Fertilised GFP chicken eggs were supplied by Prof. Helen Sang, The Roslin Institute, and University of Edinburgh, UK. The Roslin Transgenic Chicken Facility is funded by the Wellcome Trust and by the Biotechnology and Biological Sciences Research Council (BBSRC). The work was partly funded, and NT supported, by Great Ormond Street Hospital Children&#8217;s Charity, London, UK.&#160;The authors thank Ben Jevans, UCL Institute of Child Health, for help with preparing embryos for grafting.

1. Preparation of Micro-scalpel for Neural Tube Ablations
<ol>
	<li>Shape a micro-scalpel from a commercially available steel sewing needle.
	<ol>
		<li>First flatten the needle on both sides using a grinding wheel mounted on a powered bench grinder.</li>
		<li>Start shaping the scalpel, first on a coarse grade Arkansas stone using a controlled circular motion, in alternate directions, on both sides of the needle.</li>
		<li>Carry on the same sharpening movements on an extra fine grade Arkansas stone to shape an ultra fine micro-scalpel, with a well defined cutting edge (Figure 1A, B). 
		NOTE: Alternatives to the micro-scalpel could be electrolitically sharpened needles, commercially available tungsten needles, or pulled glass needles.</li>
	</ol>
	</li>
</ol>
2. Incubate Wild Type and GFP Eggs to the Desired Stage
<ol>
	<li>Store fertilized chicken eggs and transgenic GFP chicken eggs in a cooled incubator at 14 - 15 &#176;C prior to incubation as development is halted at this temperature. Store eggs for a few days, up to one week.</li>
	<li>To commence development, place wild type and GFP eggs on a tray horizontally and incubate simultaneously in a humidified (58 - 60%) incubator at 37.5 &#186;C, so that embryos are at matching stages for neural tube grafting.</li>
	<li>To obtain embryos at the 10 - 12 somite stage of development for carrying out vagal neural tube grafting, incubate eggs for 1.5 days (33 - 38 hrs) and stage embryos according to the development tables of Hamburger and Hamilton21.</li>
</ol>
3. Prepare Eggs for Windowing and Grafting
<ol>
	<li>Move one egg at a time to a custom-made egg holder for windowing. Make a small hole in the egg shell by repeatedly tapping, with straight scissors, on the upper surface of the pointed end of the egg.</li>
	<li>Remove 2 - 3 ml of albumin from the egg with an 18&#189; G hypodermic needle and 5 ml syringe. Removing the albumin lowers the yolk within the egg and facilitates subsequent windowing without causing any damage to the embryo.
	<ol>
		<li>Discard the albumin. Seal the hole with a small strip of clear tape cut to size with fine scissors.</li>
	</ol>
	</li>
	<li>Using curved scissors, tap another hole in the upper surface of the egg shell. Insert the tip of the scissors into the hole and, keeping the scissors parallel to the bench, work in a circular motion to cut a ~2 cm diameter window on top of the shell.
	<ol>
		<li>Keep the scissors in a stationary position and rotate the egg. Discard the removed disk of egg shell. At E1.5, the embryo is recognizable as a darker yellow disk on top of the yolk.</li>
		<li>Remove any shell debris that has fallen within the egg using tweezers. Discard any unfertilized eggs (identified by a small white spot on top on the otherwise light yellow yolk).</li>
	</ol>
	</li>
</ol>
4. Prepare the Host Embryo to Receive Grafted Tissue
<ol>
	<li>Adjust the stereo-microscope to eye level and optimize the orientation of gooseneck light source to adequately illuminate the embryo without causing reflections.</li>
	<li>To visualize the embryo proper, inject a small amount of Indian ink under the center of the darker yellow disk, using a mouth tube and a pulled glass micro-pipette (Figure 1C, Oii).
	<ol>
		<li>Prepare the ink 50:50 with PBS containing Penicillin/Streptomycin at 100 &#956;g/ml final concentration. Insert the micro-pipette through the yolk membrane outside the perimeter of the blastoderm then carefully angle its tip directly beneath the embryo.</li>
		<li>Deliver ink underneath the embryo by blowing on the mouth tube. If mouth pipetting is not permitted, use a 1 ml syringe instead. Be careful not to introduce any air bubbles underneath the embryo, which can lead to contamination, then carefully remove the glass micro-pipette. This is a delicate step that can lead to the death of the embryo if not done with precision.</li>
	</ol>
	</li>
	<li>Stage the embryo by reference to Hamburger and Hamilton21 and record the stage in a lab book.</li>
	<li>Using a custom made micro-scalpel (or a fine tungsten needle) mounted on a needle holder, make a very small gash in the vitelline membrane, next to the area where the micro-surgery will be performed.
	<ol>
		<li>Carefully apply 2 - 3 drops of PBS over the membrane tear (using a glass micro-pipette and the mouth tube) to create space between the embryo and the membrane. Cut a larger window in the membrane to expose the entire region where the micro-surgery will take place.</li>
	</ol>
	</li>
	<li>Remove the neural tube region of interest using the micro-scalpel, starting with rostral and caudal transverse incisions across the entire dorsal neural tube (at the level of somite 1 to 7 in the video).
	<ol>
		<li>Cut bilaterally between the neural tube and the somites to separate the neural tube from surrounding tissues, without damaging the somites.</li>
		<li>Very gently separate the neural tube from the underlying notochord, which should remain intact. Note that successful neural tube excision will leave all surrounding tissues perfectly intact (Figure 2).</li>
		<li>Remove the excised neural tube by aspirating it into a glass micro-pipette, then discard.</li>
	</ol>
	</li>
	<li>Record the level of neural tube ablation in a lab book. The host embryo is now ready to receive the donor neural tube.</li>
