The author thanks Hugo Juraver-Geslin, Marion Wassef and Anne H&#233;l&#232;ne Monsoro-Burq for their help and advice, and the Animal Facility of the Institut Curie. The author thanks Paul Johnson for his editing work on the manuscript. This work was supported by the Centre National de la Recherche Scientifique (CNRS UMR8197, INSERM U1024) and by grants from the "Association pour la Recherche sur le Cancer" (ARC 4972 and ARC 5115; FRC DOC20120605233 and LABEX Memolife) and the Fondation Pierre Gilles de Gennes (FPGG0039).

Experiments comply with National and European regulation on the protection of animals used for scientific purposes and with internationally established principles of replacement, reduction and refinement.
1. Flat-mounting of Xenopus laevis Tadpoles Anterior Neural Tube After Whole-mount In Situ Hybridization
<ol>
	<li>Obtain X. laevis embryos according to standard procedures4 and age them until they reach neurula stage 26 and older (according to Nieuwkoop and Faber developmental table17.</li>
	<li>Fix X. laevis tadpoles by incubating them in a solution of 4% paraformaldehyde (PFA). Rock and rotate the embryos in a 2 ml glass vial , 1 to 1.5 hr at RT&#160;for stage 26 to stage 38 embryos, 2 hr at RT for embryos at later stages. 
	CAUTION: PFA is toxic by contact and a suspected carcinogen. It should be manipulated under a hood. 
	NOTE: One X. laevis brood is usually fixed in a 2 ml glass vial but a larger glass vial can be used.</li>
	<li>Wash the embryos in the same vial using 1x PBS with 0.1% Tween (PBT), 3 min at RT, twice. 
	NOTE: Unless otherwise specify the PBS used is with or without calcium and magnesium.</li>
	<li>Dehydrate the embryos in 100% methanol (MeOH) for at least 12 hr at RT in the same vial. Change the MeOH 100% at least twice under the hood using a plastic micropipette. 
	NOTE: The MeOH turns slightly yellow as lipids dissolved in it. Caution the MeOH is toxic by inhalation and should be manipulated under a hood.</li>
	<li>Rehydrate the embryos using the same vial through a graded series of MeOH baths: MeOH 75% in PBS, MeOH 50% in PBS, MeOH 25% in PBS, PBS twice, each bath 5-10 min at RT. The MeOH solutions are changed under the hood using a plastic micropipette.</li>
	<li>Using a plastic pipette, transfer the embryo to be dissected in a 60 mm Petri dish filled to the top with PBT. Using two fine forceps carefully remove the eyes by inserting one fine forceps between the eyes and the neural tube, at the level of the optic stalk. Detach the eyes from the neural tube. Carefully introduce the forceps below the ectoderm overlying the neural tube. With care, peel off the ectoderm starting from behind the head and discard it.</li>
	<li>Perform a double or single ISH using digoxigenin-labeled or fluorescein-labeled probes as previously described18,19.</li>
	<li>After the ISH transfer the embryos in a new 60 mm Petri dish filled to the top with PBT using a plastic pipette.</li>
	<li>Using fine forceps carefully detach the neural tube from the rest of the embryo. At this stage, the anterior neural tube separates from the posterior neural tube. Carefully detach remaining parts of the ectoderm overlying the neural tube, the notochord that is loosely attached below the neural tube, the otic vesicles and remaining parts of the mesoderm. Ensure that the neural tube is devoid of any appendices at this stage. 
	NOTE: To avoid damage to the neural tube perform all neural tube/embryo transfer with a plastic transfer pipette or a micropipette if necessary with a tip cut at its end. The diameter of the tip end is adjusted to the size of the neural tube.</li>
	<li>Transfer the dissected neural tube (see note step 1.9) in a 1.5 ml tube filled with 50% glycerol diluted in PBS. Wait until the neural tubes have fallen to the bottom of the glycerol solution, usually O/N at 4 &#176;C.</li>
	<li>Remove the solution of 50% glycerol/PBS using a micropipette P1000 and add in the same 1.5 ml tube a solution of 90% glycerol/PBS using a micropipette P1000. Wait until the neural tubes fall to the bottom of the 90% glycerol/PBS solution, usually O/N at 4 &#176;C. 
	NOTE: for long term storage, use 90% glycerol/PBS plus antibiotics to prevent bacterial development.</li>
	<li>Transfer the neural tubes on a glass plate, or a microscope slide with a plastic transfer pipette.</li>
	<li>Dissect the neural tubes along the dorsal and ventral midlines using tungsten needles. During that step the neural tubes are kept in 90% glycerol/PBS.</li>
	<li>Mount the two sides of the neural tube in 90% glycerol/PBS using reinforcement rings covered with a glass coverslip.</li>
	<li>Fix the coverslips with varnish and keep at 4 &#176;C.</li>
</ol>
2. Animal Cap Explants Induction Using Anion Exchange Resin Beads
<ol>
	<li>Transfer 100 &#181;l of anion exchange resin beads into a 2 ml tube using a micropipette P1000. Wash the beads at least five consecutive times with sterile distilled water. Allow the beads to sediment at the bottom of the tube and replace the sterile distilled water using a micropipette P1000. Do not touch the beads.</li>
	<li>Let the beads soak O/N in a 2 ml tube filled with sterile distilled water with Bovine Serum Albumin (BSA) (10 mg/ ml) at 4 &#176;C.</li>
