This work was funded by a National Institute of Dental and Craniofacial Research (NIDCR) F32 grant (DE021929) to J.L.F. and a NIDCR R01 grant DE016402 to R.A.S.

1. Prepare Tungsten Needles
<ol>
	<li>Cut a tungsten rod in half using wire cutters so that the rod does not bend.</li>
	<li>Thread the rod through the tapered end of a 5&#160;3/4 in&#160;borosilicate glass Pasteur pipette.</li>
	<li>Leaving approximately 3/4 in&#160;of the rod sticking out of the glass, adhere the rod to the pipette tip with a small bead of &#8220;hot melt adhesive&#8221; (HMA), which is the glue stick used for a glue gun. The HMA should be quickly melted in an alcohol flame, taking care not to burn the glue and generate black smoke.</li>
	<li>Using a pair of forceps, bend the tip of the tungsten rod (about 1/4 in&#160;from the top end) until a 45&#176;&#160;angle is reached.</li>
	<li>Holding the Pasteur pipette, place about 1/8 in&#160;of the tip of the tungsten rod into the flame of a propane torch. Hold the tip steady and exactly perpendicular to the flame.</li>
	<li>Pull the needle out of the flame the moment a tiny orange speck of tungsten flies off of the needle.</li>
</ol>
2. Prepare Spemann Pipettes
<ol>
	<li>Using a Bunsen burner, heat the narrow end of a Pasteur pipette until the glass shortly beyond the taper begins to melt. Remove from flame and pull the tip until the pipette becomes sealed. Be sure that there remains a portion of the tapered part that has an O.D. of approximately 0.7-1.0 mm. Break the sealed end off about 4 cm from the taper.</li>
	<li>Attach around 2 ft&#160;(60 cm) of rubber tubing to the wide (open) end of the pipette. Blow on the other end of the rubber tubing to ensure that no air can pass through the tapered part of the Pasteur pipette.</li>
	<li>Using a propane fuel cylinder with an attached pencil flame torch, heat an oval area on one side of the pipette just below the start of the taper. Blow air very gently and constantly through the rubber tubing with a slight amount of pressure (approximately the amount of pressure needed to say the beginning of the letter &#8220;P&#8221; at conversation level). When the glass becomes soft on one side and begins to buckle outwards, quickly remove the pipette from the flame and blow hard and steady through the rubber tubing. This will cause the heated part of the glass to form a sausage-shaped bubble that may pop, which is fine. If the bubble is too small, try melting and blowing the glass again. The final open window in the glass should be around 0.25 in&#160;(6.35 mm) wide by 0.5 in&#160;(12.7 mm) long.</li>
	<li>Pointing the tapered end upwards, scrape the bubble into a glass waste container while gently blowing through the tubing to prevent glass fragments from entering the pipette.</li>
	<li>Using the propane flame, burn off the remaining edges of the bubble and fire-polish the sides of the open window. Take care not to melt the shaft of the pipette.</li>
	<li>Using a diamond point pencil, score the tapered end of the pipette approximately 1.25 in&#160;(30 mm) from the window and break off the tip with forceps so that the pipette opening is cut cleanly (discard pipettes with broken or uneven tips).</li>
	<li>Using forceps and the edge of a flame from an alcohol burner, bend the narrow portion of the pipette 0.375 in&#160;(9.5 mm) from the tip to an angle of approximately 60&#176; so that the bent portion is in a vertical plane when the window opening is at the 1:00 position for right-handed usage and 11:00 for left-handed usage. This step is best conducted in a draft-free enclosure.</li>
	<li>Fire-polish the tip using the alcohol flame under a dissecting microscope. Be sure not to close off the tip but rather keep the opening round and smooth without any constriction.</li>
	<li>Cut a 1&#160;in&#160;(25 mm) piece of rubber tubing, spray the piece of tubing with 70% ethanol, and slide the tubing over the shaft of the pipette until the window opening is completely covered. This functions as the &#8220;diaphragm&#8221; of the Spemann pipette.</li>
