We thank Radek Sindelka for his help, and Cas Bresilla for assisting with frog husbandry and embryo preparation. This work was funded by the NIH via the grant R01DE021109 to H.L.S. Laura Jacox was funded by the Herschel Smith Graduate Fellowship at Harvard University and an F30 individual fellowship grant F30DE022989-01 through the NIDCR.

1. Preparing Reagents
<ol>
	<li>10x MBS: Prepare 1 L of 10x Modified Barth&#39;s Saline (MBS) solution17. Refer to Table 1, Reagents, ingredients, and instructions. Use distilled water for all solutions. Mix in a beaker, using a stir bar, until full dissolution. All solutions should be made at room temperature.</li>
	<li>1x MBS: Dilute 100 ml of 10x MBS solution in 900 ml of distilled water to make 1 L of 1x MBS. Add 0.7 ml of 1 M CaCl2 solution.</li>
	<li>0.1x MBS: Dilute 1x MBS to prepare 1 L of 0.5x MBS solution and 2 L of 0.1x MBS solution. To 1 L of 0.1x MBS solution, add 1 ml of 10 mg/ml gentamycin solution. The 0.1x MBS solution with gentamycin will be used for long-term embryo culture.</li>
	<li>Ficoll/MBS: Add 15 grams of Ficoll 400 to 500 ml of 0.5x MBS solution. Mix vigorously. Add a stir bar and mix until Ficoll is fully dissolved (several hours).</li>
	<li>70% ethanol: Dilute 100% ethanol to 70% ethanol using distilled water.</li>
</ol>
2. Preparing Glass Operating Tools
<ol>
	<li>Needle preparation: Load capillary tubing into a needle puller.
	<ol>
		<li>Pull the needles according to the settings shown in Table 2: Needle puller settings. The settings are for a Sutter Instrument Co. Model P-80/PC micropipette puller and capillary tubings, as described in the Table of specific reagents and equipment. However, these settings are specific to the Sutter needle puller, and will need to be adjusted for other machines. Settings can be determined using a ramp test, as specified by the machine&#39;s manufacturer.</li>
		<li>Pull 4-6 needles in preparation for the procedure. The needles should be broken such that the flexible, hair-like portion of the glass tip is fully removed, which is typically 2-3 mm long. The tip must be relatively rigid, but still narrow enough to be used as a cutting tool. See photo of an ideal needle in Figure 1A.</li>
		<li>Store the needles in a Petri dish with a strip of clay down the center. Press the shaft of each needle into the clay, to hold it in place and to keep the fragile sharp tip away from the bottom and sides.</li>
	</ol>
	</li>
	<li>For additional tools, obtain glass cover slips, a pair of long standard pattern forceps, a Bunsen burner, and 3-4 glass Pasteur pipettes (Size 5 &#190; in).</li>
	<li>Pipette tool: To make a pipette tool, light the Bunsen burner and place the tip of a glass Pasteur pipette into the blue part of the flame while rotating it, such that the tip melts and the hole completely seals, forming a closed, rounded end. The rounded, sealed tip is later used to make depressions in the clay dish that holds the embryos during the operation. See Figure 1B.</li>
	<li>Glass bridges: To make glass bridges, use long standard pattern forceps to carefully break off 3 mm by 3 mm chunks of cover slip glass. While holding the piece of cover slip with tweezers, place the glass in the flame until all four edges soften and curve downward, forming a tiny glass dome. The edges should no longer be sharp. See Figure 1C.</li>
	<li>Store the glass cover slip bridges by inserting them, edges down, into modeling clay, lining the bottom of a Petri dish. Insert the bridges such that the top of the bridge remains above the clay surface. Do not push too hard such that the bridge breaks, and do not fully embed the bridge into the clay, because it can be difficult to remove. After sterilization with a flame or 70% ethanol, the bridges can be reused from experiment to experiment.</li>
</ol>
3. Preparing for the Embryo Operation
<ol>
	<li>Line a small 60 mm plastic Petri dish with modeling clay. Between uses, the dish and clay surface is thoroughly washed with distilled water and then with 70% ethanol. 
	NOTE: Use red, white or yellow, Van Aken Plastalina modeling clay, which can be bought at a local toy or art store. Black clay is not recommended because it releases a residue.</li>
	<li>Fill the dish with 3% Ficoll 0.5x MBS solution. The higher salt concentration prevents tissue dissociation, and the polysaccharide Ficoll helps to thicken the solution, which assists in holding the face in position.</li>
	<li>Use the flamed Pasteur pipette tool (see Figure 1) to make shallow, 2-3 mm depressions in the clay, about the depth of a stage 20 embryo body. Make 20-30 depressions, 1-2 mm apart, on each side of the dish, so there is a total of 40-60 depressions. Label one side LOF/GOF, and the other side wild type, by carving initials into the clay with forceps.</li>