	<li>Two hr before collecting the ACs, place half of the beads into a new 1.5 ml tube using a micropipette P1000. Replace the sterile distilled water with 500 &#181;l of the medium containing the molecule to be tested for its inductive potential, or known to induce a specific fate. Keep the other half of the beads, as they will be used as a negative control.</li>
	<li>Incubate the beads at 4 &#176;C for at least 2 hr.</li>
	<li>Carry out all subsequent steps under the stereomicroscope and perform all AC transfer with a micropipette.</li>
	<li>Transfer blastula or very early gastrula embryos into PBS with calcium and magnesium complemented with 0.2% BSA using a plastic transfer pipette. 
	NOTE: Embryos developing at 12 &#176;C reach blastula and gastrula stages in 20 to 24 hr.</li>
	<li>Remove the embryo&#39;s vitelline membrane using forceps as previously described20. Fix the embryo with one forceps. Pinch the vitelline membrane with the side of the other forceps. Holding the membrane, slowly peel it off the embryo. Perform this procedure on the ventral (vegetal) side of the embryo. Do not damage the animal side of the embryo.</li>
	<li>Isolate small ACs as previously described21. The animal pole is the embryo&#39;s pigmented part. Using fine forceps cut out a small square of tissue out of the animal pole. The AC tissue contains only the ectodermal cells. Ensure that the tissue is of uniform thickness. If not, remove the AC from the analysis.</li>
	<li>Place each AC in a Terasaki multiwell plates in 0.5x Modified Barth&#39;s Saline (MBS)4. Place the AC into the well with its pigmented animal side down, in contact with the round bottom of the well. 
	NOTE: coating pipettes with a 0.1% BSA solution prevents the adhesion of small pieces of adhesive tissue to the plastic.</li>
	<li>Place one bead on each cap using a micropipette P20. If needed, use forceps to carefully place the bead in the center of the cap. 
	NOTE: When soaked in a conditioned medium, the beads are colored in pink. Select the most colored beads.</li>
	<li>Incubate at RT (18 to 22 &#176;C) for 6 hr without moving the Terasaki multiwell plate. The bead sticks to the AC in less than 2 hr.</li>
	<li>Place the Terasaki multiwell plate on top of papers soaked in water. Cover the plate with a plastic container. 
	NOTE: This &#39;humidity chamber&#39; prevents premature evaporation of the wells.</li>
	<li>Culture the ACs in 0.5x MBS or 3/4 NAM (see step 4.1) in the humidity chamber at 15-20 &#176;C until the sibling embryos have reached the correct stage to test the effect of your factor.</li>
	<li>Fix the ACs for 1 hr in freshly prepared 4% PFA. Dehydrate the ACs in 100% MeOH as previously described (Step 1.4).</li>
</ol>
3. Animal Cap Cell Dissociation and Reaggregation Before Grafting in a Xenopuslaevis Neurula
<ol>
	<li>Coat petri dishes (60 mm) or 12 wells plates with 3% agarose in sterile water or in PBS without calcium and magnesium. Add enough agarose to cover the bottom of the petri dish or the well. Pre-warm the plates at RT (18-22 &#176;C). Optionally, stored the plates for couple of days at 4 &#176;C to prevent dehydration.</li>
	<li>Prepare Calcium-free Holtfreter&#39;s saline (60 mM NaCl, 0.7 mM KCl, 4.6 mM HEPES, 0.1% BSA (A-7888 Sigma pH 7.6)) and Holtfreter&#39;s saline (60 mM NaCl, 0.7 mM KCl, 0.9 mM CaCl2, 4.6 mM HEPES, 0.1% BSA pH 7.6)5. NOTE: the solution of CaCl2 cannot be autoclaved.</li>
	<li>Fill the agarose-coated well to the top with Calcium-free Holtfreter&#39;s saline.</li>
	<li>Prepare blastula or very early gastrula embryos to isolate their ACs as previously described (steps 2.5 to 2.7).</li>
	<li>Isolate at least 15 or up to 30, small ACs21. Using fine forceps cut out a small square of tissue out of the animal pole. The AC tissue only contains ectodermal cells and is therefore of uniform thickness. If not, remove the AC from the analysis. 
	NOTE: The animal pole is the embryo&#39;s pigmented part.</li>
	<li>Transfer the ACs into an agarose-coated well filled to the top with Calcium-free Holtfreter&#39;s saline. Place the ACs with their pigmented side facing upwards. 
	NOTE: The dissociation process starts rapidly in calcium-free Holtfreter&#39;s saline.</li>
	<li>Wait a few minutes for cells to start dissociating. Observe this through disaggregation of the tissues. Using fine forceps, separate the pigmented layer from the rest of the AC and discard them with a micropipette P20. Complete cell dissociation process indicates separation of cells from one another. 
	NOTE: One or two pigmented layers can be left within the re-aggregated explant to help visualization during grafting.</li>
	<li>Center the cells using circular movements of the plate. Using a micropipette P1000 carefully remove as much medium as possible. Be careful not to touch the cells.</li>
	<li>Add 1 ml of Holtfreter&#39;s saline with calcium to the well. Transfer the dissociated ACs into a 1.5 ml tube. 
	NOTE: 2 ml tubes cannot be used due to their round bottom.</li>
	<li>Pellet the cells by centrifugation, 5 min at a maximum speed of 2,000 rpm for a bench centrifuge (500 x g). Remove carefully the supernatant with a micropipette P1000.</li>
	<li>Add to the dissociated cells 20 &#181;l of Holtfreter&#39;s saline with calcium (60 mM NaCl, 0.7 mM KCl, 0.9 mM CaCl2, 4.6 mM HEPES, 0.1% BSA (A-7888 Sigma pH 7.6)5.</li>
	<li>Keep the dissociated ACs, 3 to 6 hr at RT (18-22 &#176;C). 
