We thank George Galaris (LUMC, Leiden, the Netherlands) for his help with chicken embryo injection. We acknowledge the support of Saskia van der Wal-Maas (Department of Anatomy &#38; Embryology, LUMC, Leiden, the Netherlands), Conny van Munsteren (Department of Anatomy &#38; Embryology, LUMC, Leiden, the Netherlands), Manon Zuurmond (LUMC, Leiden, the Netherlands), and Annemarie de Graaf (LUMC, Leiden, the Netherlands). M. Koning is supported by &#39;Nephrosearch Stichting tot steun van het wetenschappelijk onderzoek van de afdeling Nierziekten van het LUMC&#39;. This work was in part supported by the Leiden University Fund &#34;Prof. Jaap de Graeff-Lingling Wiyadhanrma Fund&#34; GWF2019-02. This work is supported by the partners of Regenerative Medicine Crossing Borders (RegMedXB) and Health Holland, Top Sector Life Sciences &#38; Health. C.W. van den Berg and T.J. Rabelink are supported by The Novo Nordisk Foundation Center for Stem Cell Medicine (reNEW), The Novo Nordisk Foundation Center for Stem Cell Medicine is supported by Novo Nordisk Foundation grants (NNF21CC0073729).

In accordance with Dutch law, approval by the animal welfare committee was not required for this research.
1. Preparing hiPSC-derived kidney organoids for transplantation
<ol>
	<li>Differentiate hiPSCs to kidney organoids using the protocol developed by Takasato et al.4,18,31. Culture the organoids following this protocol on polyester cell culture inserts with 0.4 &#181;m pores (cell culture inserts) until day 7 + 12 of differentiation. Each cell culture insert will contain three organoids.</li>
	<li>Remove the organoids from the cell culture insert.
	<ol>
		<li>Make a hole in the middle of the membrane of the cell culture insert with a pair of dissecting forceps. Using dissecting scissors, make three cuts in the membrane from the hole in the middle to the edge of the cell culture insert, cutting between the organoids. This will result in three pieces of membrane, each with one organoid attached, which are still connected to the cell culture insert at their outer edge.</li>
		<li>Take hold of one of the pieces of membrane close to its outer edge with a pair of forceps and tear it loose from the cell culture insert. Place the piece of membrane with the attached organoid in a Petri dish. Add a few drops of Dulbecco&#39;s phosphate-buffered saline (DPBS) without calcium and magnesium (DPBS-/-) to the organoid to avoid dehydration. Repeat for the other two organoids.</li>
	</ol>
	</li>
	<li>Bisect the organoids by holding their membrane in place with forceps and cutting them in half with a double-edge stainless steel razor blade (a whole organoid is too large for the celom to accommodate at this stage of development). Gently push the two organoid halves off the membrane with a dura dissector. Discard the membrane and leave the organoids in DPBS-/- until transplantation. 
	NOTE: Prepare a maximum of three organoids at a time for transplantation to avoid stress to the organoids prior to transplantation. Ideally, one person prepares the organoids while another performs the transplantation.</li>
</ol>
2. Preparing chicken embryos for transplantation
<ol>
	<li>Incubate fertilized white leghorn eggs. Start the incubation of the eggs on kidney organoid differentiation day 7 + 8 to ensure the correct timing of the transplantation.
	<ol>
		<li>Place the fertilized white leghorn eggs (Gallus domesticus) horizontally on a holder (Figure 1A; day 0), marking the middle of the upward-facing side with a pencil. 
		NOTE: Either custom-made plastic holders or egg cartons can be used as holders to incubate the eggs.</li>
		<li>Place the holder with the eggs in a humidified incubator at 38 &#177; 1 &#186;C (Figure 1A; day 0).</li>
		<li>Incubate the eggs for 3 days, keeping the water basin in the incubator filled.</li>
	</ol>
	</li>
	<li>Create a window in the eggshell on day 3 of incubation.
