576</td><td>Type IX-A. Used during embedding of embryos for observation.</td></tr><tr><td>Petri dish culture chamber</td><td>Tool</td><td>Iwaki, Asahi Techno Glass Corp.</td><td>11-004-008</td><td>Iwaki glass base dish 35mm with 12mm window. Used during embedding of embryos for observation.</td></tr><tr><td>Micromanipulator</td><td>Equipment</td><td>Narishige</td><td>N/A</td><td>Model MN151. For removal and insertion of cells during transplantation.</td></tr><tr><td>Borosilicate glass capillaries</td><td>Tool</td><td>Harvard Apparatus</td><td>GC100-10T</td><td>For making transplantation needles.</td></tr><tr><td>Flaming/Brown micropipette puller</td><td>Equipment</td><td>Sutter Instrument Co.</td><td>Model P-97</td><td>For making transplantation needles.</td></tr></tbody></table>

dechorionation of medaka embryos and cell transplantation for the generation of chimeras

Medaka is a small egg-laying freshwater fish that allows both genetic and embryological analyses and is one of the three vertebrate model organisms in which genome-wide phenotype-driven mutant screens were carried out 1. Divergence of functional overlap of related genes between medaka and zebrafish allows identification of novel phenotypes that are unidentifiable in a single species 2, thus medaka and zebrafish are complementary for genetic dissection of the vertebrate genome functions. Manipulation of medaka embryos, such as dechorionation, mounting embryos for imaging and cell transplantation, are key procedures to work on both medaka and zebrafish in a laboratory. Cell transplantation examines cell autonomy of medaka mutations. Chimeras are generated by transplanting labeled cells from donor embryos into unlabeled recipient embryos. Donor cells can be transplanted to specific areas of the recipient embryos based on the fate maps 3 so that clones from transplanted cells can be integrated in the tissue of interest during development. Due to the hard chorion and soft embryos, manipulation of medaka embryos is more involved than in zebrafish. In this video, we show detailed procedures to manipulate medaka embryos.

Watch this Scientific Journal Video about Dechorionation of Medaka Embryos and Cell Transplantation for the Generation of Chimeras at JoVE.com

Dechorionation of Medaka Embryos and Cell Transplantation for the Generation of Chimeras

Due to the hard chorion and soft embryos, manipulation of medaka embryos is more involved than in zebrafish. This video shows step-by-step procedures for how to manipulate medaka embryos, including dechorionation, mounting in agarose for imaging and cell transplantation for the production of chimeras. These procedures are essential to use medaka and zebrafish in a laboratory to take full advantage of their complementary features for the genetic dissection of vertebrate genome functions.

Medaka is a small egg-laying freshwater fish that allows both genetic and embryological analyses and is one of the three vertebrate model organisms in ...

dechorionation-medaka-embryos-cell-transplantation-for-generation

Nanyang Technological University

Research

JoVE Journal

Biology

17.7K Views.  University of Bath. Due to the hard chorion and soft embryos, manipulation of medaka embryos is more involved than in zebrafish. This video shows step-by-step procedures for how to manipulate medaka embryos, including dechorionation, mounting in agarose for imaging and cell transplantation for the production of chimeras. These procedures are essential to use medaka and zebrafish in a laboratory to take full advantage of their complementary features for the genetic dissection of vertebrate ge...

Video: Dechorionation of Medaka Embryos and Cell Transplantation for the Generation of Chimeras

Drug Treatment and In Vivo Imaging of Osteoblast-Osteoclast Interactions in a Medaka Fish Osteoporosis Model