</ol>
5. Prepare the Donor Graft Tissue
<ol>
	<li>Select a windowed, stage-matched GFP embryo by viewing under a fluorescent stereo-microscope with FITC filter. The GFP fluorescence makes it very easy to visualize the somites and stage the embryo.</li>
	<li>Once a stage-matched embryo has been identified, remove the embryo from the egg by making 4 incisions, with Pascheff-Wolff spring scissors (Figure 1C, l) in a rectangle shape around the embryo then gently pick it up with an embryo spoon.</li>
	<li>Place the embryo in a square watch glass with a sylgard polymer base. Gently shake the embryo with Dumont #5 tweezers to remove any attached yolk. Remove the vitelline membrane and pin the embryo onto the polymer base using stainless minutien pins (Figure 1C).</li>
	<li>Using the Spring Scissors, make 4 incisions in a rectangular shape around the neural tube and surrounding somites, in the same region that has been removed from the host embryo.</li>
	<li>Using a plastic transfer pipette, transfer the neural tube and somite tissues from the donor GFP embryo into a watch glass containing 0.2% pancreatin in Pen/Strep PBS.</li>
	<li>Allow enzymatic digestion to proceed for 10 min at RT to help separate the tissues. After incubation in enzyme, use stainless minutien pins mounted on a handle to manually separate the neural tube from all adjacent tissues.</li>
	<li>Using a glass micro-pipette, transfer the dissociated neural tube to another watch glass containing DMEM + 10% serum (e.g., goat, horse or fetal calf) on ice, to rinse the excess pancreatin and stop enzymatic digestion. After 5 min, the dissected neural tube is ready to be grafted orthotopically into the chick host (Figure 2 and S1).</li>
</ol>
6. Graft the Tissue
<ol>
	<li>Using a glass micro-pipette, carefully transfer the dissected neural tube from the watch glass to the host embryo. Position the neural tube in the correct anterior-posterior orientation and gently push the explant adjacent to the excised region of the chick host using the micro-scalpel. Leave a small fragment of ectoderm attached to, or by cutting a small nick in, the dorsal surface to identify the orientation of the neural tube.
	<ol>
		<li>If necessary, use the micro-scalpel to trim the explant to the exact size of the excised region.</li>
	</ol>
	</li>
	<li>Gently guide the neural tube into the ablated region and position it such that the dorsal side is correctly orientated. Use a glass micro-pipette, mounted to a mouth tube, to remove PBS and/or fluid surrounding the graft. This helps the donor and host tissues to adhere and the graft to become established.</li>
	<li>Seal the entire window with 24 mm wide clear tape to prevent dehydration and contamination.</li>
	<li>Label the chimeric embryo by marking with a pencil on the eggshell and record its number in the lab book. Return the egg to the incubator for further development.</li>
</ol>
7. Inject DiI into Blood Vessels of Host Embryo
<ol>
	<li>At the desired experimental time point (here, 3 - 10 days later), retrieve the chimeric embryo from the incubator and remove the clear tape using straight scissors to gain access to the embryo within the egg.</li>
	<li>If necessary, enlarge the window in the shell using the scissors. Be careful not to damage the chorioallantoic membrane if it is attached to the shell, which would result in hemorrhage and jeopardize the blood vessel labeling.</li>
	<li>Choose an accessible vein on the yolk making sure the blood flow is directed towards the embryo. Choose a branching point of one of the vitelline veins (Figure 3B, C). 
	NOTE: At E6.5 - E7.5, the chorioallantoic membrane may need to be gently moved aside with tweezers to access the yolk veins. After E8.5, the only option is to inject into one of the chorioallantoic membrane veins since, by this stage, the chorioallantoic membrane fully covers the embryo.</li>
	<li>Remove the vitelline membrane above the chosen injection point using two Dumont #5 tweezers by tearing in opposite directions.</li>
	<li>Break a pulled glass needle using a Dumont #5 and adjust its diameter to the approximate size of the vein prior to loading with CellTracker CM-DiI. Make the DiI stock solution at 40 &#181;g/&#181;l in DMSO and store at -20 &#176;C. Prepare the working solution in 0.3 M sucrose/PBS at a concentration of 4 &#181;g/&#181;l.
	<ol>
		<li>Aspirate between 5 - 10 &#181;l of DiI in 0.3 M sucrose/PBS into the needle using suction with a mouth tube. Older embryos might require up to 25 &#181;l or more. From E8.5, embryos have larger, more muscular veins, which may need to be held in position with a Dumont #5 before stabbing with the DiI-loaded glass needle.</li>
	</ol>
	</li>
	<li>Swiftly insert the needle into the vein and blow steadily with the mouth tube to allow the DiI to join the blood flow slowly without forming a clot. Alternatively, use a pressure injector for DiI delivery.</li>
</ol>
8. Harvest Embryos for Sectioning or Wholemount Examination
<ol>
	<li>To retain as much DiI within the embryo as possible, harvest the embryo immediately after the injection by scooping it onto a perforated spoon and cutting the blood vessels and connective tissues with a pair of straight scissors, to free the embryo from the yolk.</li>
	<li>Remove any loose membranes and dissect out the organs of interest (i.e., the lungs and digestive tract in this tutorial), taking great care not to compress the tissue, which creates diffusion of the DiI. Immediately fix the tissues by immersion in 4% PFA for 1 - 2 hrs at RT.</li>
	<li>Rinse the tissue for 5 min in PBS, then 15 min in PBS containing 5 &#956;g/ml DAPI. Mount the samples on a bridged microscope slide for whole mount examination or embed them for cryo-sectioning.</li>
</ol>