	<ol>
		<li>On day 3 of incubation, place a small piece (~1 cm x 0.5 cm) of transparent tape on the pointed tip of the egg (small end). Make a small hole in the eggshell in the middle of the transparent tape by tapping it with the sharp end of a pair of dissecting scissors.</li>
		<li>Insert a 19 G needle on a 5 mL syringe into the hole at a 45&#176; angle, avoiding damage to the yolk sac, and aspirate 2-3 mL of albumen from the egg to lower the embryo inside the egg. Seal the hole with a second piece (~1 cm x 0.5 cm) of transparent tape.</li>
		<li>Place a large piece (~5 cm x 5 cm) of transparent tape on the pencil-marked, upward-facing side of the egg. Make a hole in the eggshell in the middle of the transparent tape by tapping it with the sharp end of a pair of dissecting scissors (Figure 1A; day 3).</li>
		<li>Starting from this hole, cut a small circular window in the eggshell using curved dissecting scissors. Look through this window to locate the embryo, then enlarge the window to optimize access to the embryo (Figure 1A; day 3).</li>
		<li>Remove any large pieces of eggshell that may have fallen on top of the embryo using forceps. Remove smaller pieces by placing a few drops of DPBS with calcium and magnesium (DPBS+/+) on the embryo with a plastic transfer pipette, then aspirating the DPBS+/+ with the eggshell into the pipette.</li>
		<li>Add three drops of DPBS+/+ supplemented with 0.5% penicillin/streptomycin to the egg using a plastic transfer pipette.</li>
		<li>Carefully seal the window with a large piece (~5 cm x 5 cm) of transparent tape before placing the egg back in the incubator until day 4 of incubation, when the embryo is in Hamburger-Hamilton stage 23-24 (HH 23-24)32. 
		NOTE: Sealing the window is very important to avoid dehydration and death of the embryo.</li>
		<li>Check the embryos daily for viability by looking at them through the tape (do not remove the tape to avoid dehydration). During these first days of incubation, the egg yolk color changes from bright to matte yellow upon embryo death. Discard deceased embryos.</li>
		<li>Keep the water basin in the incubator filled.</li>
	</ol>
	</li>
</ol>
3. Intracelomic transplantation on day 4 of incubation
<ol>
	<li>Gaining access to the celomic cavity
	<ol>
		<li>Cut the tape from the window with curved dissecting scissors.</li>
		<li>Place the egg under a dissecting microscope on a rubber holder or egg carton.</li>
		<li>The chicken embryo is now in HH 23-24 and lying on its left side, with its right side facing the viewer (Figure 1A; day 4). Locate the right wing and leg bud of the embryo, as the celom will be accessed between these two limb buds. In the area between the right wing and leg bud, create an opening consecutively in the vitelline membrane, the chorion, and the amnion, by holding them with two pairs of dissecting forceps and gently pulling them in opposite directions. 
		NOTE: The vitelline membrane is the first membrane that is encountered after opening the egg, and in some cases, is already damaged after making the window on day 3. If the embryo is rotated, lying on its right side with its left side facing the viewer (Figure 1A; day 4), it is necessary to turn it around to enable transplantation. To do this, make a large opening in the vitelline and chorion membrane, then carefully turn the embryo around using forceps.</li>
		<li>Check whether there is unobstructed access to the celomic cavity: gently take hold of the edge of the body wall between the wing and limb bud with a pair of dissecting forceps and pull it slightly toward the viewer. The celomic cavity must be clearly visible. Carefully insert a blunt but slim instrument (e.g., a blunt tungsten wire in a microscalpel holder) into the celomic cavity. 
		NOTE: If insertion of the blunt instrument into the celomic cavity is not possible, it means one or more of the membranes have not been properly opened.</li>
	</ol>
	</li>
	<li>Transplantation
	<ol>
		<li>Place half an organoid inside the egg on top of the allantois using a dura dissector.</li>
		<li>Using dissecting forceps, carefully take hold of the edge of the body wall and pull it slightly toward the viewer to make the opening to the celom visible. 
		NOTE: Avoid damaging the blood vessels in the body wall.</li>
		<li>Gently move the organoid toward and through the opening in the body wall into the celom with a blunt Tungsten wire in a microscalpel holder. Push the organoid slightly cranially to lodge it inside the celom. It is now visible just behind the wing bud (Figure 1A; day 4).</li>
		<li>Add three drops of DPBS+/+ to the egg using a plastic transfer pipette.</li>
		<li>Carefully seal the window with a large piece (~5 cm x 5 cm) of transparent tape before placing the egg back in the incubator until day 12 of incubation (8 days after transplantation).</li>
		<li>Keep checking the embryos daily for viability by looking at them through the tape (do not remove the tape to avoid dehydration). Discard deceased embryos. 