</ol>
4. Preoperation Embryo Preparation
<ol>
	<li>Obtain and culture Xenopus laevis embryos using standard methods17. For a detailed description of frog husbandry, please see Sive et al.17
	<ol>
		<li>Forty-eight hours prior to the experiment obtain eggs from female frogs and perform in vitro fertilization.</li>
		<li>Inject 0.5-1 ng of capped membrane GFP mRNA, plus any desired mRNA or anti-sense morpholino-modified antisense oligonucleotides (&#34;morpholinos&#34;) at the one cell stage or into 2 cells at the 2 cell stage. The one cell stage lasts approximately 70-90 min. Inject a total volume of 1-3 nl. In place of RNA coding for a fluorescent protein (such as GFP or RFP RNA), one can inject FITC labeled morpholino or fluorescently labeled dextran. The fluorescence is important for determining whether the transplanted tissue heals and remains in the head.</li>
		<li>Capped mRNA can be prepared from a linearized plasmid using a mMessage mMachine SP6 or T7 kit. Morpholinos can be designed and ordered through Gene Tools LLC. The amount of morpholino required for a desired effect must be determined for each gene18.</li>
		<li>Store the injected embryos at 15 &#176;C for 48 hr, until they reach stage 19-20. (For all subsequent steps, stage embryos according to the Normal table of Xenopus laevis by Nieuwkoop and Faber19.) 
		NOTE: On the day of the surgery, both recipient and donor embryos must be within one stage of each other for the transplants to work optimally. However, embryos injected with morpholinos (&#34;morphants&#34;) sometimes develop more slowly than wild type or control morphant embryos, making it necessary to coordinate experiments so both morphant and wild type embryos are at the same stage. Morphants may need to be maintained at a higher temperature for 12-24 hr prior to the procedure. To increase the likelihood that embryos can be found at matching stages, embryos can be maintained at several temperatures. Twenty-four hours prior to the experiment, one should divide the embryos into a several dishes, and place morphants at 18-20 &#176;C and wild type embryos at 15-18 &#176;C.</li>
	</ol>
	</li>
	<li>On the day of the transplantation experiment, remove the embryos from the15 &#176;C incubator and stage them according to Nieuwkoop and Faber19. If they are younger than late neurula (stage 19), leave them at room temperature for 1-2 hr, until they reach stage 19.</li>
	<li>Screen the injected embryos under a fluorescent microscope. Select embryos that show uniform, bright fluorescence for the experiment.</li>
	<li>Remove clutter and possible contaminants near the operating area. Wipe down the operating surface, stereomicroscope and all tools with 70% ethanol.</li>
	<li>Once the embryos are at stage 19, remove the vitelline membrane using #5/45 Dumont forceps under a stereomicroscope.
	<ol>
		<li>Remove the vitelline membrane of 20-30 of each of donor and host embryos.</li>
		<li>For the first several face transplant experiments, one should practice with a few embryos for each condition. The method is challenging and requires careful practice before it can be used on a larger scale, quickly and successfully. One can work up to doing 20 transplants/experiment.</li>
	</ol>
	</li>
	<li>Move host embryos into the operating dish using a plastic graduated transfer pipette with the tip cut off such that the opening is much wider than an embryo (at least a few millimeters). Be careful to avoid touching embryos to bubbles or the water surface, as embryos will explode in the surface tension.</li>
	<li>Use #5/45 forceps to insert the embryos into the clay depressions, with the posterior of the embryo in the clay. Gently close the clay around the bases of the embryo using the forceps, leaving the top quarter of the embryo, the head, protruding from the depression.</li>
	<li>Once all host embryos are secured in their depressions, move to the other side of the plate and begin to insert the donor embryos into their depressions. Repeat the process of securing the embryos in their wells.</li>
</ol>
5. Performing the Face Transplant Surgery
<ol>
	<li>Cutting knife: As a cutting knife for removal of donor EAD tissue, use an appropriately broken glass capillary needle (see step 2.3 and Figure 1). 
	NOTE: One can directly hold the capillary between fingers (this works well for people with small hands) or one can mount the needle in an insect pin holder. Electrosharpened tungsten needles17 loaded into a pin holder may be used instead of a capillary tube needle. Please see suggested pin holders and tungsten needles in the Table of Specific Reagents and Equipment.</li>