	NOTE: This is necessary time for the cells to re-aggregate. The re-aggregation is visible as the cells form a small ball at the bottom of the tube.</li>
	<li>Detach the explant carefully from the bottom of the tube by adding 1 ml of 0.5x MBS or of 1 ml of 3/4 NAM (see step 4.1). Transfer the explant in an un-coated plate using a plastic transfer pipette.</li>
	<li>Stage the explants using their control siblings. For long-term culture use antibiotics: kanamycin (50 &#181;g/ml), ampicillin (50 &#181;g/ml) and gentamycine (50 &#181;g/ml).</li>
</ol>
4. Grafting of Animal Caps Explants in the Neural Plate of X. laevis Embryo
<ol>
	<li>Prepare 3/4 NAM (110 mM NaCl, 2 mM KCl, 1 mM Ca(NO3)2, 1 mM MgSO4, 0.1 mM EDTA, 1 mM NaHCO3, 0.2x PBS, 50 &#181;g/ml gentamycin). Do not keep for more than 1 week for dissection and store at 4 &#176;C. Older NAM can be used for preparing agarose-coated dissection dishes (below).</li>
	<li>Coat petri dishes (60 mm) with 3% agarose in 3/4 NAM into which small holes have been made using either a silicone mold, or the cover of a table tennis racket. 
	NOTE: The agarose covers the bottom of the petri dish. Leave some space so that the petri dish to fill with 3/4 NAM. Alternatively make dissection dishes with non-drying modeling clay. Within the clay, squeeze the embryos slightly during the dissection procedure. Dig small holes in the agarose using rounded forceps This &#39;dissection dish&#39; helps to hold the embryos during the dissection procedure.</li>
	<li>Collect dissection tools: plastic transfer pipettes, an &#39;eyebrow knife&#39; (made with eyebrow hair embedded in paraffin at the tip of a glass Pasteur pipette)22, two fine dissection forceps, P1000 micropipette and tips, a stereosmicroscope with magnification 8-40X, a bright light source with optic fiber guides. Keep all the dissecting tools clean; after each experiment, rinse them twice with distilled water, once with 100% ethanol and let dry. Store away from dust and if necessary autoclave the metal dissecting tools.</li>
	<li>Let the dissociated and re-aggregated ACs (protocol 3), and their siblings X. laevis embryos develop until they reach stage 13 (neurula) according to Nieuwkoop and Faber developmental table17. For transplantation within the neural plate stage 13 to stage 15 embryos are used.</li>
	<li>Carry out all subsequent steps under the stereomicroscope.</li>
	<li>Using a plastic transfer pipette, transfer the embryos into PBS with calcium and magnesium complemented with 0.2% BSA. Remove the embryo&#39;s vitelline membrane as previously described20. Fix the embryo with one forceps. Pinch the vitelline membrane with the side of the other forceps. Holding the membrane firmly, slowly peel it off the embryo. 
	NOTE: Do this procedure on the ventral (vegetal) side of the embryo. Do not damage the embryos neural plate. If embryos have been damaged during the vitelline membrane removal step, transfer the embryos into a 60 mm petri dish containing 3/4 NAM using a plastic transfer pipette and wait 15 min, a sufficient time for the healing process to occur.</li>
	<li>Fill in the dissection dish to the top with fresh 3/4 NAM.</li>
	<li>Using a plastic transfer pipette, transfer the embryos without their vitelline membrane into the dissection dish. Place the embryos dorsal side up into the wells. During the transfer process do not allow the embryo to enter in contact with the air-liquid interface since X. laevis are lysed by surface tension.</li>
	<li>Transfer the AC explant to be grafted into the dissection plate using a plastic transfer pipette or a micropipette P1000. Once the pipette is inside the liquid, allow the explants to slowly sink down by gravity, or push very gently.</li>
	<li>Maintain the embryos with rounded forceps. With the eyebrow knife make an incision into the neural plate where you intend to graft your explant&#39;s piece. 
	NOTE: At stage 14 the anterior bending of the neural plate can be use as a landmark, it marks the diencephalon territory (Figure 4).</li>
	<li>Cut out a small piece of neuroepithelium, using rounded forceps and an eyebrow knive. Cut a small piece of the explant with the eyebrow knife. 
	NOTE: The piece of explant should be about the same size and shape as the neuroepithelium ablated area.</li>
	<li>Place the piece of explant into the neuroepithelium incision using the eyebrow knife and fine forceps. Ensure that the explant rapidly attaches to the embryo.</li>
	<li>Alternatively place a piece of glass coverslip onto the grafted embryo to maintain the graft in place. To do this, cut a fine glass coverslip into very small pieces using coarse forceps. Ensure that the size is approximately 1.5 mm2. Immerse the pieces into a Petri dish containing 3/4 NAM or PBS using forceps. Choose a piece of glass bigger than the embryo to avoid damaging it. The embryo will be a little bit flattened.</li>
	<li>Wait for at least 30 min without moving, or only move the dissecting plate gently. Let the embryo recover for 30 min to 2 hr and gently remove the coverslip if necessary. Carefully pipette the grafted embryos into a clean dish filled with 3/4 NAM, using the plastic transfer pipette. 
	NOTE: Grafted embryos tend to develop bacteria or fungi contamination. For long-term culture use antibiotics: kanamycin (50 &#181;g/ml), ampicillin (50 &#181;g/ml) and gentamycine (50 &#181;g/ml).</li>
</ol>