		NOTE: As the embryos and their chorioallantoic membranes grow, they become more clearly visible through the tape. A lack of movement by the embryo and collapsed or sparse blood vessels are a sign of embryo death.</li>
		<li>Check the water basin in the incubator daily and keep it filled.</li>
	</ol>
	</li>
</ol>
4. Injection of fluorescently labeled lectin
<ol>
	<li>Preparing for injection
	<ol>
		<li>Make glass microinjection needles by pulling glass microcapillaries in a micropipette puller with the following settings: Heat 533, Pull 60, Velocity 150, Time 200. While looking through the dissecting microscope, carefully break the tips off the microinjection needles using dissecting forceps to create an opening. 
		NOTE: The required settings for the micropipette puller may differ depending on the machine.</li>
		<li>Assemble the injection system. Take two pieces of 38 cm silicone tubing and connect them to each other by placing a 0.2 &#181;m filter between them. Insert a mouthpiece into the end of the tube that is connected with the filter outlet (silicone tube 1), and a connector to the end of the tube that is connected with the filter inlet (silicone tube 2). Finally, insert the microinjection needle into the connector (Figure 1B).</li>
		<li>Dilute fluorescently labeled lens culinaris agglutinin (LCA) with DPBS-/- to a concentration of 2.5 mg mL-1 in a 0.5 mL tube and spin down for ~30 s in a microcentrifuge to move the aggregates to the bottom of the tube.</li>
		<li>Pipette 20 &#181;L of LCA onto a piece of parafilm.</li>
		<li>Aspirate the 20 &#181;L of LCA from the parafilm into the microcapillary needle of the assembled injection system.</li>
	</ol>
	</li>
	<li>Injection
	<ol>
		<li>Cut the tape from the window with curved dissecting scissors. Place the egg under a dissecting microscope in a rubber holder.</li>
		<li>Evaluate the vasculature and locate the veins, which can be distinguished from the arteries by their slightly brighter red color (Figure 1A; day 12). To improve access to the vasculature, carefully enlarge the window by cutting with curved dissecting scissors. Select a vein for injection based on accessibility and size.</li>
		<li>Insert the tip of the microcapillary needle into the selected vein at a 0&#176;-20&#186; angle. Ensure the needle is in the vein by gently moving the tip from side to side. Gently and steadily blow into the injection system to inject the LCA (Figure 1A; day 12). 
		NOTE: If the needle in the vein is in the correct position, it should stay within the boundaries of the vein.</li>
		<li>Place the egg back in the incubator for 10 min to let the LCA circulate. 
		&#8203;NOTE: It is not necessary to seal the egg with tape during this time.</li>
	</ol>
	</li>
</ol>
5. Collecting transplanted organoids on day 12 of incubation
<ol>
	<li>Sacrificing the chicken embryo
	<ol>
		<li>Place the egg in a rubber holder on the bench. Cut the tape from the window with curved dissecting scissors. Then, cut through the membranes surrounding the embryo with curved dissecting scissors. 
		NOTE: A dissecting microscope is not required for this step.</li>
		<li>Scoop the embryo up from the egg with a perforated spoon and immediately decapitate the embryo using scissors. Place the body of the embryo in a Petri dish under the dissection microscope.</li>
	</ol>
	</li>
	<li>Locating and collecting organoids
	<ol>
		<li>Place the embryo on its back in the Petri dish and spread its limbs.</li>
		<li>Carefully open the abdominal wall of the embryo along the longitudinal axis using forceps.</li>
		<li>Locate the organoid inside the embryo. The organoid most frequently becomes attached to the right liver lobe, either at the caudal tip or cranially just below the rib cage (Figure 1A; day 12). It is therefore recommended to start by looking in these locations.</li>
		<li>Once the organoid is located, remove it from the embryo by cutting around it with micro scissors. Place the organoid and the chicken tissue that is inevitably attached to it in a Petri dish under the dissecting microscope. Remove as much chicken tissue as possible with a double-edge stainless steel razor blade.</li>
		<li>Process the organoid depending on the desired analysis.</li>
	</ol>
	</li>
</ol>
6. Whole-mount immunofluorescence staining
<ol>
	<li>Place a transplanted organoid in a 24-well plate and fix in 500 &#181;L of 4% paraformaldehyde (PFA) at 4 &#176;C for 24 h. Wash 3x with DPBS-/-.</li>