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The authors declare that they have no competing financial interests.

The method of intraspecies neural tube grafting, combined with blood vessel labeling described here, takes full advantage of the ease of access of the avian embryo within the egg (compared with other vertebrate embryos) to study the co-development of an element of the autonomic nervous system (the ENS) and the vascular system.
For labeling NCC derivatives, the chickGFP-chick intraspecies grafting method we describe has a number of advantages over the classical quail-chick chimera method that was established over 40 years ago1-3. First, under FITC light, GFP fluorescence is extremely bright, to the extent that GFP+ cells are easily discernable in live chimeric embryos. This allows the success of the graft to be checked in ovo, whereas quail-chick grafting requires the embryo to be killed, processed and immunostained using QCPN, before the success of the graft can be ascertained2. Second, GFP expression in the transgenic chickGFP is cytoplasmic, therefore it not only labels cell bodies, but also allows the projections of the transplanted cells to be visualized22. This permits intricate neuronal networks to be observed at high resolution (note that fine projections are best visualized when the sample is immunostained with anti-GFP antibody). As QCPN labeling is restricted to the quail cell nucleus, such networks are not revealed using quail-chick chimeras. Third, intraspecies grafting eliminates any potential species differences between cells within the chimeric embryo. Since quail embryos have a shorter incubation period than chick (19 days versus 21 days) it has been suggested that quail cells have a higher proliferation rate than chick cells, which could potentially affect the development of the chimeric tissues23. Interestingly also, it has been shown in plants that interspecies grafting could produce extensive alterations in DNA methylation patterns in the host 24. Fourth, chickGFP facilitates back-transplant experiments to address topics such as NCC fate and cell commitment25. Fifth, the transgenic chickGFP is also useful for many other techniques including FACS sorting of GFP+ cell subpopulations, organotypic culture of organs containing GFP+ cells, genetic manipulation of GFP+ grafted tissue via electroporation of expression plasmids26, and other imaging technologies such as optical projection tomography27.
The neural tube transplantation approach can be modified by microsurgically replacing shorter amounts of neural tube. By using smaller segments of neural tube the microsurgery is potentially less damaging to the embryo and survival can be improved. However, the downside of transplanting less neural tube is that the numbers of GFP+ NCC in the host will be reduced. Users could try and achieve a balance between the amount of neural tube transplanted to give optimum survival of embryos, and the numbers of GFP+ NCC within host intestine sufficient to give informative results.
For vessel painting, DiI has the advantage that its fluorescence is very bright and robust. Also, it has the capacity to diffuse during fixation assuring staining of the finest opened capillaries. Since it is a vital dye, embryos can survive the injection procedure and carry on developing with a stained vascular system (up to 24 hr in our hands, although the staining becomes more punctate over time, see Figure 3E). The combination of chickGFP grafting with DiI vascular painting is therefore compatible with live imaging. Besides all these advantages, it is important to note that vascular injection only labels luminised vessels and therefore does not identify unopened capillaries, endothelial tip cells or isolated endothelial cells. However, further progress in avian transgenesis could provide new ways to circumvent such issues, as exemplified by experiments using Tg(tie1:H2B-eYFP) quail embryos to study vascular morphogenesis28. Another limitation of this technique is that, for effective vessel labeling in embryos at E7.5 and beyond, larger amounts of dye need to be injected, which can make experiments expensive. However, a modification of the technique could include low cost blood vessel labelingusing highlighter ink14, although this approach has not been tried in our hands.
Critical steps of the procedures include the process of visualizing the embryo by injecting ink underneath the blastodisc. If the membrane covering the yolk is torn by the ink-filled needle at this stage then embryo survival is severely compromised. Also, it is important, when preparing a donor neural tube, that the tissue is not left for an excessively long time in pancreatin (consider approximately 10 min&#160;as a maximum). Prolonged exposure to pancreatin damages the tissue and the neural tube is then difficult to handle and it will not incorporate well into the host.&#160;Gaining experience of the DiI injection technique on wild type embryos is essential before injecting chimeric embryos, as only one attempt at injecting is generally possible for each embryo. DiI volume and needle diameter are critical parameters for each embryo and should be assessed on wild type, stage matched controls. &#160;
In conclusion, our dual labeling method of neural tube transplantation and DiI vessel painting in live chick embryos can be used to investigate the inter-relationships between NCC and blood vessel networks during organogenesis. Considering the mechanisms responsible for establishing correct target innervation and vascularization during organ development are still largely unknown, this methodology holds potential for future discoveries in this field.