	<li>Place a latex rubber 2 ml bulb over the open bottom of the pipette. The bulb maintains negative pressure in the Spemann pipette.</li>
	<li>To practice using a Spemann pipette, transfer chromatography beads of around 100-200 &#181;m in diameter from a marked spot in one Petri dish to a marked spot in another under a microscope.</li>
	<li>To clean the Spemann pipette, remove the rubber bulb and place pipette, tip down, in a tall beaker lined with cotton or gauze on the bottom and filled with distilled water and glassware detergent.</li>
</ol>
3. Sterilize and Incubate Eggs
<ol>
	<li>Wipe eggs with 70% ethanol and place the eggs on plastic egg trays.</li>
	<li>Set eggs in a heated incubator at 37.5 &#176;C and 85-87%.</li>
	<li>Incubate eggs until they reach HH9.5, approximately 26 hr for quail and 48 hr for duck.</li>
</ol>
4. Window Eggs
<ol>
	<li>Remove a small piece of outer shell from the top of the egg with forceps, taking care not to puncture the inner shell membrane.</li>
	<li>Using a syringe with an 18 G by 1-inch needle, poke a hole at the narrowest tip of the egg and withdraw approximately 1-2 ml of albumen.</li>
	<li>Place transparent tape over the hole and puncture mark in the shell.</li>
	<li>Cut a &#8220;window&#8221; along the top surface of the egg (through the transparent tape) using curved scissors.</li>
</ol>
5. Visualize Embryos and Prepare for Surgery
<ol>
	<li>Using a glass rod, gently brush a small amount of Neutral Red (0.02 g/ml in Hank&#8217;s BSS) over the embryo.</li>
	<li>Cut the vitelline membrane and pull it over the embryo using a flame-sharpened tungsten needle.</li>
	<li>Re-seal the egg by placing transparent tape over the window.</li>
</ol>
6. Separate the Donor Tissue from the Donor Embryo
<ol>
	<li>Remove tape from the window on the egg of the donor embryo.</li>
	<li>To cut the neural fold from the adjacent ectoderm of the donor embryo, make slits on both sides of the neural tube, using a flame-sharpened tungsten needle (manufactured as described above).</li>
	<li>Then, cut across the neural tube at the desired anterior and posterior levels of the graft and separate the graft from the rest of the neural tube.</li>
	<li>Remove the neural fold from the donor embryo using either a Spemann pipette or other type of micropipette.</li>
	<li>Then, remove the tape from the window on the host egg and use the Spemann pipette to place the donor neural fold alongside the host embryo adjacent to the region of the neural tube that will receive the transplant.</li>
</ol>
7. Separate the Host Tissue and Transplant the Donor Tissue
<ol>
	<li>Separate the neural fold from the neural tube of the host embryo, as done with the donor. Take care to remove a graft of equal size as the donor tissue.&#160;</li>
	<li>Gently push this host tissue far away from the embryo.</li>
	<li>Then, with a blunt tungsten needle or the rounded tip of a micropipette, gently move the donor neural fold into the host neural tube, maintaining the proper antero-posterior and dorso-ventral orientations. Make sure the graft is tucked in along the sides but be sure not to poke or damage to the underlying tissues.&#160;To help keep track of the original orientation note any asymmetry in the donor tissue or differential staining from the Neutral Red. We also photo-document the donor embryo with the accompanying excised tissue, as well as the chimera immediately following surgery.</li>
	<li>Sterile saline should be added to the egg if the host embryo shows signs of desiccation.</li>
	<li>Now, carefully re-seal and label the egg, and gently return it to a high humidity incubator (70-80%) where the chimeric embryo can develop to the desired stage for analysis.</li>
</ol>
8. Collection of Chimeras
<ol>
	<li>Be sure to collect chimeric embryos in freshly made&#160;cold Serra&#8217;s fixative and immediately place them on a rocker at 4 &#176;C for an O/N fixation. This will allow for more&#160;sensitive detection of quail cells with the anti-quail antibody.</li>
	<li>For gene expression analyses via RT-qPCR, freeze embryos directly in liquid nitrogen prior to RNA extraction.</li>
</ol>