	<li>Permeabilize and block the organoid in 300 &#181;L of blocking solution (0.3% Triton-X in DPBS-/- containing 10% donkey serum) for 2 h at room temperature.</li>
	<li>Prepare the primary antibody mix: for one organoid, dilute primary antibodies NPHS1 (sheep-&#945;-human, dilution of 1:100), CD31 (mouse-&#945;-human, dilution of 1:100), and LTL (biotin-conjugated, dilution of 1:300) in 300 &#181;L of blocking solution. Add the antibody mix to the organoid and incubate for 72 h at 4 &#176;C.</li>
	<li>Wash 3x with 0.3% TritonX in DPBS-/-.</li>
	<li>Prepare the secondary antibody mix: for one organoid, dilute secondary antibodies donkey-&#945;-sheep Alexa Fluor 647 (dilution of 1:500), donkey-&#945;-mouse Alexa Fluor 488 (dilution of 1:500), and streptavidin Alexa Fluor 405 (dilution of 1:200) in 300 &#181;L of blocking solution. Add the secondary antibody mix to the organoid and incubate for 2-4 h at room temperature. Cover with aluminum foil to avoid exposure of the secondary antibodies to light.</li>
	<li>Wash 3x with DPBS-/-.</li>
	<li>Embed the organoid in ~30 &#181;L of mounting medium in a 35 mm glass bottom dish and allow to dry overnight at room temperature, covered with aluminum foil. Store at 4 &#176;C.</li>
	<li>Image using a confocal microscope.</li>
</ol>

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E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+development+of+melanophores+from+embryonic+mouse+tissues+grown+in+the+coelom+of+chick+embryos.">The development of melanophores from embryonic mouse tissues grown in the coelom of chick embryos.</a> Proceedings of the National Academy of Sciences. 26 (12), 673-680 (1940).</li><li>Garreta, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fine+tuning+the+extracellular+environment+accelerates+the+derivation+of+kidney+organoids+from+human+pluripotent+stem+cells.">Fine tuning the extracellular environment accelerates the derivation of kidney organoids from human pluripotent stem cells.</a> Nature Materials. 18 (4), 397-405 (2019).</li><li>Koning, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Vasculogenesis+in+kidney+organoids+upon+transplantation.">Vasculogenesis in kidney organoids upon transplantation.</a> NPJ Regenerative Medicine. 7 (1), 40 (2022).</li><li>Hamburger, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Morphogenetic+and+axial+self-differentiation+of+transplanted+limb+primordia+of+2-day+chick+embryos.">Morphogenetic and axial self-differentiation of transplanted limb primordia of 2-day chick embryos.</a> Journal of Experimental Zoology. 77 (3), 379-399 (1938).</li><li>Dossel, W. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=New+method+of+intracelomic+grafting.">New method of intracelomic grafting.</a> Science. 120 (3111), 262-263 (1954).</li><li>Takasato, M., Er, P. X., Chiu, H. S., Little, M. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Generation+of+kidney+organoids+from+human+pluripotent+stem+cells.">Generation of kidney organoids from human pluripotent stem cells.</a> Nature Protocols. 11 (9), 1681-1692 (2016).</li><li>Hamburger, V., Hamilton, H. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+series+of+normal+stages+in+the+development+of+the+chick+embryo.">A series of normal stages in the development of the chick embryo.</a> Developmental Dynamics. 195 (4), 231-272 (1992).</li><li>Aleksandrowicz, E., Herr, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ethical+euthanasia+and+short-term+anesthesia+of+the+chick+embryo.">Ethical euthanasia and short-term anesthesia of the chick embryo.</a> ALTEX. 32 (2), 143-147 (2015).</li><li>. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ACUC.">ACUC.</a> Guideline: The Use and Euthanasia Procedures of Chicken/Avian Embryos. , (2012).</li></ol>

The authors have no conflicts of interest to disclose.

In this manuscript, a protocol for intracelomic transplantation of hiPSC-derived kidney organoids in chicken embryos is demonstrated. Upon transplantation, organoids are vascularized by perfused blood vessels that consist of a combination of human organoid-derived and chicken-derived ECs. These are spread throughout the organoid and invade the glomerular structures, enabling interaction between the ECs and podocytes. It was previously shown that this leads to enhanced maturation of the organoid glomerular and tubular structures28. The transplantation is very efficient, taking ~5 min per embryo, and the only maintenance that the embryos require is a regular refill of the water basin in the incubator. This method is therefore very suitable for the analysis of large numbers of organoids.