Bone-forming osteoblasts interact with bone-resorbing osteoclasts to coordinate the turnover of bone matrix and to control skeletal homeostasis. Medaka and zebrafish larvae are widely used to analyze the behavior of bone cells during bone formation, degeneration, and repair. Their optical clarity allows the visualization of fluorescently labeled bone cells and fluorescent dyes bound to the mineralized skeletal matrix. Our lab has generated transgenic medaka fish that express the osteoclast-inducing factor Receptor Activator of Nuclear-factor &#954;B Ligand (RANKL) under the control of a heat&#160;shock-inducible promoter. Ectopic expression of RANKL results in the excess formation of activated osteoclasts, which can be visualized in reporter lines with nlGFP expression under the control of the cathepsin K (ctsk) promoter. RANKL induction and ectopic osteoclast formation leads to severe osteoporosis-like phenotypes. Compound transgenic medaka lines that express ctsk:nlGFP in osteoclasts, as well as mCherry under the control of the osterix (osx) promoter in premature osteoblasts, can be used to study the interaction of both cell types. This facilitates the in vivo observation of cellular behavior under conditions of bone degeneration and repair. Here, we describe the use of this system to test a drug commonly used in human osteoporosis therapy and describe a protocol for live imaging. The medaka model complements studies in cell culture and mice, and offers a novel system for the in vivo analysis of drug action in the skeletal system.

Drug Treatment and In Vivo Imaging of Osteoblast-Osteoclast Interactions in a Medaka Fish Osteoporosis Model

Small laboratory fish have become popular models for bone research on the mechanisms underlying human bone disorders and for the screening of bone-modulating drugs. In this report, we describe a protocol to assess the effect of alendronate on bone cells in medaka larvae with osteoporotic lesions.

Bone-forming osteoblasts interact with bone-resorbing osteoclasts to coordinate the turnover of bone matrix and to control skeletal homeostasis. Medaka ...

Developmental Biology

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

The genetic and technical strengths have made the zebrafish vertebrate a key model organism in which the consequences of gene manipulations can be traced in vivo throughout the rapid developmental period. Multiple processes can be studied including cell proliferation, gene expression, cell migration and morphogenesis. Importantly, the generation of chimeras through transplantations can be easily performed, allowing mosaic labeling and tracking of individual cells under the influence of the host environment. For example, by combining functional gene manipulations of the host embryo (e.g., through morpholino microinjection) and live imaging, the effects of extrinsic, cell nonautonomous signals (provided by the genetically modified environment) on individual transplanted donor cells can be assessed. Here we demonstrate how this approach is used to compare the onset of fluorescent transgene expression as a proxy for the timing of cell fate determination in different genetic host environments.
In this article, we provide the protocol for microinjecting zebrafish embryos to mark donor cells and to cause gene knockdown in host embryos, a description of the transplantation technique used to generate chimeric embryos, and the protocol for preparing and running in vivo time-lapse confocal imaging of multiple embryos. In particular, performing multiposition imaging is crucial when comparing timing of events such as the onset of gene expression. This requires data collection from multiple control and experimental embryos processed simultaneously. Such an approach can easily be extended for studies of extrinsic influences in any organ or tissue of choice accessible to live imaging, provided that transplantations can be targeted easily according to established embryonic fate maps.

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

Live tracking of individual WT retinal progenitors in distinct genetic backgrounds allows for the assessment of the contribution of cell non-autonomous signaling during neurogenesis. Here, a combination of gene knockdown, chimera generation via embryo transplantation and in vivo time-lapse confocal imaging was utilized for this purpose.

The genetic and technical strengths have made the zebrafish vertebrate a key model organism in which the consequences of gene manipulations can be traced ...

Analysis of Neural Crest Migration and Differentiation by Cross-species Transplantation