The authors have nothing to disclose.

The neural crest is a transient embryonic cell population that migrates extensively throughout the embryo and differentiates into diverse cell types, including chondrocytes and osteoblasts, that contribute to the craniofacial skeleton. Transplanting neural crest in the quail-duck chimeric system has contributed greatly to our understanding of the tissue interactions and signaling pathways that regulate development of the craniofacial skeleton. However, given the vast potential of neural crest to also generate smooth muscle cells, adipocytes, melanocytes, schwann cells, and neurons, the quail-duck chimera system has tremendous potential for future applications, particularly in conjunction with the rapid advancement of stem cell biology and regenerative medicine. Since quail and duck are both commercially bred species, a ready supply of relatively inexpensive fertilized eggs is available from a variety of farms. Thus, this technique should be accessible to researchers operating within a wide range of budgets and facility space.
Although this technique is very powerful, there remain several limitations. Like other surgical techniques, the quality and viability of quail-duck chimeras rely on the surgical skills of the researcher, and therefore, there will be more inter- and intra-individual variation between experiments as compared to other models, such as those utilizing mouse genetics. Moreover, there is also variation in the rates of development and stages of individual embryos that contributes to the reproducibility and success of each transplant. Avian embryos are also very susceptible to dehydration and therefore critical steps during surgery include keeping the light levels low, the time under the microscope to a minimum, the eggs sealed with tape as much as possible, and high humidity in the post-operative incubator to avoid desiccation.&#160;
In terms of viability of the chimeras, usually between 50-75% survive, although these percentages can decrease the older the collection stage.&#160;In a typical 4-6 hr session of surgery, an experienced surgeon can generate 10-15 chimeras.&#160;The success of the transplants also depends greatly on the quality of the tools. Good tools lead to more consistent, reproducible results. Using a propane torch to make tungsten needles allows extremely sharp needles to be made.&#160;The type of torch used makes a big difference because it controls the size of the flame. Electrolytic sharpening can also be used, but this approach does not even come close to producing needles as sharp.&#160;Use tungsten rods instead of spooled wire so that the needles can be made straight.
The Spemann micropipette, while time-consuming and difficult to make, is an ideal instrument for tissue&#160;transfer.&#160;The pipette can be customized with different-sized openings, and can be used repeatedly. A critical factor for using a Spemann&#160;micropipette is to have some fluid in the pipette before touching the tip to the surface of the embryo. Some of the fluid will always flow out when contact is made with the meniscus over the embryo.&#160;Pressing&#160;on the diaphragm allows fluid and graft tissue to be ejected very precisely, whereas slightly letting up on the diaphragm gently sucks the donor graft tissue into the pipette.&#160;Maintaining a bit of positive pressure on the diaphragm keeps the donor graft tissue at the tip of pipette during transfer, and a little additional pressure on the diaphragm allows the donor graft tissue to be deliberately placed in the host.&#160;
For protection of the Spemann pipette during storage and sterilization, remove the bulb from the wide end and carefully insert the tapered tip into the bulb.&#160;Place the Spemann pipette in a glass test tube, cover the top with aluminum foil, and autoclave prior to surgery. After multiple pipettes are ready to be sterilized again for surgery, invert the pipettes, heat them nearly to&#160;a boil in the same solution of distilled water and glassware detergent, and then rinse repeatedly with distilled water. Autoclave pipettes in their individual&#160;tubes. The rubber tubing that forms the diaphragm and the rubber bulb should be replaced after several sterilizations or when they become darkened and stiff.
Many of the components of this protocol involve dangerous equipment. For example, the procedure for making a Spemann Micropipette involves three types of flames as well as heating, pulling, blowing, bending, cutting, and polishing glass. Therefore, wearing proper personal protective equipment (PPE) that increases safety such as goggles and a lab coat is critical. Additionally, because many people suffer from, or have the potential to develop, egg allergies, always use gloves when handling eggs. With these precautions in mind, the quail-duck chimeric system is a safe, efficient, and relatively accessible method that has many future applications.