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<li>Puelles, L., Rubenstein, J. L. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Forebrain+gene+expression+domains+and+the+evolving+prosomeric+model%5BTitle%5D)+AND+%22Trends+in+Neurosciences%22%5BJournal%5D)">Forebrain gene expression domains and the evolving prosomeric model.</a> Trends in Neurosciences. 26, 469-476 (2003).</li>
<li>Martinez-Ferre, A., Martinez, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Molecular+regionalization+of+the+diencephalon%5BTitle%5D)+AND+%22Frontiers+In+Neuroscience%22%5BJournal%5D)">Molecular regionalization of the diencephalon.</a> Frontiers In Neuroscience. 6, 73-73 (2012).</li>
<li>Scholpp, S., Lumsden, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Review%3A+Building+a+bridal+chamber%3A+development+of+the+thalamus%5BTitle%5D)+AND+%22Trends+in+Neurosciences%22%5BJournal%5D)">Review: Building a bridal chamber: development of the thalamus.</a> Trends in Neurosciences. 33, 373-380 (2010).</li>
<li>Coffman, C., Harris, W., Kintner, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Xotch%2C+the+Xenopus+homolog+of+Drosophila+notch%5BTitle%5D)+AND+%22Science%22%5BJournal%5D)">Xotch, the Xenopus homolog of Drosophila notch.</a> Science. 249, 1438-1441 (1990).</li>
<li>Pera, E. M., Acosta, H., Gouignard, N., Climent, M., Arregi, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Active+signals%2C+gradient+formation+and+regional+specificity+in+neural+induction%5BTitle%5D)+AND+%22Exp+Cell+Res%22%5BJournal%5D)">Active signals, gradient formation and regional specificity in neural induction.</a> Exp Cell Res. 321, 25-31 (2014).</li>
<li>Stern, C. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Neural+induction%3A+old+problem%2C+new+findings%2C+yet+more+questions%5BTitle%5D)+AND+%22Development%22%5BJournal%5D)">Neural induction: old problem, new findings, yet more questions.</a> Development. 132, 2007-2021 (2005).</li>
<li>Juraver-Geslin, H. A., Ausseil, J. J., Wassef, M., Durand, B. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Barhl2+limits+growth+of+the+diencephalic+primordium+through+Caspase3+inhibition+of+beta-catenin+activation%5BTitle%5D)+AND+%22Proc+Natl+Acad+Sci+U+S+A%22%5BJournal%5D)">Barhl2 limits growth of the diencephalic primordium through Caspase3 inhibition of beta-catenin activation.</a> Proc Natl Acad Sci U S A. 108, 2288-2293 (2011).</li>
<li>Sive, H. L., Grainger, R. M., Harland, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Dissociation+and+Reaggregation+of+Xenopus+laevis+Animal+Caps%5BTitle%5D)+AND+%22CSH+protocols%22%5BJournal%5D)">Dissociation and Reaggregation of Xenopus laevis Animal Caps.</a> CSH protocols. , (2007).</li>
<li>Harland, R. M., Grainger, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Xenopus+research%3A+metamorphosed+by+genetics+and+genomics%5BTitle%5D)+AND+%22Trends+Genet%22%5BJournal%5D)">Xenopus research: metamorphosed by genetics and genomics.</a> Trends Genet. 27, 507-515 (2011).</li>
<li>Beccari, L., Marco-Ferreres, R., Bovolenta, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+logic+of+gene+regulatory+networks+in+early+vertebrate+forebrain+patterning%5BTitle%5D)+AND+%22Mech+Dev%22%5BJournal%5D)">The logic of gene regulatory networks in early vertebrate forebrain patterning.</a> Mech Dev. 130, 95-111 (2013).</li>
<li>Pani, A. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Ancient+deuterostome+origins+of+vertebrate+brain+signalling+centres%5BTitle%5D)+AND+%22Nature%22%5BJournal%5D)">Ancient deuterostome origins of vertebrate brain signalling centres.</a> Nature. 483, 289-294 (2012).</li>
<li>Holland, L. Z., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Evolution+of+bilaterian+central+nervous+systems%3A+a+single+origin%3F%5BTitle%5D)+AND+%22Evodevo%22%5BJournal%5D)">Evolution of bilaterian central nervous systems: a single origin?</a> Evodevo. 4, 27(2013).</li>
<li>Pratt, K. G., Khakhalin, A. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Modeling+human+neurodevelopmental+disorders+in+the+Xenopus+tadpole%3A+from+mechanisms+to+therapeutic+targets%5BTitle%5D)+AND+%22Disease+models+%26+mechanisms%22%5BJournal%5D)">Modeling human neurodevelopmental disorders in the Xenopus tadpole: from mechanisms to therapeutic targets.</a> Disease models & mechanisms. 6, 1057-1065 (2013).</li>
<li>Sasai, Y., Ogushi, M., Nagase, T., Ando, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Bridging+the+gap+from+frog+research+to+human+therapy%3A+a+tale+of+neural+differentiation+in+Xenopus+animal+caps+and+human+pluripotent+cells%5BTitle%5D)+AND+%22Development%2C+growth+%26+differentiation%22%5BJournal%5D)">Bridging the gap from frog research to human therapy: a tale of neural differentiation in Xenopus animal caps and human pluripotent cells.</a> Development, growth & differentiation. 50, s47-s55 (2008).</li>
</ol>