The chicken embryo is an invaluable model organism in vertebrate developmental biology, not least because its development in ovo permits experimental manipulations that are otherwise impossible to perform in vertebrates that develop in utero. This accessibility and ease of manipulations has led to the chick embryo playing key roles in many seminal discoveries in the field of developmental biology. Amongst the most powerful techniques has been the use of quail-chick chimeric embryos to study cell fate, a method pioneered by Professor Nicole Le Douarin in the 1970&#8217;s1-3. In particular, quail-chick chimeras have been especially useful to genetically mark and follow highly migratory neural crest cell (NCC) populations during early development. NCC are a multipotent population of migratory cells, arising in the dorsal ectoderm at the margins of the neural tube, that give rise to a wide range of cell types throughout the vertebrate embryo. These include craniofacial structures (cartilage, bone, muscles), neurons and glia (in the sensory and autonomic nervous systems), melanocytes, and a subpopulation of cells of the endocrine system2,4,5. One of the most important factors influencing NCC fate is their initial location along the anterior-posterior axis of the neural tube. For example, enteric NCC, which give rise to the neurons and glia of the enteric nervous system (ENS), arise from two discrete sub-populations: the first located in the vagal (caudal hindbrain) region, and the second in the sacral region of the neural tube6-13. Inter or intra-species grafting of the corresponding regions of the neural tube have been the techniques of choice to permanently label these cells and subsequently allow tracking, from their birth on the margins of the neural tube, to their final destinations within the digestive tract6,7,10.
Another embryonic manipulation easier to perform in chick, compared to other animal models, is the vital labeling of the vascular system. Indeed, as the chick embryo develops, it lays on top of an extra-embryonic vascular network that circulates oxygen and nutrients from the yolk. This accessible vascular network, located on the surface of the yolk, can be used as a gateway to label the embryo&#8217;s developing vascular system during organogenesis12,14-17. Intravascular injection of various dyes, such as the lipophilic dye DiI, makes it possible to delineate/stain all the luminised vessels of the nascent vascular network.
Because developing organs need to be connected to both the nervous system (for sensory and motor control) as well as the vascular system (for gas exchange, fluid and nutrient supply), the two networks develop alongside each other and share striking similarities in their branching architecture18-20. Here we report embryonic manipulations that allow us to study the simultaneous development of the NCC-derived ENS, along with the vascular system, during organogenesis. This is achieved by generating chicken chimeras via transplantation of discrete segments of the neural tube, including the neural crest, combined with vascular DiI injection. As an advance from quail-chicken chimeras, our method uses transgenic GFP chick embryos for intraspecies grafting, making the transplant technique more powerful, in terms of imaging cells and their projections, and eliminating any potential bias linked to species differences.