Avian embryos provide a unique platform for studying many vertebrate developmental processes, due to the easy access of the embryos within the egg. Chimeric avian embryos, in which quail donor tissue is transplanted into a chick embryo in ovo, combine the power of indelible genetic labeling of cell populations with the ease of manipulation presented by the avian embryo.
Quail-chick chimeras are a classical tool for tracing migratory neural crest cells (NCCs)1-3. NCCs are a transient migratory population of cells in the embryo, which originate in the dorsal region of the developing neural tube4. They undergo an epithelial to mesenchymal transition and subsequently migrate to other regions of the embryo, where they differentiate into various cell types including cartilage5-13, melanocytes11,14-20, neurons and glia21-32. NCCs are multipotent, and their ultimate fate is influenced by 1) the region of the neural tube in which they originate along the rostro-caudal axis of the embryo11,33-37, 2) signals from neighboring cells as they migrate38-44, and 3) the microenvironment of their ultimate destination within the embryo45,46. Tracing these cells from their point of origin at the neural tube, to their final position and fate within the embryo, provides important insight into the developmental processes that regulate patterning and organogenesis.
Transplantation of complementary regions of donor neural tube (homotopic grafting) or different regions of donor neural tube (heterotopic grafting) can reveal differences in pre-specification of NCCs along the rostro-caudal axis2,47. This technique can be further adapted to transplant a unilateral compartment of the neural tube, such that one side is derived from donor tissue, and the contralateral side remains unperturbed in the host embryo, yielding an internal control within the same sample2,47. It can also be adapted for transplantation of brain segments in later embryos, after HH10, when the anterior neural tube has closed47.
Here we report techniques for generating quail-chick chimeras via neural tube transplantation, which allow for tracing of migratory NCCs derived from a discrete segment of the neural tube. Species-specific labeling of the donor-derived cells with the quail-specific QCPN antibody48-56 allows the researcher to distinguish donor and host cells at the experimental end point. This technique is straightforward, inexpensive, and has many applications, including fate-mapping, cell lineage tracing, and identifying pre-patterning events along the rostro-caudal axis45. Because of the ease of access to the avian embryo, the quail-chick graft technique may be combined with other manipulations, including but not limited to lens ablation40, injection of inhibitory molecules57,58, or genetic manipulation via electroporation of expression plasmids59-61, to identify the response of particular migratory streams of NCCs to perturbations in the embryo's developmental program. Furthermore, this grafting technique may also be used to generate other interspecific chimeric embryos such as quail-duck chimeras to study NCC contribution to craniofacial morphogenesis, or mouse-chick chimeras to combine the power of mouse genetics with the ease of manipulation of the avian embryo.62

An approach for analyzing migration and eventual fate of avian neural crest cells in quail-chick chimeric embryos is described. This method is a simple and straightforward technique for tracing neural crest cells during migration and differentiation that are otherwise difficult to distinguish within an unmanipulated chick embryo.

Avian embryos provide a unique platform for studying many vertebrate developmental processes, due to the easy access of the embryos within the egg. ...

Neuroscience

Generation of Chimeric Axolotls with Mutant Haploid Limbs Through Embryonic Grafting

A growing set of genetic techniques and resources enable researchers to probe the molecular origins of the ability of some species of salamanders, such as axolotls, to regenerate entire limbs as adults. Here, we outline techniques used to generate chimeric axolotls with Cas9-mutagenized haploid forelimbs that can be used for exploring gene function and the fidelity of limb regeneration. We combine several embryological and genetic techniques, including haploid generation via in vitro activation, CRISPR/Cas9 mutagenesis, and tissue grafting into one protocol to produce a unique system for haploid genetic screening in a model organism of regeneration. This strategy reduces the number of animals, space, and time required for the functional analysis of genes in limb regeneration. This also permits the investigation of regeneration-specific functions of genes that may be required for other essential processes, such as organogenesis, tissue morphogenesis, and other essential embryonic processes. The method described here is a unique platform for conducting haploid genetic screening in a vertebrate model system.

This goal of this protocol is to produce chimeric axolotls with haploid forelimbs derived from Cas9-mutagenized donor tissue using embryonic tissue grafting techniques.

A growing set of genetic techniques and resources enable researchers to probe the molecular origins of the ability of some species of salamanders, such as ...

Genetics

Generating Chimeric Zebrafish Embryos by Transplantation

One of the most powerful tools used to gain insight into complex developmental processes is the analysis of chimeric embryos. A chimera is defined as an organism that contains cells from more than one animal; mosaics are one type of chimera in which cells from more than one genotype are mixed, usually wild-type and mutant. In the zebrafish, chimeras can be readily made by transplantation of cells from a donor embryo into a host embryo at the appropriate embryonic stage. Labeled donor cells are generated by injection of a lineage marker, such as a fluorescent dye, into the one-cell stage embryo. Labeled donor cells are removed from donor embryos and introduced into unlabeled host embryos using an oil-controlled glass pipette mounted on either a compound or dissecting microscope. Donor cells can in some cases be targeted to a specific region or tissue of the developing blastula or gastrula stage host embryo by choosing a transplantation site in the host embryo based on well-established fate maps.