The authors have nothing to disclose.

Neural development is orchestrated by a complex interplay between cellular developmental programs and signals from the surrounding tissues (Reviewed in3,31,32). Here we describe a set of protocols that can be used in X. laevis embryos to explore extrinsic and intrinsic factors involved in neural fate determination and neural morphogenesis in vitro and in vivo. These protocols can be used as such on X. tropicalis embryos, however X. tropicalis embryos are four times ...

The vertebrate nervous system emerges from the neural plate as a homogeneous layer of neuroepithelial cells. Understanding how developmental programs are induced, encoded, and established during regionalization of the neural plate is, at present, a major goal in developmental biology. Compared to other systems, the experimentally amenable Xenopus embryo is a model of choice for analyzing early steps of neural development1,2. It is easy to obtain large numbers of embryos, and external development gives access to the very first steps of neurulation3. Many tools are available to experimentally manipulate Xenopuslaevis (X. laevis) embryonic development. Micro-injection of mRNAs or morpholinos (MO), including inducible MOs, together with biochemical and pharmacological tools, allows controlled gain of function (GOF) and loss of function (LOF) and specific alteration of signaling pathways4,5. The blastocoel roof ectoderm, located around the animal pole of a blastula, or a very early gastrula embryo, and referred to as the &#39;Animal Cap&#39; (AC), is a source of pluripotent cells that can be programmed by manipulation of gene expression prior to explants preparation. In this manuscript are detailed protocols to use X. laevis AC explants to test in vitro and in vivo molecular mechanisms and cellular processes underlying neural development.
A technique is presented, allowing fine observation of gene expression patterns in a Xenopus tadpole neural tube, a preliminary step in the identification of fate determination cues. Whereas the observation of flat-mounted tissues is commonly used in the study of chick embryos6, it has not been properly described in Xenopus. Manipulation of gene expression by injecting synthetic mRNA or MO into the blastomeres of 2 or 4 cell stage embryos allows programming of AC explants4. For example inhibition of the Bone Morphogenetic Protein (BMP) pathway by expression of the anti-BMP factor Noggin, gives a neural identity to AC cells3. The protocol is detailed for performing local and time-controlled exposure of AC explants to extrinsic cues via direct contact with an anion exchange resin bead. Finally a technique is described for testing developmental features of neural progenitors in vivo by transplantation of mixed explants prepared from distinct programmed cells dissociated and re-associated.
The frog embryo is a powerful model to study early vertebrate neural development. Combining manipulation of gene expression to explant in vitro cultures provides important information in the study of neuroepithelium regionalization, proliferation, and morphogenesis7-12. The programming of AC explants permitted development of a functional heart ex vivo13,14. The use of explant grafting15 led to the identification of the minimal transcriptional switch inducing the neural crest differentiation program16. The zona limitans intrathalamica (ZLI) is a signaling center that secretes sonic hedgehog (Shh) to control the growth and regionalization of the caudal forebrain. When continuously exposed to Shh, neuroepithelial cells coexpressing the three transcription factor genes - barH-like homeobox-2(barhl2),orthodenticle-2 (otx2) and iroquois-3 (irx3) - acquire two characteristics of the ZLI compartment: the competence to express shh, and the ability to segregate from anterior neural plate cells. As a model system, the induction of a ZLI fate into neuroepithelial cells will be presented8.
These protocols aim at providing simple, cheap, and efficient tools for developmental biologists and other researchers to explore the fundamental mechanisms of key neural cell behaviors. These protocols are very versatile and allow the investigation of a large range of extrinsic and intrinsic neural determination cues. It permits long term in vivo analysis of neural lineage commitment, inductive interactions and cell behaviors.

<table><tbody><tr><td>Paraformaldehyde</td><td>VWR</td><td>&amp;nbsp;20909.290</td><td>Toxic</td></tr><tr><td>anion exchange resin beads</td><td>Biorad</td><td>140- 1231</td><td></td></tr><tr><td>Bovine Serum Albumin&amp;nbsp;</td><td>SIGMA</td><td>A-7888</td><td>For culture of animal cappH 7.6</td></tr><tr><td>Gentamycine&amp;nbsp;</td><td>GIBCO</td><td>15751-045&amp;nbsp;</td><td>antibiotic</td></tr><tr><td>Bovine Serum Albumin</td><td>SIGMA</td><td>A7906</td><td>&amp;nbsp;for bead preparation</td></tr></tbody></table>

stem cell-like xenopus embryonic explants to study early neural developmental features in vitro and in vivo

Understanding the genetic programs underlying neural development is an important goal of developmental and stem cell biology. In the amphibian blastula, cells from the roof of the blastocoel are pluripotent. These cells can be isolated, and programmed to generate various tissues through manipulation of genes expression or induction by morphogens. In this manuscript protocols are described for the use of Xenopus laevis blastocoel roof explants as an assay system to investigate key in vivo and in vitro features of early neural development. These protocols allow the investigation of fate acquisition, cell migration behaviors, and cell autonomous and non-autonomous properties. The blastocoel roof explants can be cultured in a serum-free defined medium and grafted into host embryos. This transplantation into an embryo allows the investigation of the long-term lineage commitment, the inductive properties, and the behavior of transplanted cells in vivo. These assays can be exploited to investigate molecular mechanisms, cellular processes and gene regulatory networks underlying neural development. In the context of regenerative medicine, these assays provide a means to generate neural-derived cell types in vitro that could be used in drug screening.

Based on morphological considerations in different species, embryological manipulations, and the expression pattern of regulatory genes, a conceptual model holds that the neural plate is divided into transverse and longitudinal segments that define a developmental grid generating distinct histogenic fields. In the neural plate, the primordia of the forebrain, midbrain, hindbrain and spinal cord are all already established along the antero-posterior (AP) axis during gastrulation (reviewed ...

Watch this Scientific Journal Video about Stem cell-like Xenopus Embryonic Explants to Study Early Neural Developmental Features In Vitro and In Vivo at JoVE.com

Stem cell-like Xenopus Embryonic Explants to Study Early Neural Developmental Features In Vitro and In Vivo

In Xenopus embryos, cells from the roof of the blastocoel are pluripotent and can be programmed to generate various tissues. Here, we describe protocols to use amphibian blastocoel roof explants as an assay system to investigate key in vivo and in vitro features of early neural development.

Stem cell-like Xenopus Embryonic Explants to Study Early Neural Developmental Features In Vitro and In Vivo

Understanding the genetic programs underlying neural development is an important goal of developmental and stem cell biology. In the amphibian blastula, ...

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Research

JoVE Journal

Developmental Biology

8.0K Views.  Institut Curie. In Xenopus embryos, cells from the roof of the blastocoel are pluripotent and can be programmed to generate various tissues. Here, we describe protocols to use amphibian blastocoel roof explants as an assay system to investigate key in vivo and in vitro features of early neural development. 

Video: Stem cell-like Xenopus Embryonic Explants to Study Early Neural Developmental Features In Vitro and In Vivo

Understanding Early Organogenesis Using a Simplified In Situ Hybridization Protocol in Xenopus

Organogenesis is the study of how organs are specified and then acquire their specific shape and functions during development. The Xenopuslaevis embryo is very useful for studying organogenesis because their large size makes them very suitable for identifying organs at the earliest steps in organogenesis. At this time, the primary method used for identifying a specific organ or primordium is whole mount in situ hybridization with labeled antisense RNA probes specific to a gene that is expressed in the organ of interest. In addition, it is relatively easy to manipulate genes or signaling pathways in Xenopus and in situ hybridization allows one to then assay for changes in the presence or morphology of a target organ. Whole mount in situ hybridization is a multi-day protocol with many steps involved. Here we provide a simplified protocol with reduced numbers of steps and reagents used that works well for routine assays. In situ hybridization robots have greatly facilitated the process and we detail how and when we utilize that technology in the process. Once an in situ hybridization is complete, capturing the best image of the result can be frustrating. We provide advice on how to optimize imaging of in situ hybridization results. Although the protocol describes assessing organogenesis in Xenopus laevis, the same basic protocol can almost certainly be adapted to Xenopus tropicalis and other model systems.