<table><tbody><tr><td>Fertilised chick eggs</td><td>Henry Stewart and Co, Louth, UK</td><td></td><td></td></tr><tr><td>Fertilised GFP chick eggs</td><td>The Transgenic Chicken Facility, The Roslin Institute, The University of Edinburgh</td><td></td><td></td></tr><tr><td>Egg incubator (Profi-H Hatcher)</td><td>Lyon Technologies, CA, USA</td><td>910-033</td><td></td></tr><tr><td>14C Incubator</td><td>Precision Cooled Incubator, Leec Ltd., Nottingham, UK</td><td>Model LT2</td><td></td></tr><tr><td>Stereo-microscope&amp;nbsp;</td><td>LEICA</td><td>Model MZ 12.5</td><td></td></tr><tr><td>Digital Camera</td><td>LEICA</td><td>DC500</td><td></td></tr><tr><td>Image acquisition software</td><td>LEICA</td><td>IM50</td><td></td></tr><tr><td>Goose neck halogen cold light source</td><td>Advanced Imaging Concepts, Inc</td><td>KL 1500 LCD&amp;nbsp;</td><td></td></tr><tr><td>181&amp;frasl;2 G hypodermic needle&amp;nbsp;</td><td>SIGMA - ALDRICH</td><td>HSWNH181</td><td></td></tr><tr><td>Pancreatin</td><td>SIGMA - ALDRICH</td><td>P3292</td><td></td></tr><tr><td>DMEM</td><td>SIGMA - ALDRICH</td><td>D5030</td><td></td></tr><tr><td>Goat serum</td><td>SIGMA - ALDRICH</td><td>G6767</td><td></td></tr><tr><td>5 ml syringe</td><td>SIGMA - ALDRICH</td><td>Z248010</td><td></td></tr><tr><td>Mouth tube</td><td>SIGMA - ALDRICH</td><td>A5177</td><td></td></tr><tr><td>Sigma Pasteur pipettes non-plugged, L 5 3/4 in.</td><td>SIGMA - ALDRICH</td><td>S6018</td><td></td></tr><tr><td>Transfer pipettes, polyethylene</td><td>SIGMA - ALDRICH</td><td>Z350796&amp;nbsp;</td><td></td></tr><tr><td>Borosillicate glass capillaries, thin wall without filament</td><td>Harvard apparatus</td><td>PY8 30-0035</td><td></td></tr><tr><td>Iris Scissors - ToughCut</td><td>Fine Science Tools</td><td>14058-09</td><td></td></tr><tr><td>Curved Iris Scissors - ToughCut</td><td>Fine Science Tools</td><td>14059-09</td><td></td></tr><tr><td>Needle holders (Nickel-plated pin holder)&amp;nbsp;</td><td>Fine Science Tools</td><td>26018-17&amp;nbsp;</td><td></td></tr><tr><td>Pascheff-Wolff Spring Scissors&amp;nbsp;</td><td>Fine Science Tools&amp;nbsp;</td><td>15371-92</td><td></td></tr><tr><td>Dumont #5 forceps&amp;nbsp;</td><td>Fine Science Tools&amp;nbsp;</td><td>11251-30&amp;nbsp;</td><td></td></tr><tr><td>Minutien pins&amp;nbsp;</td><td>Fine Science Tools</td><td>26002-15</td><td></td></tr><tr><td>Dumont AA forceps, Inox Epoxy- coated&amp;nbsp;</td><td>Fine Science Tools</td><td>11210-10&amp;nbsp;</td><td></td></tr><tr><td>Perforated spoon</td><td>Fine Science Tools</td><td>10370-18</td><td></td></tr><tr><td>Tungsten needles (0.125mm diameter)</td><td>Fine Science Tools</td><td>10130-05</td><td></td></tr><tr><td>Sellotape (clear, 24 mm width)</td><td>Any Supplier</td><td></td><td></td></tr><tr><td>Pen/Strep (Penicillin, Streptomycin) Solution</td><td>VWR international</td><td>101447-068&amp;nbsp;</td><td></td></tr><tr><td>Sylgard 184 silicone elastomer kit</td><td>Dow Corning</td><td>S09 512 516</td><td></td></tr><tr><td>Pelikan black ink</td><td>Pelikan</td><td>211-169</td><td></td></tr><tr><td>CellTracker CM-DiI&amp;nbsp;</td><td>Molecular Probes</td><td>C-7001</td><td></td></tr><tr><td>DAPI (4&#039;,6-Diamidino-2-Phenylindole, Dihydrochloride)</td><td>Molecular Probes</td><td>D1306</td><td></td></tr><tr><td>Settings for glass needle puller&amp;nbsp;</td><td>Sutter Instruments</td><td>Flaming/Brown micropipette puller model P-86</td><td></td></tr><tr><td>Heat 950; Pull 150; Velocity 100; Time 200; Pressure 500</td><td></td><td></td><td></td></tr></tbody></table>

dual labeling of neural crest cells and blood vessels within chicken embryos using chickgfp neural tube grafting and carbocyanine dye dii injection

All developing organs need to be connected to both the nervous system (for sensory and motor control) as well as the vascular system (for gas exchange, fluid and nutrient supply). Consequently both the nervous and vascular systems develop alongside each other and share striking similarities in their branching architecture. Here we report embryonic manipulations that allow us to study the simultaneous development of neural crest-derived nervous tissue (in this case the enteric nervous system), and the vascular system. This is achieved by generating chicken chimeras via transplantation of discrete segments of the neural tube, and associated neural crest, combined with vascular DiI injection in the same embryo. Our method uses transgenic chickGFP embryos for intraspecies grafting, making the transplant technique more powerful than the classical quail-chick interspecies grafting protocol used with great effect since the 1970s. ChickGFP-chick intraspecies grafting facilitates imaging of transplanted cells and their projections in intact tissues, and eliminates any potential bias in cell development linked to species differences. This method takes full advantage of the ease of access of the avian embryo (compared with other vertebrate embryos) to study the co-development of the enteric nervous system and the vascular system.