A step-by-step guide to generating targeted chimeric zebrafish embryos by transplantation at the blastula or gastrula stage.

One of the most powerful tools used to gain insight into complex developmental processes is the analysis of chimeric embryos. A chimera is defined as an ...

Tissue Targeted Embryonic Chimeras: Zebrafish Gastrula Cell Transplantation

Certain fundamental questions in the field of developmental biology can only be answered when cells are placed in novel environments or when small groups of cells in a larger context are altered.  Watching how one cell interacts with and behaves in a unique environment is essential to characterizing cell functions.  Determining how the localized misexpression of a specific protein influences surrounding cells provides insightful information on the roles that protein plays in a variety of developmental processes.  Our lab uses the zebrafish model system to uniquely combine genetic approaches with classical transplantation techniques to generate genotypic or phenotypic chimeras.  We study neuron-glial cell interactions during the formation of forebrain commissures in zebrafish. This video describes a method that allows our lab to investigate the role of astroglial populations in the diencephalon and the roles of specific guidance cues that influence projecting axons as they cross the midline. Due to their transparency zebrafish embryos are ideal models for this type of ectopic cell placement or localized gene misexpression. Tracking transplanted cells can be accomplished using a vital dye or a transgenic fish line expressing a fluorescent protein. We demonstrate here how to prepare donor embryos with a vital dye tracer for transplantation, as well as how to extract and transplant cells from one gastrula staged embryo to another.  We present data showing ectopic GFP+ transgenic cells within the forebrain of zebrafish embryos and characterize the location of these cells with respect to forebrain commissures.  In addition, we show laser scanning confocal timelapse microscopy of Alexa 594 labeled cells transplanted into a GFP+ transgenic host embryo.  These data provide evidence that gastrula staged transplantation enables the targeted positioning of ectopic cells to address a variety of questions in Developmental Biology.

Zebrafish cell transplantation enables the combination of genetics and embryology to generate tissue specific chimeras. This video demonstrates gastrula staged cell transplantations that have allowed our lab to investigate the roles of astroglial populations and specific guidance cues during commissure formation in the forebrain.

Certain fundamental questions in the field of developmental biology can only be answered when cells are placed in novel environments or when small groups ...

Transplantation of GFP-expressing Blastomeres for Live Imaging of Retinal and Brain Development in Chimeric Zebrafish Embryos

Cells change extensively in their locations and property during embryogenesis. These changes are regulated by the interactions between the cells and their environment. Chimeric embryos, which are composed of cells of different genetic background, are great tools to study the cell-cell interactions mediated by genes of interest. The embryonic transparency of zebrafish at early developmental stages permits direct visualization of the morphogenesis of tissues and organs at the cellular level. Here, we demonstrate a protocol to generate chimeric retinas and brains in zebrafish embryos and to perform live imaging of the donor cells. The protocol covers the preparation of transplantation needles, the transplantation of GFP-expressing donor blastomeres to GFP-negative hosts, and the examination of donor cell behavior under live confocal microscopy. With slight modifications, this protocol can also be used to study the embryonic development of other tissues and organs in zebrafish. The advantages of using GFP to label donor cells are also discussed. 

We demonstrate a protocol to generate chimeric zebrafish embryos for live imaging cellular behavior during embryogenesis.

Cells change extensively in their locations and property during embryogenesis. These changes are regulated by the interactions between the cells and their ...