	<li>Under a stereomicroscope, insert the needle into the head of the embryo to the left of the cement gland. The needle should be inserted deeply, so that it passes from outside the embryo, through the head, and into the foregut. To transplant the entire EAD, cuts should extend from the outside of the embryo to the foregut. For ectoderm-only transplants, cuts should be shallower and extend only through the ectoderm.
	<ol>
		<li>Flick the needle from the left to the right side of the head, across the entire width of the cement gland. The flicking is important as the motion gives a clean cut. Refer to Figure 2A for a summary of the technique and Figure 2B for a demonstration of the cuts. The cement gland and eyes are important landmarks for the cuts. The order of the cuts does not affect the outcome and can vary based on the user&#39;s preference or handedness.</li>
	</ol>
	</li>
	<li>Place the needle at the start of the previous cut, at the left border of the cement gland, and flick the needle upward until it reaches the bottom of the left eye. This will create a vertical cut from the left border of the cement gland to the bottom of the left eye.</li>
	<li>Place the needle at the right border of the cement gland, and flick the needle upward until it reaches the bottom of the right eye. This will create a vertical cut from the right border of the cement gland to the bottom of the right eye.</li>
	<li>To fully excise the tissue, flick the needle from the bottom of the left eye to the bottom of the right eye, creating a horizontal cut that will free the tissue. Cuts can extend from the outside of the embryo to the foregut, including both ectoderm and endoderm in the EAD explant. Alternately, shallow cuts can be used for ectoderm-only EAD transplants. Once the EAD tissue is removed, there should be a rectangular hole from the outside of the embryo into the foregut, stretching from the cement gland to just below the eyes (top to bottom). The hole should extend from the inside border of the left eye to the inside border of the right eye (side to side). See Figure 2Bb.</li>
	<li>Gently push the excised tissue on the tip of the needle, and lift it through the buffer to the part of the dish containing host embryos. Do not expose the tissue to the surface air or it will become damaged.</li>
	<li>Excise the same tissue from the host embryo, as for the donor. Discard the host EAD explant or save it to insert into the donor face, for reciprocal transplants.</li>
	<li>Insert the donor explant into the resulting host hole using #5/45 forceps.</li>
	<li>Once the donor tissue is correctly positioned and fully inserted, carefully place a glass bridge (see Figure 1 and Figure 2Bc) over the embryo&#39;s face to hold the transplant in place. The ends of the bridge should insert into the clay, holding it in place. The bridge should gently apply pressure to the transplanted tissue, such that the transplant lies flush with the host head, without it sticking out of the head or lying deep within the head. The head can be slightly flattened by the cover slip, but be careful not to damage the embryo with too much pressure. Transplants should be performed within 5 minutes. 
	NOTE: An experienced investigator can perform around twenty transplants per experiment, over 2-3 hr. During this period the embryos will progress from stage 20-22. Completing the face transplants at stage 21 or 22 does not affect outcomes. Later transplants (at stages 22-26) can be done but are more difficult as the crest-free EAD midline region is narrowed as cranial neural crest moves into the face. Consistency across transplants is critical.</li>
</ol>
6. Face Transplant Post-operation Recovery
<ol>
	<li>Healing typically takes 2-3 hr. Leave the embryos at room temperature undisturbed in their clay depressions with the glass bridges holding the donor tissue in place.</li>
	<li>Once the transplants have healed, carefully remove the glass bridges, remove the clay from around the base of the embryos using forceps, and use a plastic graduated transfer pipette to gently vacuum the embryos out of their depressions.</li>
	<li>Put the embryos into an appropriately labeled Petri dish, half filled with clean 0.1x MBS with gentamycin.</li>
	<li>Grow the embryos at 15 &#176;C or 18 &#176;C for several days until they reach feeding tadpole stage at stage 40, when facial phenotypes can be scored.
	<ol>
		<li>The 0.1x MBS solution with gentamycin should be changed daily, and any dead embryos should be removed promptly to prevent contamination and death of other embryos.</li>
		<li>The mouth opens at stage 40. At stage 40-41, one must check that the transplanted tissue remains healed in place by viewing its fluorescence. Transplants occasionally fall out, so one must ensure that all scored embryos have the donor tissue in place.</li>
	</ol>
	</li>
</ol>