Vascularization and maturation of kidney organoids have been previously demonstrated through transplantation in mice18,20. In these labor-intensive mouse models, organoids were transplanted for 2 to up to 12 weeks. In the intracelomic transplantation experiments shown here, the duration of transplantation was limited to 8 days. This allows for the sacrifice of the embryos before day 13 of incubation, when they are thought to start experiencing pain33,34. Since chicken embryos hatch on day 21, the duration of transplantation could be extended to 15 days, sacrificing the embryos on day 19 to avoid hatching. This would, however, require the use of anesthetics. Despite the relatively short transplantation duration in this model, it induces extensive vascularization and significant maturation of organoid nephrons compared to in vitro organoids, including the formation of a GBM between organoid podocytes and the invading ECs28.
When performing intracelomic transplantation experiments, it is important to consider that not all chicken embryos develop normally and survive. Usually, ~65% of the embryos placed in the incubator at day 0 reach the end point of the experiment. This is due to a combination of acute bleeding caused by vessel damage during transplantation (5%-10% in our hands) and the stress that is induced by the windowing and transplantation procedures. When embryos are in an earlier or later stage than HH 23-24, transplantation becomes complicated due to limited space and more extensive vasculature, respectively. If embryos are not in the correct stage at the expected time, this could be due to the temperature of the incubation, as a higher temperature generally leads to faster development.
Moreover, keeping fertilized eggs at room temperature for a prolonged period before incubation induces more variability in development. To avoid this, the temperature of the incubator must be kept stable throughout and between experiments, and incubation should be started within 3 days after delivery of the fertilized eggs. Unexpectedly high percentages of embryo death between procedures can be caused by dehydration. To avoid this, it is essential to add two or three drops of DPBS+/+ to each egg after opening it and after transplantation and seal the egg very carefully with tape, smoothing out creases in the tape as much as possible. Combining the windowing and transplantation steps to reduce the number of times the eggs are opened is not recommended, as windowing on day 4 considerably increases embryo death. This is the result of damage to blood vessels that have frequently become attached to the eggshell at this stage.
In conclusion, this method provides an efficient and powerful tool to induce vascularization and enhance maturation in kidney organoids. It can be used to study these processes in large numbers of organoids and has potential for the modeling of renal diseases that require a higher level of maturation than can currently be acquired in vitro.

Human induced pluripotent stem cell (hiPSC)-derived kidney organoids have been shown to have potential for developmental studies1,2,3,4, toxicity screening5,6, and disease modeling5,7,8,9,10,11,12,13. However, their applicability for these and eventual clinical transplantation purposes is limited by the lack of a vascular network. During embryonic kidney development, podocytes, mesangial cells, and vascular endothelial cells (ECs) interact to form the intricate structure of the glomerulus. Without this interaction, the glomerular filtration barrier, consisting of podocytes, the glomerular basement membrane (GBM), and ECs, cannot develop properly14,15,16. Although kidney organoids in vitro do contain some ECs, these fail to form a proper vascular network and diminish over time17. It is therefore not surprising that the organoids remain immature. Transplantation in mice induces vascularization and maturation of the kidney organoids18,19,20,21. Unfortunately, this is a labor-intensive process that is unsuitable for the analysis of large numbers of organoids.
Chicken embryos have been used to study vascularization and development for over a century22. They are easily accessible, require low maintenance, lack a fully functional immune system, and can develop normally after opening the eggshell23,24,25,26. The transplantation of organoids on their chorioallantoic membrane (CAM) has been shown to lead to vascularization27. However, the duration of transplantation on the CAM, as well as the level of maturation of the graft, are limited by CAM formation, which takes until embryonic day 7 to complete. Therefore, a method was recently developed to efficiently vascularize and mature kidney organoids through intracelomic transplantation in chicken embryos28. The celomic cavity of chicken embryos has been known since the 1930s to be a favorable environment for the differentiation of embryonic tissues29,30. It can be accessed early in embryonic development and allows for relatively unlimited expansion of the graft in all directions.