&#160;The facial skeleton develops from the growth and fusion of multiple facial processes that are composed of neural crest and mesodermal mesenchyme surrounded by ectodermal and endodermal epithelial layers1-11. Morphogenetic events within each process are governed by distinct signaling interactions between the mesenchyme and surrounding epithelia12-16. Alterations to these signaling interactions and/or their downstream effectors contribute to disease phenotypes and may also be relevant to evolution of the craniofacial skeleton17,18. Therefore, elucidating the timing and nature of tissue interactions has great potential to increase our understanding of the developmental and evolutionary biology of the facial skeleton.
The use of chimeric embryos to investigate tissue interactions has a long history in developmental biology. This approach was pioneered by Hans Spemann and his lab who discovered embryonic &#8220;organizers&#8221; by transplanting tissues between embryos of different amphibian species. Spemann was a master of micro-surgical techniques whose hand-skills were complemented by his development of specialized tools, notably the Spemann pipette. Viktor Hamburger was a graduate student in the laboratory of Hans Spemann in Freiburg during the 1920s, which is when the original transplant experiments that led to Spemann&#8217;s Nobel Prize were performed.&#160;When Hamburger moved to Washington University in St. Louis in 1935, he detailed the process of making a Spemann micropipette in his Manual of Experimental Embryology19.&#160;Drew Noden was a graduate student in Hamburger&#8217;s lab at Washington University until 1972.&#160;After moving to the University of Massachusetts, Amherst and then to Cornell University, Noden continued fabricating and using Spemann micropipettes for his surgical transplants involving quail-chick chimeras.&#160;While a graduate student, one of the authors (Rich Schneider) trained with Drew Noden at Cornell from 1995 to 1998.&#160;The following protocol for making a Spemann micropipette is based on descriptions written by Hamburger and Noden, and includes subsequent modifications made by Schneider.
The use of quail-chick chimeras for the study of craniofacial development and especially for understanding the contributions of neural crest cells was pioneered by Noden and by Le Douarin in the early 1970s, reviewed in Le Douarin et al20. This approach has been broadly adopted in many studies and by numerous other investigators1,4,5,21-38. The equivalent rates of growth and morphology of quail and chick make transplants within them ideal for the study of cell fate and lineage tracing. However, because of the similarities between quail and chick, morphological changes induced by donor cells are difficult to decipher. In contrast, other avian chimeric systems have included domestic duck as a way to study mechanisms that make embryos anatomically distinct39-50.&#160;More specifically, the quail-duck chimeric system offers multiple benefits for discerning the effects of the donor on the host, and vice versa. First, quail and duck embryos are distinct in body size and shape, which provides a direct way to explore donor- or host-specific mechanisms of pattern formation by assaying for differential domains of gene expression (Figures 1 A&#160;and B). Second, quail and duck embryos have considerably different rates of maturation, with quail hatching in 17 days and duck hatching in 28 days. Transplanted neural crest maintains its intrinsic maturation rate within the host environment, and thus, identification of temporal changes in gene expression, tissue interactions, histogenesis, and morphogenesis is possible51-57. Finally, the anti-quail nuclear antibody (Q&#162;PN) allows donor and host cellular contributions to be permanently distinguished from one another by recognizing a protein that is ubiquitously expressed in quail cells but absent from duck cells.

<table><tbody><tr><td>1x PBS</td><td>TEK</td><td>TEKZR114</td><td></td></tr><tr><td>Hank&amp;rsquo;s BSS w/o phenol red</td><td>Invitrogen</td><td>14025-092</td><td></td></tr><tr><td>Neutral Red</td><td>Sigma Aldrich</td><td>N4638-5G</td><td>0.22 &amp;micro;m filter-sterilized</td></tr><tr><td>18 G Needles</td><td>BD</td><td>305195</td><td></td></tr><tr><td>5 ml syringe</td><td>BD</td><td>309646</td><td></td></tr><tr><td>No. 5 Dumont forceps</td><td>Fine Science Tools</td><td>11252-20</td><td></td></tr><tr><td>Straight Scissors</td><td>Fine Science Tools</td><td>14028-10</td><td></td></tr><tr><td>Curved Scissors</td><td>Fine Science Tools</td><td>14029-10</td><td></td></tr><tr><td>Spemann Pipette</td><td></td><td></td><td>Hand-made in lab</td></tr><tr><td>Egg Holder</td><td></td><td></td><td>Glass ashtray and modeling clay</td></tr><tr><td>Alcohol Burner</td><td>Fisher</td><td>04-245-1</td><td></td></tr><tr><td>Transparent Tape</td><td>3M Scotch</td><td>600</td><td></td></tr><tr><td>Glass Stirring Rod</td><td>Fisher</td><td>11-380C</td><td>Tip is narrowed and rounded using a flame</td></tr><tr><td>Tungston wire (0.004 x 3 inches)</td><td>A-M Systems</td><td>7190</td><td>Tip is flame-sharpened in a propane torch</td></tr><tr><td>Bunsun Burner</td><td>Fisher Scientific</td><td>S49117</td><td></td></tr><tr><td>Pasteur Pipette</td><td>Fisherbrand</td><td>22-183-632</td><td>9-inch (229 mm)</td></tr><tr><td>Rubber Tubing</td><td>Fisher Scientific</td><td>14-178C</td><td>amber, thin wall natural rubber; wall thickness: 0.0625 in/1.6 mm; O.D.: 0.375 in/9.5 mm; I.D.: 0.25 in/6.4 mm</td></tr><tr><td>Propane Fuel Cylinder</td><td>BernzOmatic</td><td>UL2317</td><td>TX-9 with torch style &quot;A&quot; with a screw-on brass &quot;pencil flame&quot; torch</td></tr><tr><td>Diamond Point Pencil</td><td>Fisher Scientific</td><td>22-268912</td><td></td></tr><tr><td>Rubber Bulbs</td><td>Fisherbrand</td><td>S32325</td><td></td></tr></tbody></table>