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L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+the+primary+mouth+in+Xenopus+laevis.">Development of the primary mouth in Xenopus laevis.</a> Dev. Bio. 295, 700-713 (2006).</li><li>Dickinson, A. J., Sive, H. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Positioning+the+extreme+anterior+in+Xenopus:+cement+gland,+primary+mouth+and+anterior+pituitary.">Positioning the extreme anterior in Xenopus: cement gland, primary mouth and anterior pituitary.</a> Sem. Cell Dev. Bio. 18, 525-533 (2007).</li><li>Dickinson, A. J., Sive, H. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Wnt+antagonists+Frzb-1+and+Crescent+locally+regulate+basement+membrane+dissolution+in+the+developing+primary+mouth.">The Wnt antagonists Frzb-1 and Crescent locally regulate basement membrane dissolution in the developing primary mouth.</a> Dev. 136, 1071-1081 (2009).</li><li>Gilbert, S. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Developmental Biology. , (2010).</li><li>Borchers, A., Epperlein, H. H., Wedlich, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+assay+system+to+study+migratory+behavior+of+cranial+neural+crest+cells+in+Xenopus.">An assay system to study migratory behavior of cranial neural crest cells in Xenopus.</a> Dev. Genes Evol. 210, 217-222 (2000).</li><li>Grunz, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Homoiogenetic+neural+inducing+activity+of+the+presumptive+neural+plate+of+Xenopus+laevis.">Homoiogenetic neural inducing activity of the presumptive neural plate of Xenopus laevis.</a> Dev. Growth Differ. 32, 583-589 (1990).</li><li>Servetnick, M., Grainger, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Changes+in+neural+and+lens+competence+in+Xenopus+ectoderm:+evidence+for+an+autonomous+developmental+timer.">Changes in neural and lens competence in Xenopus ectoderm: evidence for an autonomous developmental timer.</a> Dev. Bio. 112, 177-188 (1991).</li><li>Servetnick, M., Grainger, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Homeogenetic+neural+induction+in+Xenopus.">Homeogenetic neural induction in Xenopus.</a> Dev. Bio. 147, 73-82 (1991).</li><li>Couly, G., Coltey, P., Le Douarin, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+triple+origin+of+skull+in+higher+vertebrates:+a+study+in+quail-chick+chimeras.">The triple origin of skull in higher vertebrates: a study in quail-chick chimeras.</a> Dev. 117, 409-429 (1993).</li><li>Couly, G. F., Le Douarin, N. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mapping+of+the+early+neural+primordium+in+quail-chick+chimeras+:+I.">Mapping of the early neural primordium in quail-chick chimeras : I.</a> Developmental relationships between placodes, facial ectoderm. 110, 422-439 (1985).</li><li>Couly, G. F., Le Douarin, N. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mapping+of+the+early+neural+primordium+in+quail-chick+chimeras+II.+The+prosencephalic+neural+plate+and+neural+folds:+Implications+for+the+genesis+of+cephalic+human+congenital+abnormalities.">Mapping of the early neural primordium in quail-chick chimeras II. The prosencephalic neural plate and neural folds: Implications for the genesis of cephalic human congenital abnormalities.</a> Dev. Bio. 120, 198-214 (1987).</li><li>Lievre, A. L., Le Douarin, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+early+development+of+cranial+sensory+ganglia+and+the+potentialities+of+their+component+cells+studied+in+quail-chick+chimeras.">The early development of cranial sensory ganglia and the potentialities of their component cells studied in quail-chick chimeras.</a> Dev. Bio. 94, 291-310 (1982).</li><li>Kennedy, A., Dickinson, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Median+facial+clefts+in+Xenopus+laevis:+roles+of+retinoic+acid+signaling+and+homeobox+genes.">Median facial clefts in Xenopus laevis: roles of retinoic acid signaling and homeobox genes.</a> Dev. Bio. 365, 229-240 (2012).</li><li>Trainor, P., Tam, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cranial+paraxial+mesoderm+and+neural+crest+of+the+mouse+embryo-+codistribution+in+the+craniofacial+mesenchyme+but+distinct+segregation+in+the+branchial+arches.">Cranial paraxial mesoderm and neural crest of the mouse embryo- codistribution in the craniofacial mesenchyme but distinct segregation in the branchial arches.</a> Dev. 121, 2569-2582 (1995).</li><li>Sive, H. L., Grainger, R. M., Harland, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Early Development of Xenopus laevis. , (2000).</li><li>Tandon, P., Showell, C., Christine, K., Conlon, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Morpholino+injection+in+Xenopus.">Morpholino injection in Xenopus.</a> Methods Mol. Biol. 843, 29-46 (2012).</li><li>Nieuwkoop, P. D., Faber, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Normal Table of Xenopus laevis (Daudin). , (1994).</li></ol>