Understanding Early Organogenesis Using a Simplified In Situ Hybridization Protocol in Xenopus

The Xenopus laevis embryo continues to be exceptionally useful in the study of early development due to its large size and ease of manipulation. A simplified protocol for whole mount in situ hybridization protocol is provided that can be used in the identification of specific organs in this model system.

Organogenesis is the study of how organs are specified and then acquire their specific shape and functions during development.  The Xenopuslaevis embryo ...

Manipulation and In Vitro Maturation of Xenopus laevis Oocytes, Followed by Intracytoplasmic Sperm Injection, to Study Embryonic Development

Amphibian eggs have been widely used to study embryonic development. Early embryonic development is driven by maternally stored factors accumulated during oogenesis. In order to study roles of such maternal factors in early embryonic development, it is desirable to manipulate their functions from the very beginning of embryonic development. Conventional ways of gene interference are achieved by injection of antisense oligonucleotides (oligos) or mRNA into fertilized eggs, enabling under- or over-expression of specific proteins, respectively. However, these methods normally require more than several hours until protein expression is affected, and, hence, the interference of gene functions is not effective during early embryonic stages. Here, we introduce an experimental system in which expression levels of maternal proteins can be altered before fertilization. Xenopus laevis oocytes obtained from ovaries are defolliculated by incubating with enzymes. Antisense oligos or mRNAs are injected into defolliculated oocytes at the germinal vesicle (GV) stage. These oocytes are in vitro matured to eggs at the metaphase II (MII) stage, followed by intracytoplasmic sperm injection (ICSI). By this way, up to 10% of ICSI embryos can reach the swimming tadpole stage, thus allowing functional tests of specific gene knockdown or overexpression. This approach can be a useful way to study roles of maternally stored factors in early embryonic development.

Manipulation and In Vitro Maturation of Xenopus laevis Oocytes, Followed by Intracytoplasmic Sperm Injection, to Study Embryonic Development

We describe methods of manipulating Xenopus laevis immature oocytes, in vitro maturation of oocytes to eggs, and intracytoplasmic sperm injection. This protocol allows degradation of some maternal proteins and overexpression of genes of interest at fertilization, and hence is valuable to study roles of specific factors in early embryonic development.

Amphibian eggs have been widely used to study embryonic development. Early embryonic development is driven by maternally stored factors accumulated during ...

Methods for Study of Neuronal Morphogenesis: Ex vivo RNAi Electroporation in Embryonic Murine Cerebral Cortex

The cerebral cortex directs higher cognitive functions. This six layered structure is generated in an inside-first, outside-last manner, in which the first born neurons remain closer to the ventricle while the last born neurons migrate past the first born neurons towards the surface of the brain1. In addition to neuronal migration2, a key process for normal cortical function is the regulation of neuronal morphogenesis3. While neuronal morphogenesis can be studied in vitro in primary cultures, there is much to be learned from how these processes are regulated in tissue environments.
We describe techniques to analyze neuronal migration and/or morphogenesis in organotypic slices of the cerebral cortex4,6. A pSilencer modified vector is used which contains both a U6 promoter that drives the double stranded hairpin RNA and a separate expression cassette that encodes GFP protein driven by a CMV promoter7-9. Our approach allows for the rapid assessment of defects in neurite outgrowth upon specific knockdown of candidate genes and has been successfully used in a screen for regulators of neurite outgrowth8. Because only a subset of cells will express the RNAi constructs, the organotypic slices allow for a mosaic analysis of the potential phenotypes. Moreover, because this analysis is done in a near approximation of the in vivo environment, it provides a low cost and rapid alternative to the generation of transgenic or knockout animals for genes of unknown cortical function. Finally, in comparison with in vivo electroporation technology, the success of ex vivo electroporation experiments is not dependant upon proficient surgery skill development and can be performed with a shorter training time and skill.

Methods for Study of Neuronal Morphogenesis: Ex vivo RNAi Electroporation in Embryonic Murine Cerebral Cortex

To conduct a rapid assessment of the function of genes in the development of cerebral cortex, we describe methods involving the ex vivo electroporation of plasmids co-expressing inhibitory RNA (RNAi) and GFP in murine embryonic cortex. This protocol is amenable to the study of various aspects of neurodevelopment such as neurogenesis, neuronal migration and neuronal morphogenesis including dendrite and axon outgrowth.

The cerebral cortex directs higher cognitive functions. This six layered structure is generated in an inside-first, outside-last manner, in which the ...

Neuroscience

High-resolution Live Imaging of Cell Behavior in the Developing Neuroepithelium

The embryonic spinal cord consists of cycling neural progenitor cells that give rise to a large percentage of the neuronal and glial cells of the central nervous system (CNS). Although much is known about the molecular mechanisms that pattern the spinal cord and elicit neuronal differentiation1, 2, we lack a deep understanding of these early events at the level of cell behavior. It is thus critical to study the behavior of neural progenitors in real time as they undergo neurogenesis.
In the past, real-time imaging of early embryonic tissue has been limited by cell/tissue viability in culture as well as the phototoxic effects of fluorescent imaging. Here we present a novel assay for imaging such tissue for long periods of time, utilizing a novel ex vivo slice culture protocol and wide-field fluorescence microscopy (Fig. 1). This approach achieves long-term time-lapse monitoring of chick embryonic spinal cord progenitor cells with high spatial and temporal resolution.
This assay may be modified to image a range of embryonic tissues3, 4 In addition to the observation of cellular and sub-cellular behaviors, the development of novel and highly sensitive reporters for gene activity (for example, Notch signaling5) makes this assay a powerful tool with which to understand how signaling regulates cell behavior during embryonic development.