Figure 1 shows typical instruments required to carry out microsurgical isolation and transplantation of the neural tube. Figure 2 shows the transplant procedure. Following transplantation embryos are screened for transplant success. This involves examining the embryo under a stereo fluorescence microscope, typically the morning after microsurgery, for the presence of graft-derived (GFP+) NCC. If transplantation has been a success, then GFP+ NCC can be observed in the vicinity of the neural tube and in early migration pathways leading toward the foregut. If the procedure has not been successful, GFP+ NCC will not be observed outside the neural tube, or if they are present in the host they may be in smaller numbers. These unsuccessful embryos are discarded. Typically, 5-8 neural tube transplants are performed in one day, and of these 80% are successful. Reasons for unsuccessful neural tube transplantation include death of the embryo due to tissue damage incurred during microsurgery, or failure of the neural tube to integrate into the host embryo. The latter can result from poor placement of the neural tube within the host or from a poor quality neural tube due to poor dissection technique or from excessive exposure to dissociation enzyme. The initial screening step, as well as similar later examinations for GFP+ cells, is useful as it means that time and resources are not wasted by performing experiments on embryos that do not have GFP-labelled NCC within the gut.
Figure 3 shows the procedure for DiI injection of the blood vessels.&#160;The efficiency/success of the DiI injection technique depends on: first, cutting the injection needle to the optimum diameter for the targeted vein, second a precise gesture when inserting the needle in the vein (so not to pierce through the other side), and third avoiding the needle getting plugged during the injection by blowing at a constant rate. If any of these three parameters is done incorrectly, the embryo will bleed out or will need several hours to recover before a second attempt is made as the hemorrhage makes it almost impossible to re-inject immediately. Successful embryos should be selected immediately by viewing under a stereofluorescence microscope and must be dissected rapidly. In successful embryos, DiI labeled blood vessels are present throughout the embryo (Figure 3C,D) including capillary beds (Figure 3D).
Upon harvesting of embryos and examination of tissue sections or wholemount gastrointestinal tracts, typical results reveal GFP+ NCC within the primitive ENS and the fine structure of the DiI-labeled gut blood vessel networks (Figure 4) Wholemount preparations can be examined using confocal microscopy whereby image stacks produce three dimensional (3D) reconstructions showing the interrelationships between the fine projections of GFP+ ENS cells and the DiI stained vascular system (Figure 4 A-C; G-I; Videos 1 and 2).
<img alt="Figure 1" src="/files/ftp_upload/52514/52514fig1.jpg" /> 
Figure 1. Recommended microsurgery instruments.&#160;(A) micro-scalpel shaped from a sewing needle. (B) fine Arkansas stone for shaping a micro-scalpel. (C) a) straight scissors, b) curved scissors, c) 5 ml syringe with 181/2 G hypodermic needle, d) plastic pipette, e) custom made egg holder, f) black ink, g) square watch glass, h) square watch glass with black sylgard base, i) micro-scalpel on needle holder, j) minutien pins, k) minutien or tungsten needle on needle holder, l) Pascheff-Wolff spring scissors, m) Dumont #5 tweezers, n) perforated spoon, oi) short fire-pulled transfer needle, oii) long fire-pulled inking needle, p) mouth tube. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52514/52514fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/52514/52514fig2.jpg" /> 
Figure 2. Intraspecies neural tube transplant. Chick embryo/GFP neural tube images have been modified from Delalande et al.12. Vascularization is not necessary for gut colonization by enteric neural crest cells. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52514/52514fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/52514/52514fig3.jpg" /> 
Figure 3. Intravenous DiI injection. (A)&#160;Recommended instruments: a) CellTracker CM-DiI drop on parafilm, b) pulled glass injection needle, c) mouth tube. (B)&#160;Schematic diagram of intravenous DiI injection into E4 chimeric chick embryo. (C)&#160;in ovo DiI intravenous injection showing fine glass needle containing DiI inserted into vein (arrow). (D) E4 chimeric&#160;embryo post DiI injection (red) with GFP+ neural tube (arrow).&#160;(E) DiI stained fine blood vessel network in a live embryo, 24 hr post-injection. Br: brain; H: heart; LB: limb bud; A: allantois. Images in (C) and (D) have been modified from Delalande et al.12 Vascularization is not necessary for gut colonization by enteric neural crest cells. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52514/52514fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/52514/52514fig4.jpg" /> 
Figure 4: Representative results in the stomach and caecum of an E5.5 chick embryo. (A-C)&#160;3 dimensional (3D) reconstruction of a confocal image stack in the region of the stomach showing (D) the GFP+ enteric neural crest cells (ENCC) (E) the DiI stained vascular system and (F) a merged image of both networks D-F Histological sections at the level of the stomach showing (G) the GFP+ ENCC (H) the DiI stained vascular system and (I) a merged image of both networks. Nuclei are stained with DAPI (cyan).(G-H) 3D reconstruction of a confocal image stack in the caecum region showing (A) the GFP+ ENCC migration front in green, (B) the DiI stained vascular system in red, and (C) a merged image of both networks. Images (A-F) have been modified from Delalande et al.12 Vascularization is not necessary for gut colonization by enteric neural crest cells. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52514/52514fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure S1" src="/files/ftp_upload/52514/52514figS1.jpg" /> Figure S1.&#160;Isolation of a donor GFP+ neural tube from the surrounding tissues by enzymatic digestion and micro-dissection. (A)&#160;GFP+ neural tube and adjacent somites dissected from the donor embryo. (B) Isolated neural tube after pancreatin digestion and micro-dissection using stainless minutien pins. So: somites; NT: neural tube; Nc: Notochord.
<img alt="video 1" src="/files/ftp_upload/52514/52514video1.jpg" /> Video 1.&#160;3-dimensional 360&#176; rotation of the image in Figure 4C, showing the vascular system and the ENCC in the stomach at E5.5 (HH27-28). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52514/JoVE_Video1.avi" target="_blank">Please click here to view this video.</a>
<img alt="Video 2" src="/files/ftp_upload/52514/52514video2.jpg" /> Video 2.&#160;3-dimensional 360&#176; rotation of the image in Figure 4I, showing the vascular system and the ENCC migration front in the region of the caecum at E5.5 (HH27-28). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52514/JoVE_Video2.avi" target="_blank">Please click here to view this video.</a>