This paper outlines a protocol for the transplantation of hiPSC-derived kidney organoids in the celomic cavity of day 4 chicken embryos. This method induces vascularization and enhanced maturation of the organoids within 8 days. Injection of fluorescently labeled lens culinaris agglutinin (LCA) prior to sacrificing the embryos enables visualization of perfused blood vessels within the organoids through confocal microscopy.

<table><tbody><tr><td>0.2 &amp;micro;m filter: Whatman Puradisc 30 syringe filter 0.2 &amp;micro;m</td><td>Whatman</td><td>10462200</td><td></td></tr><tr><td>35 mm glass bottom dishes&amp;nbsp;</td><td>MatTek Corporation</td><td>P35G-1.5-14-C</td><td></td></tr><tr><td>Aspirator tube assemblies for calibrated microcapillary pipettes</td><td>Sigma-Aldrich</td><td>A5177-5EA</td><td>Contains silicone tubes, mouth piece and connector</td></tr><tr><td>Confocal microscope: Leica White Light Laser Confocal Microscope&amp;nbsp;</td><td>Leica</td><td>TCS SP8</td><td></td></tr><tr><td>Dissecting forceps, simple type. Titanium, curved, with fine sharp tips</td><td>Hammacher Karl</td><td>HAMMHTC091-10</td><td></td></tr><tr><td>Dissecting forceps, simple type. Titanium, straight, with fine sharp tips</td><td>Hammacher Karl</td><td>HAMMHTC090-11</td><td></td></tr><tr><td>Dissecting microscope&amp;nbsp;</td><td>Wild Heerbrugg</td><td>355110</td><td></td></tr><tr><td>Dissecting scissors, curved, OP-special, extra sharp/sharp</td><td>Hammacher Karl</td><td>HAMMHSB391-10</td><td></td></tr><tr><td>Donkey serum</td><td>Sigma-Aldrich</td><td>D9663</td><td></td></tr><tr><td>Donkey-&amp;alpha;-mouse Alexa Fluor 488</td><td>ThermoFisher Scientific</td><td>A-212-02</td><td>dilution 1:500</td></tr><tr><td>Donkey-&amp;alpha;-sheep Alexa Fluor 647</td><td>ThermoFisher Scientific</td><td>A-21448</td><td>dilution 1:500</td></tr><tr><td>Double edged stainless steel razor blades</td><td>Electron Microsopy Sciences</td><td>72000</td><td></td></tr><tr><td>DPBS, calcium, magnesium (DPBS-/-)</td><td>ThermoFisher Scientific</td><td>14040133</td><td></td></tr><tr><td>DPBS, no calcium, no magnesium (DPBS+/+)</td><td>ThermoFisher Scientific</td><td>14190094</td><td></td></tr><tr><td>Egg cartons or custom made egg holders&amp;nbsp;</td><td>NA</td><td>NA</td><td></td></tr><tr><td>Fertilized white leghorn eggs (&lt;em&gt;Gallus Gallus Domesticus&lt;/em&gt;)&amp;nbsp;</td><td>Drost Loosdrecht B.V.</td><td>NA</td><td></td></tr><tr><td>Incubator</td><td>Elbanton BV</td><td>ET-3 combi</td><td></td></tr><tr><td>Lotus Tetragonolobus lectin (LTL) Biotinylated</td><td>Vector Laboratories</td><td>B-1325</td><td>dilution 1:300</td></tr><tr><td>Micro scissors, straight, sharp/sharp, cutting length 10 mm</td><td>Hammacher Karl</td><td>HAMMHSB500-09</td><td></td></tr><tr><td>Microcapillaries: Thin wall glass capillaries 1.5 mm, filament</td><td>World Precision Instruments</td><td>TW150F-3</td><td></td></tr><tr><td>Micropipette puller</td><td>Sutter Instrument Company</td><td>Model P-97</td><td>We use the following settings: Heat 533, Pull 60, Velocity 150, Time 200</td></tr><tr><td>Microscalpel holder: Castroviejo blade and pins holder, 12 cm, round handle, conical 10 mm jaws.</td><td>Euronexia</td><td>L-120</td><td></td></tr><tr><td>Mounting medium: Prolong Gold Antifade Mountant&amp;nbsp;</td><td>ThermoFisher Scientific</td><td>P36930</td><td></td></tr><tr><td>Olivecrona dura dissector 18 cm&amp;nbsp;</td><td>Reda</td><td>41146-18</td><td></td></tr><tr><td>Parafilm&amp;nbsp;</td><td>Heathrow Scientific</td><td>HS234526B</td><td></td></tr><tr><td>Penicillin-streptomycin 5,000 U/mL</td><td>ThermoFisher Scientific</td><td>15070063</td><td></td></tr><tr><td>Perforated spoon&amp;nbsp;</td><td>Euronexia</td><td>S-20-P</td><td></td></tr><tr><td>Petri dish 60 x 15 