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.

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<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 H2O/10 liter RO water (50X stock solution: NaCl 14.7g, KCl 0.6g, CaCl2 2H2O 2.4g, MgSO4 7H2O 4.0g/1 litre RO water.)</td>
</tr>
<tr>
<td>Micro-dissecting forceps</td>
<td>Tools</td>
<td>55 INOX A.DUMONT&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 MgSO4.7H2O, 4g CaCl2.2H2O, 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>
		</div>

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

Research

JoVE Journal

Biology

17.6K Views.  University of Bath. Из-за жестких и мягких хориона эмбрионов, манипуляции с эмбрионами оризии более сложный, чем в данио. Это видео показывает, шаг за шагом процедуры, как манипулировать оризии эмбрионов, в том числе dechorionation, монтаж в агарозном для работы с изображениями и клеточн...

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

Isolation and Transplantation of Hematopoietic Stem Cells (HSCs)

Isolation and Transplantation of Hematopoietic Stem Cells HSCs

Dissection of 6.5 dpc Mouse Embryos

Analysis of gene expression patterns during early stages of mammalian embryonic development can provide important clues about gene function, cell-cell interaction and signaling mechanisms that guide embryonic patterning. However, dissection of the mouse embryo from the decidua shortly after implantation can be a challenging procedure, and detailed step-by-step documentation of this process is lacking.

Here we demonstrate how post-implantation (6.5 dpc) embryos are isolated by first dissecting the uterus of a pregnant mouse (detection of the vaginal plug was designated day 0.5 poist coitum) and subsequently dissecting the embryo from maternal decidua. The dissection of Reichert's membrane is described as well as the removal of the ectoplacental cone.

Isolation of postimplantation-stage embryos allows one to study gene patterning and analyze cell-lineage decision making processes during embryonic development, but proper dissection of the early embryo can be challenging. This protocol describes a method for isolating early primitive-streak-stage embryos (~6.5 days post coitum [dpc]).

Analysis of gene expression patterns during early stages of mammalian embryonic development can provide important clues about gene function, cell-cell ...

Tracheotomy: A Method for Transplantation of Stem Cells to the Lung

Lung disease is a leading cause of death and likely to become an epidemic given increases in pollution and smoking worldwide. Advances in stem cell therapy may alleviate many of the symptoms associated with lung disease and induce alveolar repair in adults. Concurrent with the ongoing search for stem cells applicable for human treatment, precise delivery and homing (to the site of disease) must be reassured for successful therapy. Here, I report that stem cells can safely be instilled via the trachea opening a non-stop route to the lung. This method involves a skin incision, caudal insertion of a cannula into and along the tracheal lumen, and injection of a stem cell vehicle mixture into airways of the lung. A broad range of media solutions and stabilizers can be instilled via tracheotomy, resulting in the ability to deliver a wider range of cell types. With alveolar epithelium confining these cells to the lumen, lung expansion and negative pressure during inhalation may also assist in stem cell integration. Tracheal delivery of stem cells, with a quick uptake and the ability to handle a large range of treatments, could accelerate the development of cell-based therapies, opening new avenues for treatment of lung disease.

Significant breakthroughs in stem cell identification are continuously being made. To translate these discoveries, however, novel methods for cellular delivery must be devised. Here I report that the airways provide a safe route for stem cell transplantation to the lungs.

Lung disease is a leading cause of death and likely to become an epidemic given increases in pollution and smoking worldwide. Advances in stem cell ...

Plastic Embedding and Sectioning of Xenopus laevis Embryos

Plastic sections maintain true tissue morphology in thin sections of tissue that can be immunostained with fluorescent secondary antibodies, making this method more useful than paraffin-embedded or frozen sections for many types of tissue. The method for staining, plastic embedding, and sectioning is demonstrated in this video.

Generation of Human CD40-activated B cells

CD40-activated B cells (CD40-B cells) have been identified as an alternative source of immuno-stimulatory antigen-presenting cells (APC) for cancer immunotherapy 1-3. Compared to Dendritic cells (DCs), the best characterized APC, CD40-B cells have several distinct biological and technical properties. Similar to DCs, B cells show an increased expression of MHC and co-stimulatory molecules (Fig.1b), exhibit a strong migratory capacity and present antigen presentation efficiently to T cells, after stimulation with interleukin-4 and CD40 ligand (CD40L). However, in contrast to immature or mature DCs, CD40-B cells express the full lymph node homing triad consisting of CD62L, CCR7/CXCR4, and leukocyte function antigen-1 (LFA1, CD11a/CD18), necessary for homing to secondary lymphoid organs (Fig.1a) 3. CD40-B cells can be generated without difficulties from very small amounts of peripheral blood which can be further expanded in vitro to very large amounts of highly-pure CD40-B cells (&gt;109 cells per patient) from healthy donors as well as cancer patients (Fig.1c,d) 1,4. 
In this protocol we demonstrate how to obtain fully activated CD40-B cells from human PBMC. Key molecules for the cell culture are CD40 ligand, interleukin-4 (IL-4) and cyclosporin A (CsA), which are replenished in a 3-4 day culture cycle. For laboratory purposes CD40-stimulation is provided by NIH/3T3 cells expressing recombinant human CD40 ligand (tCD40L NIH/3T3) 5. To avoid contamination with non-transfected cells, expression of the human CD40 ligand on the transfectants has to be checked regularly (Fig.2). 
After 14 days CD40-B cell cultures consist of more than 95% pure B cells and an expansion of CD40-B cells over 65 days is frequently possible without any loss of function 1, 4. CD40-B cells efficiently take up, process and present antigens to T cells 6. They do not only prime na&#970;ve, but also expand memory T cells 7,8. CD40-activated B cells can be used to study B-cell activation, differentiation and function. Moreover, they represent a promising tool for therapeutic or preventive vaccination against tumors 9.

In this video we present the ex vivo generation and expansion of human CD40-activated B cells (CD40-B) from peripheral blood mononuclear cells (PBMC) by stimulation with CD40 ligand and interleukin-4.

CD40-activated B cells (CD40-B cells) have been identified as an alternative source of immuno-stimulatory antigen-presenting cells (APC) for cancer ...

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 ...

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

The morphogenesis of the Drosophila embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell shape changes during embryonic development. One of the challenges faced in studying this structure is that the lumen of the heart tube, as well as the membrane features that are crucial to heart tube formation, are difficult to visualize in whole mount embryos, due to the small size of the heart tube and intra-lumenal space relative to the embryo. The use of transmission electron microscopy allows for higher magnification of these structures and gives the advantage of examining the embryos in cross section, which easily reveals the size and shape of the lumen. In this video, we detail the process for reliable fixation, embedding, and sectioning of late stage Drosophila embryos in order to visualize the heart tube lumen as well as important cellular structures including cell-cell junctions and the basement membrane.

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

We describe a process for fixation, embedding, sectioning, and imaging of late stage Drosophila embryos for Trasmission Electron Microscopy of the embryonic heart tube. This technique allows for the visualization of the heart tube lumen as well as the basement membrane, which lines the lumen of the heart.

The morphogenesis of the Drosophila  embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell ...

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 ...