The authors have nothing to disclose.

Critical Steps and Limitations: The EAD face transplant procedure is time and work intensive. It requires practice, steady hands, and dexterity to perfect. The face transplant protocol relies on the researcher's ability to efficiently remove and transplant tissue. If one takes too long to insert the transplant into the host's face, the host face will begin to contract and heal. Forceps can be used to delicately expand the facial region. However, if significant wound contraction has occurred, the transplant will no...

To understand mechanisms underlying craniofacial birth defects1-2, important tissues and their signaling contributions during craniofacial development must be identified. In the frog Xenopus laevis, part of the face, including the mouth and nostrils form from the "Extreme Anterior Domain" (EAD), where ectoderm and endoderm are directly juxtaposed3-4. The EAD also acts as a signaling center to influence surrounding tissues, including the cranial neural crest, which forms the jaws and other facial regions5. To identify genes that contribute to EAD function, a 'face transplant' technique was developed, where tissue is transplanted from a donor into a host embryo, after removing the corresponding host region. Following the transplant, resulting facial development is assessed. Thus, the effects of loss of function (LOF) or gain of function (GOF) for a specific gene&#160;in the EAD are analyzed locally, where the rest of the head and body is composed of wild type tissue. The reciprocal transplant can be performed, where wild type tissue is transplanted into embryos with global LOF or GOF in specific genes. Transplantation has been frequently used in Xenopus and chick studies6. For example, Xenopus transplantation has addressed homogenetic neural induction, lens and neural competence, and neural crest migration7-10. Quail-chick chimeric grafting has analyzed development of the anterior neural plate, anterior neural ridge, neural crest, and cranial bones11-14. This is the first transplant technique for study of craniofacial development in Xenopus. This technique has demonstrated a novel role for the Wnt inhibitors Frzb1 and Crescent in regulating basement membrane formation in the presumptive mouth5. Xenopus laevis is an ideal model for study of craniofacial development as embryos are large, develop externally, and the face is readily visible, allowing micromanipulation and imaging of development. Mechanisms underlying facial development appear conserved, indicating that findings made in the frog provide insight into human development4,15-16.

<table><tbody><tr><td>Pasteur pipette</td><td>VWR</td><td>14672-400</td><td>Lime Glass&amp;nbsp;</td></tr><tr><td>Size 5 3/4 in</td><td>Cotton Plugged</td><td></td><td></td></tr><tr><td>Graduated Transfer Pipette</td><td>VWR</td><td>16001-180</td><td>Disposable&amp;nbsp;</td></tr><tr><td>#5/45 forceps</td><td>Fine Science Tools by Dupont medical</td><td>11251-35</td><td>Angled 45&amp;deg;</td></tr><tr><td>Standard Pattern Forceps</td><td>Fine Science Tools</td><td>11000-20</td><td>Straight; serrated tip; stainless steel</td></tr><tr><td>Capillary Tubing (for needles)</td><td>FHC</td><td>30-30-1</td><td>Borosil 1.0 mm OD x 0.5 mm ID/Fiber</td></tr><tr><td>Cover slip&amp;nbsp;</td><td>VWR</td><td>48393 252</td><td>24 x 60 mm&amp;nbsp;micro cover glass;</td></tr><tr><td>Ficoll 400</td><td>Sigma-Aldrich</td><td>F9378</td><td></td></tr><tr><td>Needle Puller&amp;nbsp;</td><td>Sutter Instrument Co</td><td>Needle Puller: discontinued Filament: FB300B</td><td>The most similar, currently available&amp;nbsp; needle puller is the P-97. For filaments, use Sutter 3.00 mm square box filaments, 3.0 mm wide.</td></tr><tr><td>Model P-80</td><td>Flaming / Brown micropipette puller</td><td></td><td></td></tr><tr><td>Stereomicroscope</td><td>Zeiss</td><td></td><td></td></tr><tr><td>Stereomicroscope Lighting by Fostec</td><td>Fostec</td><td></td><td>Use a light box with 2 fiberoptic arms.&amp;nbsp;&amp;nbsp;</td></tr><tr><td>Nickel Plated Pin Holder</td><td>Fine Science Tools</td><td>26018-17</td><td>Jaw Opening Diameter: 0-1 mm</td></tr><tr><td>Moria Nickel Plated Pin Holder</td><td>Fine Science Tools</td><td>26016-12</td><td>Jaw opening Diameter: 0-1 mm</td></tr><tr><td>Tungsten Needles</td><td>Fine Science Tools</td><td>10130-05</td><td>0.125 mm Rod diameter</td></tr><tr><td>Van Aken Plastalina</td><td>Blick</td><td>#33268-2981</td><td></td></tr><tr><td>Modeling Clay- white, red, or yellow</td><td></td><td></td><td></td></tr><tr><td>mMessage mMashine SP6 or T7 Kit</td><td>Ambion</td><td>AM1340</td><td></td></tr></tbody></table>

facial transplants in xenopus laevis embryos

Craniofacial birth defects occur in 1 out of every 700 live births, but etiology is rarely known due to limited understanding of craniofacial development. To identify where signaling pathways and tissues act during patterning of the developing face, a 'face transplant' technique has been developed in embryos of the frog Xenopus laevis. A region of presumptive facial tissue (the "Extreme Anterior Domain" (EAD)) is removed from a donor embryo at tailbud stage, and transplanted to a host embryo of the same stage, from which the equivalent region has been removed. This can be used to generate a chimeric face where the host or donor tissue has a loss or gain of function in a gene, and/or includes a lineage label. After healing, the outcome of development is monitored, and indicates roles of the signaling pathway within the donor or surrounding host tissues. Xenopus is a valuable model for face development, as the facial region is large and readily accessible for micromanipulation. Many embryos can be assayed, over a short time period since development occurs rapidly. Findings in the frog are relevant to human development, since craniofacial processes appear conserved between Xenopus and mammals.