assessing species-specific contributions to craniofacial development using quail-duck chimeras

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

Prior to further analysis, the efficiency of the transplant needs to be assayed. For histological, morphological, or gene expression analyses on tissue samples, quail cells should be detected by immunohistochemistry using Q&#162;PN antibody as described36. For RNA analyses, species-specific contributions to tissues of interest can be calculated using a PCR-based strategy58. After the efficacy of the transplant has been validated, further morphological or molecular outcome measures can be evaluated in chimeras. Interactions between the donor neural crest cells and surrounding host-derived tissues that underlie proper histogenesis and morphogenesis of the craniofacial complex have previously been extensively studied. In particular, neural crest mesenchyme directs species-specific morphology of the face7,13,51,59, feather pattern52, muscle pattern56, and cartilage53,57 through the regulation of host gene expression.&#160;For example, neural crest mesenchyme dictates when bone forms in the mandible by temporally regulating Bmp4 expression55.
Recently, investigations have centered on tissue interactions occurring very early in craniofacial development. In this regard, experiments utilizing quail-duck chimeras have shown host embryos influence neural crest migration by determining morphological boundaries. That is, in chimeric quck embryos, donor quail neural crest cells migrate into the host duck mandibular arch in a duck-like pattern (Figure 1D). Despite this host contribution to the size of the neural crest population, the donor neural crest continues to generate a mandibular skeleton that is quail-like in size and shape (Figure 1E).
<img alt="Figure 1" fo:content-width="6in" fo:src="/files/ftp_upload/51534/51534fig1highres.jpg" src="/files/ftp_upload/51534/51534fig1.jpg" /> 
Figure 1.&#160;Quail-Duck Chimeric System. (A) Quail and (B) duck skulls display considerable differences in size and shape, and thus, are ideally suited for using a chimeric system to study craniofacial development (modified from Tokia&#160;et al.56).&#160;(C) Experimental design for generating unilateral chimeric quck embryos from stage-matched HH9.5 quail and duck embryos. The neural fold is removed from one side of quail embryos and transplanted into duck embryos after an equivalent piece of the neural fold has been removed.&#160;(D) Quail donor cells (green) can be followed in chimeras using an anti-quail antibody (Q&#162;PN) as shown in ventral view of HH12 chimeric embryo.&#160;(F) In quck mandibles at HH38, the quail donor-derived Meckel&#8217;s cartilage is shorter and straighter than that observed for the contralateral duck host-derived Meckel&#8217;s cartilage, which is larger and curved. Reprinted Tokita et al., Dev.&#160;Bio.&#160;306, 377 (2007) with permission from Elsevier.

Watch this Scientific Journal Video about Assessing Species-specific Contributions To Craniofacial Development Using Quail-duck Chimeras at JoVE.com

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

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

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

assessing-species-specific-contributions-to-craniofacial-development

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

10.9K Views.  University of California at San Francisco. This article describes a method to generate chimeric embryos that are designed to test the species-specific contributions of neural crest and/or other tissues to craniofacial development. 