Watch this Scientific Journal Video about Dual Labeling of Neural Crest Cells and Blood Vessels Within Chicken Embryos Using ChickGFP Neural Tube Grafting and Carbocyanine Dye DiI Injection at JoVE.co

Dual Labeling of Neural Crest Cells and Blood Vessels Within Chicken Embryos Using ChickGFP Neural Tube Grafting and Carbocyanine Dye DiI Injection

Here we report dual labeling of neural crest cells and blood vessels using chickGFP neural tube intraspecies grafting combined with intra-vascular DiI injection. This experimental technique allows us to simultaneously visualize and study development of the NCC-derived (enteric) nervous system and the vascular system, during organogenesis.

Dual Labeling of Neural Crest Cells and Blood Vessels Within Chicken Embryos Using ChickGFP Neural Tube Grafting and Carbocyanine Dye DiI Injection

All developing organs need to be connected to both the nervous system (for sensory and motor control) as well as the vascular system (for gas exchange, ...

dual-labeling-neural-crest-cells-blood-vessels-within-chicken-embryos

Research

JoVE Journal

Developmental Biology

8.3K Views.  UCL Institute of Child Health. Hier melden we dubbele etikettering van neurale kuifcellen en bloedvaten met behulp van chickGFP neurale buis intraspecies enting in combinatie met intra-vasculaire DiI injectie. Deze experimentele techniek stelt ons in staat om tegelijkertijd de ontwikkeling van het NCC-afgeleide (enterische) zenuwstelsel en het vasculaire systeem te visualiseren en te bestuderen, tijdens organogenese. 

Video: Dual Labeling of Neural Crest Cells and Blood Vessels Within Chicken Embryos Using ChickGFP Neural Tube Grafting and Carbocyanine Dye DiI Injection

Double Whole Mount in situ Hybridization of Early Chick Embryos

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying  gene expression in brain, neural tube, somite and heart primordia formation. This video demonstrates the different steps in 2-color whole mount in situ hybridization; First, the embryo is dissected from the egg and fixed in paraformaldehyde. Second, the embryo is processed for prehybridization. The embryo is then hybridized with two different probes, one coupled to DIG, and one coupled to FITC.  Following overnight hybridization, the embryo is incubated with DIG coupled antibody. Color reaction for DIG substrate is performed, and the region of interest appears blue. The embryo is then incubated with FITC coupled antibody. The embryo is processed for color reaction with FITC, and the region of interest appears red. Finally, the embryo is fixed and processed for phtograph and sectioning. A troubleshooting guide is also presented.

This video demonstrates 2-color whole mount in situ hybridization, a method by which the spatial and temporal expression pattern of 2 different genes can be visualized in young chick embryos. This method was originally introduced by David Wilkinson, Domingos Henrique, Phil Ingham and David Ish -Horowicz.

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ...

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

Assay for Neural Induction in the Chick Embryo

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying the formation and initial patterning of the nervous system. This video demonstrates how to graft organizer tissue into a host, a method by which Hensen s node (the organizer in the chick embryo) is grafted to a host competent ectoderm. The organizer graft instructs overlying na ve tissue to adopt a neural fate via neural inducing signals. This mechanism is referred to as neural induction, and constitutes the initial step in the formation of brain and spinal cord in amniotes. This method is essentially used for the characterization of putative neural inducing molecules in chick. This video demonstrates the different steps in the assay for neural induction; First, the donnor embryo is explanted and pinned on a dish. Then, the host embryo is prepared for New culture. The graft is excised and transplanted to the host area pellucida margin. The host is cultured for 18-22 hrs. The assembly is fixed and processed for further applications (e.g. in situ hybridization). This method was originally devised by Waddington 1,2 and Gallera 3,4.

Neural induction is the first step in the formation of the brain. It is a mechanism by which Hensen's node (organizer), instructs adjacent tissue to adopt a neural fate, i.e. to give rise to the nervous system. This video demonstrates an assay for neural induction in chick embryo.

Electroporation of the Hindbrain to Trace Axonal Trajectories and Synaptic Targets in the Chick Embryo

Electroporation of the chick embryonic neural tube has many advantages such as being quick and efficient for the expression of foreign genes into neuronal cells. In this manuscript we provide a method that demonstrates uniquely how to electroporate DNA into the avian hindbrain at E2.75 in order to specifically label a subset of neuronal progenitors, and how to follow their axonal projections and synaptic targets at much advanced stages of development, up to E14.5. We have utilized novel genetic tools including specific enhancer elements, Cre/Lox - based plasmids and the PiggyBac-mediated DNA transposition system to drive GFP expression in a subtype of hindbrain cells (the dorsal most subgroup of interneurons, dA1). Axonal trajectories and targets of dA1 axons are followed at early and late embryonic stages at various brainstem regions. This strategy contributes advanced techniques for targeting cells of interest in the embryonic hindbrain and for tracing circuit formation at multiple stages of development.