Imaging embryonic tissue in real-time is challenging over long periods of time. Here we present an assay for monitoring cellular and sub-cellular changes in chick spinal cord for long periods with high spatial and temporal resolution. This technique can be adapted for other regions of the nervous system and developing embryo.

The embryonic spinal cord consists of cycling neural progenitor cells that give rise to a large percentage of the neuronal and glial cells of the central ...

Expansion of Embryonic and Adult Neural Stem Cells by In Utero Electroporation or Viral Stereotaxic Injection

Somatic stem cells can divide to generate additional stem cells (expansion) or more differentiated cell types (differentiation), which is fundamental for tissue formation during embryonic development and tissue homeostasis during adulthood 1. Currently, great efforts are invested towards controlling the switch of somatic stem cells from expansion to differentiation because this is thought to be fundamental for developing novel strategies for regenerative medicine 1,2. However, a major challenge in the study and use of somatic stem cell is that their expansion has been proven very difficult to control.
Here we describe a system that allows the control of neural stem/progenitor cell (altogether referred to as NSC) expansion in the mouse embryonic cortex or the adult hippocampus by manipulating the expression of the cdk4/cyclinD1 complex, a major regulator of the G1 phase of the cell cycle and somatic stem cell differentiation 3,4. Specifically, two different approaches are described by which the cdk4/cyclinD1 complex is overexpressed in NSC in vivo. By the first approach, overexpression of the cell cycle regulators is obtained by injecting plasmids encoding for cdk4/cyclinD1 in the lumen of the mouse telencephalon followed by in utero electroporation to deliver them to NSC of the lateral cortex, thus, triggering episomal expression of the transgenes 5-8. By the second approach, highly concentrated HIV-derived viruses are stereotaxically injected in the dentate gyrus of the adult mouse hippocampus, thus, triggering constitutive expression of the cell cycle regulators after integration of the viral construct in the genome of infected cells 9. Both approaches, whose basic principles were already described by other video protocols 10-14, were here optimized to i) reduce tissue damage, ii) target wide portions of very specific brain regions, iii) obtain high numbers of manipulated cells within each region, and iv) trigger high expression levels of the transgenes within each cell. Transient overexpression of the transgenes using the two approaches is obtained by different means i.e. by natural dilution of the electroporated plasmids due to cell division or tamoxifen administration in Cre-expressing NSC infected with viruses carrying cdk4/cyclinD1 flanked by loxP sites, respectively 9,15.
These methods provide a very powerful platform to acutely and tissue-specifically manipulate the expression of any gene in the mouse brain. In particular, by manipulating the expression of the cdk4/cyclinD1 complex, our system allows the temporal control of NSC expansion and their switch to differentiation, thus, ultimately increasing the number of neurons generated in the mammalian brain. Our approach may be critically important for basic research and using somatic stem cells for therapy of the mammalian central nervous system while providing a better understanding of i) stem cell contribution to tissue formation during development, ii) tissue homeostasis during adulthood, iii) the role of adult neurogenesis in cognitive functions, and perhaps, iv) better using somatic stem cells in models of neurodegenerative diseases.

Expansion of Embryonic and Adult Neural Stem Cells by In Utero Electroporation or Viral Stereotaxic Injection

Controlling the expansion of somatic stem cells is a major factor hampering their study and use in therapy. Here we describe a system to temporally control neural stem cells expansion during development and adulthood, which can be used to increase the number of neurons generated in the mouse brain.

Somatic stem cells can divide to generate additional stem cells (expansion) or more differentiated cell types (differentiation), which is fundamental for ...

Neural Explant Cultures from Xenopus laevis

The complex process of axon guidance is largely driven by the growth cone, which is the dynamic motile structure at the tip of the growing axon. During axon outgrowth, the growth cone must integrate multiple sources of guidance cue information to modulate its cytoskeleton in order to propel the growth cone forward and accurately navigate to find its specific targets1. How this integration occurs at the cytoskeletal level is still emerging, and examination of cytoskeletal protein and effector dynamics within the growth cone can allow the elucidation of these mechanisms. Xenopus laevis growth cones are large enough (10-30 microns in diameter) to perform high-resolution live imaging of cytoskeletal dynamics (e.g.2-4 ) and are easy to isolate and manipulate in a lab setting compared to other vertebrates. The frog is a classic model system for developmental neurobiology studies, and important early insights into growth cone microtubule dynamics were initially found using this system5-7 . In this method8, eggs are collected and fertilized in vitro, injected with RNA encoding fluorescently tagged cytoskeletal fusion proteins or other constructs to manipulate gene expression, and then allowed to develop to the neural tube stage. Neural tubes are isolated by dissection and then are cultured, and growth cones on outgrowing neurites are imaged. In this article, we describe how to perform this method, the goal of which is to culture Xenopus laevis growth cones for subsequent high-resolution image analysis. While we provide the example of +TIP fusion protein EB1-GFP, this method can be applied to any number of proteins to elucidate their behaviors within the growth cone.

Neural Explant Cultures from Xenopus laevis

Culturing neural explants from dissected Xenopus laevis embryos that express fluorescent fusion proteins allows for imaging of growth cone cytoskeletal dynamics.

The complex process of axon guidance is largely driven by the growth cone, which is the dynamic motile structure at the tip of the growing axon. During ...