Transplanted tissue should be fully inserted into the host head after transplantation as shown in Figure 3A, and have a glass bridge appropriately placed on the embryo's face, as shown in Figure 2Bc. The transplanted donor tissue must be correctly sized for the host opening, for the transplant to be successful. The EAD tissue should not protrude from the head, in any way, as seen in Figures 3B and 3C. Additionally, the face transplant should not be rotat...

Watch this Scientific Journal Video about Facial Transplants in Xenopus laevis Embryos at JoVE.com

Facial Transplants in Xenopus laevis Embryos

A technique for transplanting "Extreme Anterior Domain" facial tissue between Xenopus laevis embryos has been developed. Tissue can be moved from one gene expression background into another, allowing the study of local requirements for craniofacial development and for signaling interactions between facial regions.

Facial Transplants in Xenopus laevis Embryos

Craniofacial birth defects occur in 1 out of every 700 live births, but etiology is rarely known due to limited understanding of craniofacial development. ...

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Biology

10.2K Views.  Harvard University.  Een techniek voor het transplanteren "Extreme Voorste Domain" tissues tussen Xenopus laevis embryo ontwikkeld. Weefsel kan worden verplaatst van de ene genexpressie achtergrond in de andere, waardoor de studie van de lokale vereisten voor craniofaciale ontwikkeling en voor het signaleren van interacties tussen het gezicht regio. 

Video: Facial Transplants in Xenopus laevis Embryos

Primordial Germ Cell Transplantation for CRISPR/Cas9-based Leapfrogging in Xenopus

The creation of mutant lines by genome editing is accelerating genetic analysis in many organisms. CRISPR/Cas9 methods have been adapted for use in the African clawed frog, Xenopus, a longstanding model organism for biomedical research. Traditional breeding schemes for creating homozygous mutant lines with CRISPR/Cas9-targeted mutagenesis have several time-consuming and laborious steps. To facilitate the creation of mutant embryos, particularly to overcome the obstacles associated with knocking out genes that are essential for embryogenesis, a new method called leapfrogging was developed. This technique leverages the robustness of Xenopus embryos to &#34;cut and paste&#34; embryological methods. Leapfrogging utilizes the transfer of primordial germ cells (PGCs) from efficiently-mutagenized donor embryos into PGC-ablated wildtype siblings. This method allows for the efficient mutation of essential genes by creating chimeric animals with wildtype somatic cells that carry a mutant germline. When two F0 animals carrying &#34;leapfrog transplants&#34; (i.e., mutant germ cells) are intercrossed, they produce homozygous, or compound heterozygous, null F1 embryos, thus saving a full generation time to obtain phenotypic data. Leapfrogging also provides a new approach for analyzing maternal effect genes, which are refractory to F0 phenotypic analysis following CRISPR/Cas9 mutagenesis. This manuscript details the method of leapfrogging, with special emphasis on how to successfully perform PGC transplantation.

Primordial Germ Cell Transplantation for CRISPR/Cas9-based Leapfrogging in Xenopus

Genes essential for survival pose technical hurdles for creating mutant lines. Leapfrogging circumvents lethality by combining genome editing with primordial germ cell transplantation to create wild-type animals carrying germline mutations. Leapfrogging also permits the efficient generation of homozygous null mutants in the F1 generation. Here, the transplantation step is demonstrated.

The creation of mutant lines by genome editing is accelerating genetic analysis in many organisms. CRISPR/Cas9 methods have been adapted for use in the ...

Genetics

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.

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.

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

Developmental Biology

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 ...

Microinjection of DNA into Eyebuds in Xenopus laevis Embryos and Imaging of GFP Expressing Optic Axonal Arbors in Intact, Living Xenopus Tadpoles

The primary visual projection of tadpoles of the aquatic frog Xenopus laevis serves as an excellent model system for studying mechanisms that regulate the development of neuronal connectivity. During establishment of the retino-tectal projection, optic axons extend from the eye and navigate through distinct regions of the brain to reach their target tissue, the optic tectum. Once optic axons enter the tectum, they elaborate terminal arbors that function to increase the number of synaptic connections they can make with target interneurons in the tectum. Here, we describe a method to express DNA encoding green fluorescent protein (GFP), and gain- and loss-of-function gene constructs, in optic neurons (retinal ganglion cells) in Xenopus embryos. We explain how to microinject a combined DNA/lipofection reagent into eyebuds of one day old embryos such that exogenous genes are expressed in single or small numbers of optic neurons. By tagging genes with GFP or co-injecting with a GFP plasmid, terminal axonal arbors of individual optic neurons with altered molecular signaling can be imaged directly in brains of intact, living Xenopus tadpoles several days later, and their morphology can be quantified. This protocol allows for determination of cell-autonomous molecular mechanisms that underlie the development of optic axon arborization in vivo.