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

Visualization of Chondrocyte Intercalation and Directional Proliferation via Zebrabow Clonal Cell Analysis in the Embryonic Meckel&#8217;s Cartilage

Development of the vertebrate craniofacial structures requires precise coordination of cell migration, proliferation, adhesion and differentiation. Patterning of the Meckel&#39;s cartilage, a first pharyngeal arch derivative, involves the migration of cranial neural crest (CNC) cells and the progressive partitioning, proliferation and organization of differentiated chondrocytes. Several studies have described CNC migration during lower jaw&#160;morphogenesis, but the details of how the chondrocytes achieve organization in the growth and extension of Meckel&#8217;s cartilage remains unclear. The sox10 restricted and chemically induced Cre recombinase-mediated recombination generates permutations of distinct fluorescent proteins (RFP, YFP and CFP), thereby creating a multi-spectral labeling of progenitor cells and their progeny, reflecting distinct clonal populations. Using confocal time-lapse photography, it is possible to observe the chondrocytes behavior during the development of the zebrafish Meckel&#8217;s cartilage.
Multispectral cell labeling enables scientists to demonstrate extension of the Meckel&#8217;s chondrocytes.&#160;During extension phase of the Meckel&#8217;s cartilage, which prefigures the mandible, chondrocytes intercalate to effect extension as they stack in an organized single-cell layered row. Failure of this organized intercalating process to mediate cell extension provides the cellular mechanistic explanation for hypoplastic mandible that we observe in mandibular malformations.

Cell organization of craniofacial bones has long been hypothesized but never directly visualized. Multi-spectral cell labeling and in vivo live imaging allows visualization of dynamic cell behavior in zebrafish lower jaw. Here, we detail the protocol to manipulate Zebrabow transgenic fish and directly observe cell intercalation and morphological changes of chondrocytes in the Meckel&#8217;s cartilage.

Development of the vertebrate craniofacial structures requires precise coordination of cell migration, proliferation, adhesion and differentiation. ...

Developmental Biology

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

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

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

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

Neuroscience

Two-step Approach to Explore Early- and Late-stages of Organ Formation in the Avian Model: The Thymus and Parathyroid Glands Organogenesis Paradigm

The avian embryo, as an experimental model, has been of utmost importance for seminal discoveries in developmental biology. Among several approaches, the formation of quail-chicken chimeras and the use of the chorioallantoic membrane (CAM) to sustain the development of ectopic tissues date back to the last century. Nowadays, the combination of these classical techniques with recent in vitro methodologies offers novel prospects to further explore organ formation.
Here we describe a two-step approach to study early- and late-stages of organogenesis. Briefly, the embryonic region containing the presumptive territory of the organ is isolated from quail embryos and grown in vitro in an organotypic system (up to 48 h). Cultured tissues are subsequently grafted onto the CAM of a chicken embryo. After 10 days of in ovo development, fully formed organs are obtained from grafted tissues. This method also allows the modulation of signaling pathways by the regular administration of pharmacological agents and tissue genetic manipulation throughout in vitro and in ovo developmental steps. Additionally, developing tissues can be collected at any time-window to analyze their gene-expression profile (using quantitative PCR (qPCR), microarrays, etc.) and morphology (assessed with conventional histology and immunochemistry).
The described experimental procedure can be used as a tool to follow organ formation outside the avian embryo, from the early stages of organogenesis to fully formed and functional organs.

This article provides an updated approach to the classical quail-chicken chimera system to study organ formation, by combining novel in vitro and in ovo experimental procedures.

The avian embryo, as an experimental model, has been of utmost importance for seminal discoveries in developmental biology. Among several approaches, the ...