Fluorescent Calcium Imaging and Subsequent In Situ Hybridization for Neuronal Precursor Characterization in Xenopus laevis

Spontaneous intracellular calcium activity can be observed in a variety of cell types and is proposed to play critical roles in a variety of physiological processes. In particular, appropriate regulation of calcium activity patterns during embryogenesis is necessary for many aspects of vertebrate neural development, including proper neural tube closure, synaptogenesis, and neurotransmitter phenotype specification. While the observation that calcium activity patterns can differ in both frequency and amplitude suggests a compelling mechanism by which these fluxes might transmit encoded signals to downstream effectors and regulate gene expression, existing population-level approaches have lacked the precision necessary to further explore this possibility. Furthermore, these approaches limit studies of the role of cell-cell interactions by precluding the ability to assay the state of neuronal determination in the absence of cell-cell contact. Therefore, we have established an experimental workflow that pairs time-lapse calcium imaging of dissociated neuronal explants with a fluorescence in situ hybridization assay, allowing the unambiguous correlation of calcium activity pattern with molecular phenotype on a single-cell level. We were successfully able to use this approach to distinguish and characterize specific calcium activity patterns associated with differentiating neural cells and neural progenitor cells, respectively; beyond this, however, the experimental framework described in this article could be readily adapted to investigate correlations between any time-series activity profile and expression of a gene or genes of interest.

Fluorescent Calcium Imaging and Subsequent In Situ Hybridization for Neuronal Precursor Characterization in Xenopus laevis

We present a two-part protocol that combines fluorescent calcium imaging with in situ hybridization, allowing the experimenter to correlate patterns of calcium activity with gene expression profiles on a single-cell level.

Spontaneous intracellular calcium activity can be observed in a variety of cell types and is proposed to play critical roles in a variety of physiological ...

Generation of Na&#239;ve Blastoderm Explants from Zebrafish Embryos

Due to their optical clarity and rapid development, zebrafish embryos are an excellent system for examining cell behaviors and developmental processes. However, because of the complexity and redundancy of embryonic signals, it can be challenging to discern the complete role of any single signal during early embryogenesis. By explanting the animal region of the zebrafish blastoderm, relatively na&#239;ve clusters of embryonic cells are generated that can be easily cultured and manipulated ex vivo. By introducing a gene of interest by RNA injection before explantation, one can assess the effect of this molecule on gene expression, cell behaviors, and other developmental processes in relative isolation. Furthermore, cells from embryos of different genotypes or conditions can be combined in a single chimeric explant to examine cell/tissue interactions and tissue-specific gene functions. This article provides instructions for generating zebrafish blastoderm explants and demonstrates that a single signaling molecule - a Nodal ligand - is sufficient to induce germ layer formation and extension morphogenesis in otherwise na&#239;ve embryonic tissues. Due to their ability to recapitulate embryonic cell behaviors, morphogen gradients, and gene expression patterns in a simplified ex vivo system, these explants are anticipated to be of great utility to many zebrafish researchers.

Zebrafish blastoderm explants are generated by isolating embryonic cells from endogenous signaling centers within the early embryo, producing relatively na&#239;ve cell clusters easily manipulated and cultured ex vivo. This article provides instructions for making such explants and demonstrates their utility by interrogating roles for Nodal signaling during gastrulation.

Due to their optical clarity and rapid development, zebrafish embryos are an excellent system for examining cell behaviors and developmental processes. ...

Assessing Primary Neurogenesis in Xenopus Embryos Using Immunostaining

Primary neurogenesis is a dynamic and complex process during embryonic development that sets up the initial layout of the central nervous system. During this process, a portion of neural stem cells undergo differentiation and give rise to the first populations of differentiated primary neurons within the nascent central nervous system. Several vertebrate model organisms have been used to explore the mechanisms of neural cell fate specification, patterning, and differentiation. Among these is the African clawed frog, Xenopus, which provides a powerful system for investigating the molecular and cellular mechanisms responsible for primary neurogenesis due to its rapid and accessible development and ease of embryological and molecular manipulations. Here, we present a convenient and rapid method to observe the different populations of neuronal cells within Xenopus central nervous system. Using antibody staining and immunofluorescence on sections of Xenopus embryos, we are able to observe the locations of neural stem cells and differentiated primary neurons during primary neurogenesis.

Assessing Primary Neurogenesis in Xenopus Embryos Using Immunostaining

This article presents a convenient and rapid method for visualizing different neuronal cell populations in the central nervous system of Xenopus embryos using immunofluorescent staining on sections.

Primary neurogenesis is a dynamic and complex process during embryonic development that sets up the initial layout of the central nervous system. During ...

Dissection of Xenopus laevis Neural Crest for in vitro Explant Culture or in vivo Transplantation

The neural crest (NC) is a transient dorsal neural tube cell population that undergoes an epithelium-to-mesenchyme transition (EMT) at the end of neurulation, migrates extensively towards various organs, and differentiates into many types of derivatives (neurons, glia, cartilage and bone, pigmented and endocrine cells). In this protocol, we describe how to dissect the premigratory cranial NC from Xenopus laevis embryos, in order to study NC development in vivo and in vitro. The frog model offers many advantages to study early development; abundant batches are available, embryos develop rapidly, in vivo gain and loss of function strategies allow manipulation of gene expression prior to NC dissection in donor and/or host embryos. The NC explants can be plated on fibronectin and used for in vitro studies. They can be cultured for several days in a serum-free defined medium. We also describe how to graft NC explants back into host embryos for studying NC migration and differentiation in vivo.

Dissection of Xenopus laevis Neural Crest for in vitro Explant Culture or in vivo Transplantation

This protocol describes how to dissect premigratory cranial neural crest (NC) from Xenopus laevis neurulas. These explants can be plated on fibronectin and cultured in vitro, or grafted back into host embryos. This technique allows studying the mechanisms of NC epithelium-to-mesenchyme transition, migration, and differentiation.

Stem cell-like Xenopus Embryonic Explants to Study Early Neural Developmental Features In Vitro and In Vivo

Summary

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Dissection of Xenopus laevis Neural Crest for in vitro Explant Culture or in vivo Transplantation