Microinjection of DNA into Eyebuds in Xenopus laevis Embryos and Imaging of GFP Expressing Optic Axonal Arbors in Intact, Living Xenopus Tadpoles

This protocol aims to demonstrate how to microinject a DNA/DOTAP mixture into eyebuds of one day old Xenopus laevis embryos, and how to image and reconstruct individual green fluorescent protein (GFP) expressing optic axonal arbors in tectal midbrains of intact, living Xenopus tadpoles.

The primary visual projection of tadpoles of the aquatic frog Xenopus laevis serves as an excellent model system for studying mechanisms that regulate the ...

Tissue Determination Using the Animal Cap Transplant (ACT) Assay in Xenopus laevis

Many proteins play a dual role in embryonic development. Those that regulate cell fate determination in a specific tissue can also affect the development of a larger region of the embryo. This makes defining its role in a particular tissue difficult to analyze. For example, noggin overexpression in Xenopus laevis embryos causes the expansion of the entire anterior region, including the eye1,2. From this result, it is not known if Noggin plays a direct role in eye determination or that by causing an expansion of neural tissue, Noggin indirectly affects eye formation. Having this complex phenotype makes studying its eye-specific role in cell fate determination difficult to analyze. We have developed an assay that overcomes this problem. Taking advantage of the pluripotent nature of the Xenopus laevis animal cap 3, we have developed an assay to test the ability of gene product(s), like noggin or the eye field transcription factors (EFTFs), to transform caps into particular tissue or cell types by transplanting this tissue onto the side of the embryo 4. While we have found either Noggin protein treatment or a collection of transcription factors can determine retinal cell fate in animal caps, this procedure could be used to identify gene product(s) involved in specifying other tissues as well.

Tissue Determination Using the Animal Cap Transplant ACT Assay in Xenopus laevis

Animal caps overexpressing gene product(s) are transplanted to the flank of developing Xenopus laevis embryos in order to establish whether tissue is determined.

Many proteins play a dual role in embryonic development. Those that regulate cell fate determination in a specific tissue can also affect the development ...

Production of Transgenic Xenopus laevis by Restriction Enzyme Mediated Integration and Nuclear Transplantation

Stable integration of cloned gene products into the Xenopus genome is necessary to control the time and place of expression, to express genes at later stages of embryonic development, and to define how enhancers and promoters regulate gene expression within the embryo. The protocol demonstrated here can be used to efficiently produce transgenic Xenopus laevis embryos. This transgenesis approach involves three parts: 1. Sperm nuclei are isolated from adult X. laevis testis by treatment with lysolecithin, which permeabilizes the sperm plasma membrane. 2. Egg extract is prepared by low speed centrifugation, addition of calcium to cause the extract to progress to interphase of the cell cycle, and a high-speed centrifugation to isolate interphase cytosol. 3. Nuclear transplantation: the nuclei and extract are combined with the linearized plasmid DNA to be introduced as the transgene and a small amount of restriction enzyme. During a short reaction, egg extract partially decondenses the sperm chromatin and the restriction enzyme generates chromosomal breaks that promote recombination of the transgene into the genome. The treated sperm nuclei are then transplanted into unfertilized eggs. Integration of the transgene usually occurs prior to the first embryonic cleavage such that the resulting embryos are not chimeric. These embryos can be analyzed without any need to breed to the next generation, allowing for efficient and rapid generation of transgenic embryos for analyses of promoter and gene function. Adult X. laevis resulting from this procedure also propagate the transgene through the germline and can be used to generate lines of transgenic animals for multiple purposes.

Production of Transgenic Xenopus laevis by Restriction Enzyme Mediated Integration and Nuclear Transplantation

This video protocol demonstrates a method for generating transgenic Xenopus laevis by introduction of transgenes into sperm nuclei followed by nuclear transplantation into unfertilized eggs.

Stable integration of cloned gene products into the Xenopus genome is necessary to control the time and place of expression, to express genes at later ...