Isolation of Embryonic Tissues and Formation of Quail-Chicken Chimeric Organs Using The Thymus Example

The capacity to isolate embryonic tissues was an essential step for establishing the quail-chicken chimera system, which in turn has provided undisputed contributions to unveiling key processes in developmental biology.
Herein is described an optimized method to isolate embryonic tissues from quail and chickens by microsurgery and enzymatic digestion while preserving its biological properties. After isolation, tissues from both species are associated in an in vitro organotypic assay for 48 h. Quail and chicken tissues can be discriminated by distinct nuclear features and molecular markers allowing the study of the cellular cross-talk between heterospecific association of tissues. This approach is, therefore, a useful tool for studying complex tissue interactions in developmental processes with highly dynamic spatial modifications, such as those occurring during pharyngeal morphogenesis and the formation of the foregut endoderm-derived organs. This experimental approach was first developed to study the epithelial-mesenchymal interactions during early-stages of thymus formation. In this, the endoderm-derived prospective thymic rudiment and mesoderm-derived mesenchyme, were isolated from quail and chicken embryos, respectively.
The capacity of the associated tissues to generate organs can be further tested by grafting them onto the chorioallantoic membrane (CAM) of a chicken embryo. The CAM provides nutrients and allows gas exchanges to the explanted tissues. After 10 days of in ovo development, the chimeric organs can be analyzed in the harvested explants by conventional morphological methods. This procedure also allows studying tissue-specific contributions during organ formation, from its initial development (in vitro development) to the final stages of organogenesis (in ovo development).
Finally, the improved isolation method also provides three-dimensionally (3D) preserved embryonic tissues, that can also be used for high-resolution topographical analysis of tissue-specific gene-expression patterns.

This article provides a method to isolate pure embryonic tissues from quail and chicken embryos that can be combined to form ex vivo chimeric organs.

The capacity to isolate embryonic tissues was an essential step for establishing the quail-chicken chimera system, which in turn has provided undisputed ...

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 ...

A Label-free Xenograft Model for Investigating the Behavior of Human Stem Cell Spheroids in Chick Embryos

Xenografts are valuable methods to investigate the behavior of human cells in vivo. In particular, the embryonic environment provides cues for cell migration, differentiation, and morphogenesis, with unique instructive signals and germ layer identity that are often absent from adult xenograft models. In addition, embryonic models cannot discriminate self versus non-self tissues, eliminating the risk of rejection of the graft and the need for immune suppression of the host. This paper presents a methodology for transplantation of spheroids of human cells into chicken embryos, which are accessible, amenable to manipulation, and develop at 37 &#176;C. 
Spheroids allow the selection of a specific region of the embryo for transplantation. After being grafted, the cells become integrated into the host tissue, allowing the follow-up of their migration, growth, and differentiation. This model is flexible enough to allow the utilization of different adherent populations, including heterogeneous primary cell populations and cancer cells. To circumvent the need for prior cell labeling, a protocol for the identification of donor cells through hybridization of human-specific Alu probes is also described, which is particularly important when investigating heterogeneous cell populations. Furthermore, DNA probes can be easily adapted to identify other donor species. This protocol will describe the general methods for preparing spheroids, grafting into chicken embryos, fixing and processing tissue for paraffin sectioning, and finally identifying the human cells using DNA in situ hybridization. Suggested controls, examples of interpretation of results and various cell behaviors that can be assayed will be discussed in addition to the limitations of this method.

Here, we describe a method to transplant and identify human cell spheroids into chick embryos. This xenograft model uses the embryonic microenvironment as a source of instructive signals to assay cell migration, differentiation, and tropism and is especially suited for the study of primary and/or heterogeneous cell populations.

Xenografts are valuable methods to investigate the behavior of human cells in vivo. In particular, the embryonic environment provides cues for cell ...

Assessing Signaling Properties of Ectodermal Epithelia During Craniofacial Development

The accessibility of avian embryos has helped experimental embryologists understand the fates of cells during development and the role of tissue interactions that regulate patterning and morphogenesis of vertebrates (e.g., 1, 2, 3, 4). Here, we illustrate a method that exploits this accessibility to test the signaling and patterning properties of ectodermal tissues during facial development. In these experiments, we create quail-chick 5 or mouse-chick 6 chimeras by transplanting the surface cephalic ectoderm that covers the upper jaw from quail or mouse onto either the same region or an ectopic region of chick embryos. The use of quail as donor tissue for transplantation into chicks was developed to take advantage of a nucleolar marker present in quail but not chick cells, thus allowing investigators to distinguish host and donor tissues 7. Similarly, a repetitive element is present in the mouse genome and is expressed ubiquitously, which allows us to distinguish host and donor tissues in mouse-chick chimeras 8. The use of mouse ectoderm as donor tissue will greatly extend our understanding of these tissue interactions, because this will allow us to test the signaling properties of ectoderm derived from various mutant embryos.