Microinjection of Medaka Embryos for use as a Model Genetic Organism

In this video, we demonstrate the technique of microinjection into one-cell stage medaka embryos. Medaka is a small egg-laying freshwater fish that allows both genetic and embryological analyses and is one of the vertebrate model organisms in which genome-wide phenotype-driven mutant screens were carried out 1, as in zebrafish and the mouse. Divergence of functional overlap of related genes between medaka and zebrafish allows identification of novel phenotypes that are unidentifiable in a single species 2, thus medaka and zebrafish are complementary for genetic dissection of vertebrate genome functions.
To take advantage of medaka fish whose embryos are transparent and develop externally, microinjection is an essential technique to inject cell-tracers for labeling cells, mRNAs or anti-sense oligonucleotides for over-expressing and knocking-down genes of interest, and DNAs for making transgenic lines.

Medaka and zebrafish are complementary for genetic dissection of vertebrate genome functions. This protocol highlights the key points for successful microinjection into medaka embryos, an important technique for embryological and genetic analysis using medaka and zebrafish in a laboratory.

In this video, we demonstrate the technique of microinjection into one-cell stage medaka embryos. Medaka is a small egg-laying freshwater fish that allows ...

A Simple and Effective Transplantation Device for Zebrafish Embryos

Classical embryological manipulations, such as removing cells and transplanting cells within or between embryos, are powerful techniques to study complex developmental processes. Zebrafish embryos are ideally suited for these manipulations since they are easily accessible, relatively large in size, and transparent. However, previously developed devices for cell removal and transplantation are cumbersome to use or expensive to purchase. In contrast, the transplantation device presented here is economical, easy to assemble, and simple to use. In this protocol, we first introduce the handling of the transplantation device as well as its assembly from commercially and widely available parts. We then present three applications for its use: generation of ectopic clones to study signal dispersal from localized sources, extirpation of cells to produce size-reduced embryos, and germline transplantation to generate maternal-zygotic mutants. Finally, we show that the tool can also be used for embryological manipulations in other species such as the Japanese rice fish medaka.

Embryological manipulations such as extirpation and transplantation of cells are important tools to study early development. This protocol describes a simple and effective transplantation device to perform these manipulations in zebrafish embryos.

Classical embryological manipulations, such as removing cells and transplanting cells within or between embryos, are powerful techniques to study complex ...

Assessing Species-specific Contributions To Craniofacial Development Using Quail-duck Chimeras

The generation of chimeric embryos is a widespread and powerful approach to study cell fates, tissue interactions, and species-specific contributions to the histological and morphological development of vertebrate embryos. In particular, the use of chimeric embryos has established the importance of neural crest in directing the species-specific morphology of the craniofacial complex. The method described herein utilizes two avian species, duck and quail, with remarkably different craniofacial morphology. This method greatly facilitates the investigation of molecular and cellular regulation of species-specific pattern in the craniofacial complex.&#160;Experiments in quail and duck chimeric embryos have already revealed neural crest-mediated tissue interactions and cell-autonomous behaviors that regulate species-specific pattern in the craniofacial skeleton, musculature, and integument. The great diversity of neural crest derivatives suggests significant potential for future applications of the quail-duck chimeric system to understanding vertebrate development, disease, and evolution.

This article describes a method to generate chimeric embryos that are&#160;designed to test the species-specific contributions of neural crest and/or other tissues to craniofacial development.

The generation of chimeric embryos is a widespread and powerful approach to study cell fates, tissue interactions, and species-specific contributions to ...

Dechorionation of Medaka Embryos and Cell Transplantation for the Generation of Chimeras

Summary

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Drug Treatment and In Vivo Imaging of Osteoblast-Osteoclast Interactions in a Medaka Fish Osteoporosis Model

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

Analysis of Neural Crest Migration and Differentiation by Cross-species Transplantation

Generation of Chimeric Axolotls with Mutant Haploid Limbs Through Embryonic Grafting

Generating Chimeric Zebrafish Embryos by Transplantation

Tissue Targeted Embryonic Chimeras: Zebrafish Gastrula Cell Transplantation

Transplantation of GFP-expressing Blastomeres for Live Imaging of Retinal and Brain Development in Chimeric Zebrafish Embryos

Microinjection of Medaka Embryos for use as a Model Genetic Organism

A Simple and Effective Transplantation Device for Zebrafish Embryos

Assessing Species-specific Contributions To Craniofacial Development Using Quail-duck Chimeras