How neuronal networks are established in the embryonic brain is a fundamental question in developmental neurobiology. Here we combined an electroporation technique with novel genetic tools, such as Cre/Lox&#x2013;plasmids and PiggyBac-mediated DNA transposition system in the avian hindbrain to label dorsal interneurons and track their axonal projections and synaptic targets at various developmental stages.

Electroporation of  the chick embryonic neural tube has many advantages such as being quick and efficient  for the expression of foreign genes into ...

Deciphering Axonal Pathways of Genetically Defined Groups of Neurons in the Chick Neural Tube Utilizing in ovo Electroporation

Employment of enhancer elements to drive expression of reporter genes in neurons is a widely used paradigm for tracking axonal projection. For tracking axonal projection of spinal interneurons in vertebrates, germ line-targeted reporter genes yield bilaterally symmetric labeling. Therefore, it is hard to distinguish between the ipsi- and contra-laterally projecting axons. Unilateral electroporation into the chick neural tube provides a useful means to restrict expression of a reporter gene to one side of the central nervous system, and to follow axonal projection on both sides 1 ,2-5. This video demonstrates first how to handle the eggs prior to injection. At HH stage 18-20, DNA is injected into the sacral level of the neural tube, then tungsten electrodes are placed parallel to the embryo and short electrical pulses are administered with a pulse generator. The egg is sealed with tape and placed back into an incubator for further development. Three days later (E6) the spinal cord is removed as an open book preparation from embryo, fixed, and processed for whole mount antibody staining. The stained spinal cord is mounted on slide and visualized using confocal microscopy.

Deciphering Axonal Pathways of Genetically Defined Groups of Neurons in the Chick Neural Tube Utilizing in ovo Electroporation

This video demonstrates how to visualize axonal pathways of genetically defined groups of neurons in the embryonic chick spinal cord utilizing in ovo electroporation of reporter genes under the control of specific enhancer elements.

Employment of enhancer elements to drive expression of reporter genes in neurons is a widely used paradigm for tracking axonal projection. For tracking ...

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 ...

Single Cell Transfection in Chick Embryos

A central theme in developmental biology is the diversification of lineages and the elucidation of underlying molecular mechanisms. This entails a thorough analysis of the fates of single cells under normal and experimental conditions. To this end, transfection methods that target single progenitors are a prerequisite. We describe here a technically straightforward method for transfecting single cells in chicken tissues in-ovo, allowing reliable lineage tracing as well as genetic manipulation. Specific tissue domains are targeted within the somite or neural tube, and DNA is injected directly into the epithelium of interest, resulting in sporadic transfection of single cells. Using reporters, clonal populations may consequently be traced for up to three days, and behavior of genetically manipulated clonal populations can be compared with that of controls. This method takes advantage of the accessibility of the chick embryo along with emerging tools for genetic manipulation. We compare and discuss its advantages over the widely-used electroporation method, and possible applications and use in additional in-vivo models are also suggested. We advocate the use of this method as a significant addition and complement for existing lineage tracing and genetic interference tools.

Using fine tip micropipettes we inject plasmid DNA into subdomains of chicken somites or neural tubes. The concentration of the plasmid is adjusted to generate single transfected cells. We then allow the cells to develop into clonal populations.

A central theme in developmental biology is the diversification of lineages and the elucidation of underlying molecular mechanisms. This entails a ...

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 ...

In Ovo Intravascular Injection in Chicken Embryos

As a classical model system of embryo biology, the chicken embryo has been used to investigate embryonic development and differentiation. Delivering exogenous materials into chicken embryos has a great advantage for studying gene function, transgenic breeding, and chimera preparation during embryonic development. Here we show the method of in ovo intravascular injection whereby exogenous materials such as plasmid vectors or modified primordial germ cells (PGCs) can be transferred into donor chicken embryos at early developmental stages. The results show that the intravascular injection through the dorsal aorta and head allows injected materials to diffuse into the whole embryo through the blood circulatory system. In the presented protocol, the efficacy of exogenous plasmid and lentiviral vector introduction, and the colonization of injected exogenous PGCs in the recipient gonad, were determined by observing fluorescence in the embryos. This article describes detailed procedures of this method, thereby providing an excellent approach to studying gene function, embryo and developmental biology, and gonad-chimeric chicken production. In conclusion, this article will allow researchers to perform in ovo intravascular injection of exogenous materials into chicken embryos with great success and reproducibility.

In Ovo Intravascular Injection in Chicken Embryos

The overall goal of this paper is to describe how to perform in ovo intracellular injection of exogenous materials into chicken embryos. This approach is very useful to study the developmental biology of chicken embryos.

As a classical model system of embryo biology, the chicken embryo has been used to investigate embryonic development and differentiation. Delivering ...

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.

Dual Labeling of Neural Crest Cells and Blood Vessels Within Chicken Embryos Using Chick^GFPNeural Tube Grafting and Carbocyanine Dye DiI Injection

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