This article describes a tissue transplantation technique that was designed to test the signaling and patterning properties of surface cephalic ectoderm during craniofacial development.

The accessibility of avian embryos has helped experimental embryologists understand the fates of cells during development and the role of tissue ...

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.

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.

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

Analyzing Craniofacial Morphogenesis in Zebrafish Using 4D Confocal Microscopy

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical clarity and amenability to genetic manipulation, the zebrafish embryo has become a popular model organism with which to perform time-lapse analysis of morphogenesis in living embryos. Confocal imaging of a live zebrafish embryo requires that a tissue of interest is persistently labeled with a fluorescent marker, such as a transgene or injected dye. The process demands that the embryo is anesthetized and held in place in such a way that healthy development proceeds normally. Parameters for imaging must be set to account for three-dimensional growth and to balance the demands of resolving individual cells while getting quick snapshots of development. Our results demonstrate the ability to perform long-term in vivo imaging of fluorescence-labeled zebrafish embryos and to detect varied tissue behaviors in the cranial neural crest that cause craniofacial abnormalities. Developmental delays caused by anesthesia and mounting are minimal, and embryos are unharmed by the process. Time-lapse imaged embryos can be returned to liquid medium and subsequently imaged or fixed at later points in development. With an increasing abundance of transgenic zebrafish lines and well-characterized fate mapping and transplantation techniques, imaging any desired tissue is possible. As such, time-lapse in vivo imaging combines powerfully with zebrafish genetic methods, including analyses of mutant and microinjected embryos.

Time-lapse confocal imaging is a powerful technique useful for characterizing embryonic development. Here, we describe the methodology and characterize craniofacial morphogenesis in wild-type, as well as pdgfra, smad5, and smo mutant embryos.

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical ...

Dissection and Downstream Analysis of Zebra Finch Embryos at Early Stages of Development

The zebra finch (Taeniopygiaguttata) has become an increasingly important model organism in many areas of research including toxicology1,2, behavior3, and memory and learning4,5,6. As the only songbird with a sequenced genome, the zebra finch has great potential for use in developmental studies; however, the early stages of zebra finch development have not been well studied. Lack of research in zebra finch development can be attributed to the difficulty of dissecting the small egg and embryo. The following dissection method minimizes embryonic tissue damage, which allows for investigation of morphology and gene expression at all stages of embryonic development. This permits both bright field and fluorescence quality imaging of embryos, use in molecular procedures such as in situ hybridization (ISH), cell proliferation assays, and RNA extraction for quantitative assays such as quantitative real-time PCR (qtRT-PCR).&#160;This technique allows investigators to study early stages of development that were previously difficult to access.

The zebra finch (Taeniopygiaguttata) is a valuable model organism; however, early stages of zebra finch development have not been extensively studied. The protocol describes how to dissect early embryos for developmental and molecular applications.

The zebra finch (Taeniopygiaguttata) has become an increasingly important model organism in many areas of research including toxicology1,2, behavior3, and ...

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

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Visualization of Chondrocyte Intercalation and Directional Proliferation via Zebrabow Clonal Cell Analysis in the Embryonic Meckel’s Cartilage

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

Two-step Approach to Explore Early- and Late-stages of Organ Formation in the Avian Model: The Thymus and Parathyroid Glands Organogenesis Paradigm

Isolation of Embryonic Tissues and Formation of Quail-Chicken Chimeric Organs Using The Thymus Example

Creating Avian Forebrain Chimeras to Assess Facial Development

A Label-free Xenograft Model for Investigating the Behavior of Human Stem Cell Spheroids in Chick Embryos

Assessing Signaling Properties of Ectodermal Epithelia During Craniofacial Development

Facial Transplants in Xenopus laevis Embryos

Analyzing Craniofacial Morphogenesis in Zebrafish Using 4D Confocal Microscopy

Dissection and Downstream Analysis of Zebra Finch Embryos at Early Stages of Development