Creating Avian Forebrain Chimeras to Assess Facial Development

The avian embryo has been used as a model system for more than a century and has led to fundamental understanding of vertebrate development. One of the strengths of this model system is that the effect of, and interaction among, tissues can be directly assessed in chimeric embryos. We have previously shown that signals from the forebrain contribute to facial morphogenesis by regulating the shape of the expression domain of Sonic hedgehog (SHH) in the Frontonasal Ectodermal Zone (FEZ). In this article, the method of generating the forebrain chimeras and provide illustrations of the outcomes of these experiments is described.

This article describes a tissue transplantation technique that was designed to test the signaling and patterning properties of basal forebrain&#160;during craniofacial development.

The avian embryo has been used as a model system for more than a century and has led to fundamental understanding of vertebrate development. One of the ...

Polarized Translocation of Fluorescent Proteins in Xenopus Ectoderm in Response to Wnt Signaling

Cell polarity is a fundamental property of eukaryotic cells that is dynamically regulated by both intrinsic and extrinsic factors during embryonic development 1, 2. One of the signaling pathways involved in this regulation is the Wnt pathway, which is used many times during embryogenesis and critical for human disease3, 4, 5. Multiple molecular components of this pathway coordinately regulate signaling in a spatially-restricted manner, but the underlying mechanisms are not fully understood. Xenopus embryonic epithelial cells is an excellent system to study subcellular localization of various signaling proteins. Fluorescent fusion proteins are expressed in Xenopus embryos by RNA microinjection, ectodermal explants are prepared and protein localization is evaluated by epifluorescence. In this experimental protocol we describe how subcellular localization of Diversin, a cytoplasmic protein that has been implicated in signaling and cell polarity determination6, 7 is visualized in Xenopus ectodermal cells to study Wnt signal transduction8. Coexpression of a Wnt ligand or a Frizzled receptor alters the distribution of Diversin fused with red fluorescent protein, RFP, and recruits it to the cell membrane in a polarized fashion 8, 9. This ex vivo protocol should be a useful addition to in vitro studies of cultured mammalian cells, in which spatial control of signaling differs from that of the intact tissue and is much more difficult to analyze.

Polarized Translocation of Fluorescent Proteins in Xenopus Ectoderm in Response to Wnt Signaling

Xenopus embryonic ectoderm has become an attractive model for studies of cell polarity. An assay is described, in which subcellular distribution of fluorescent proteins is assessed in ectoderm cells. This protocol will help address questions related to spatial control of signaling.

Cell polarity is a fundamental property of eukaryotic cells that is dynamically regulated by both intrinsic and extrinsic factors during embryonic ...

Electroporation of Craniofacial Mesenchyme

Electroporation is an efficient method of delivering DNA and other charged macromolecules into tissues at precise time points and in precise locations. For example, electroporation has been used with great success to study neural and retinal development in Xenopus, chicken and mouse 1-10. However, it is important to note that in all of these studies, investigators were not targeting soft tissues. Because we are interested in craniofacial development, we adapted a method to target facial mesenchyme.
When we searched the literature, we found, to our surprise, very few reports of successful gene transfer into cartilaginous tissue. The majority of these studies were gene therapy studies, such as siRNA or protein delivery into chondrogenic cell lines, or, animal models of arthritis 11-13. In other systems, such as chicken or mouse, electroporation of facial mesenchyme has been challenging (personal communications, Dept of Craniofacial Development, KCL). We hypothesized that electroporation into procartilaginous and cartilaginous tissues in Xenopus might work better. In our studies, we show that gene transfer into the facial cartilages occurs efficiently at early stages (28), when the facial primordium is still comprised of soft tissue prior to cartilage differentiation.
Xenopus is a very accessible vertebrate system for analysis of craniofacial development. Craniofacial structures are more readily visible in Xenopus than in any other vertebrate model, primarily because Xenopus embryos are fertilized externally, allowing analyses of the earliest stages, and facilitating live imaging at single cell resolution, as well as reuse of the mothers 14. Among vertebrate models developing externally, Xenopus is more useful for craniofacial analysis than zebrafish, as Xenopus larvae are larger and easier to dissect, and the developing facial region is more accessible to imaging than the equivalent region in fish. In addition, Xenopus is evolutionarily closer to humans than zebrafish (&tilde;100 million years closer) 15. Finally, at these stages, Xenopus tadpoles are transparent, and concurrent expression of fluorescent proteins or molecules will allow easy visualization of the developing cartilages. We anticipate that this approach will allow us to rapidly and efficiently test candidate molecules in an in vivo model system.

Craniofacial cartilages develop in close contact with other tissues and are difficult to manipulate in live animals. We are using electroporation to deliver molecular tools during growth of the craniofacial skeleton while bypassing early embryonic effects. This approach will allow us to efficiently test candidate molecules in vivo.

Electroporation is an efficient method of delivering DNA and other charged macromolecules into tissues at precise time points and in precise locations. ...

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.

Facial Transplants in Xenopus laevis Embryos

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