mm&amp;nbsp;</td><td>CELLSTAR</td><td>628160</td><td></td></tr><tr><td>Plastic transfer pipettes&amp;nbsp;</td><td>ThermoFisher Scientific</td><td>PP89SB</td><td></td></tr><tr><td>Purified mouse anti-human CD31 antibody</td><td>BD Biosciences</td><td>555444</td><td>dilution 1:100</td></tr><tr><td>Rhodamine labeled Lens Culinaris Agglutinin (LCA)</td><td>Vector Laboratories</td><td>RL-1042</td><td>This product has recently been discontinued. Vectorlabs does still produce Dylight 649 labeled LCA (DL-1048-1) and fluorescein labeled LCA (FL-1041-5)</td></tr><tr><td>Sheep anti-human NPHS1 antibody</td><td>R&amp;amp;D systems</td><td>AF4269</td><td>dilution 1:100</td></tr><tr><td>Sterile hypodermic needles, 19 G</td><td>BD microlance</td><td>301500</td><td></td></tr><tr><td>Streptavidin Alexa Fluor 405</td><td>ThermoFisher Scientific</td><td>S32351</td><td>dilution 1:200</td></tr><tr><td>Syringe 5 mL</td><td>BD Emerald</td><td>307731</td><td></td></tr><tr><td>Transparent tape&amp;nbsp;</td><td>Tesa</td><td>4124</td><td>Available at most hardware stores</td></tr><tr><td>Triton X</td><td>Sigma-Aldrich</td><td>T9284</td><td></td></tr><tr><td>Tungsten wire, 0.25 mm dia&amp;nbsp;</td><td>ThermoFisher Scientific</td><td>010404.H2</td><td></td></tr></tbody></table>

efficient vascularization of kidney organoids through intracelomic transplantation in chicken embryos

Kidney organoids derived from human induced pluripotent stem cells contain nephron-like structures that resemble those in the adult kidney to a certain degree. Unfortunately, their clinical applicability is hampered by the lack of a functional vasculature and consequently limited maturation in vitro. The transplantation of kidney organoids in the celomic cavity of chicken embryos induces vascularization by perfused blood vessels, including the formation of glomerular capillaries, and enhances their maturation. This technique is very efficient, allowing for the transplantation and analysis of large numbers of organoids. This paper describes a detailed protocol for the intracelomic transplantation of kidney organoids in chicken embryos, followed by the injection of fluorescently labeled lectin to stain the perfused vasculature, and the collection of transplanted organoids for imaging analysis. This method can be used to induce and study organoid vascularization and maturation to find clues for enhancing these processes in vitro and improve disease modeling.

The method and timeline for the differentiation of hiPSCs to kidney organoids, incubation of fertilized chicken eggs, transplantation of kidney organoids, injection of LCA, and collection of the organoids are summarized in Figure 1A. It is important to coordinate the timing of organoid differentiation and chicken egg incubation, starting differentiation 15 days before incubation.&#160;The actions on day 0, 3, 4, and 12 of incubation are illustrated by photographs below the timeline. Organoids are transplanted at day 7 + 12 of differentiation into day 4 (HH 23-24) chicken embryos. LCA is injected into the venous system of the embryo 8 days after transplantation to stain the perfused vasculature, before sacrificing the embryo and retrieving the organoid. The assembled injection system is shown in Figure 1B.
Chicken embryos are sacrificed on day 12 of incubation (Figure 1A). Upon careful dissection of the embryo, the transplanted organoid can usually be found attached to the liver. Looking through a dissection microscope, it appears vascularized (Figure 1A; day 12;&#160;step 5.2.3). Confocal imaging of transplanted organoids injected with LCA and stained for nephron structures and human ECs confirms vascularization by perfused blood vessels (Figure 2A), which also invade glomerular structures (Figure 2B). The vasculature is chimeric: perfused human ECs (CD31+, LCA+), unperfused human ECs (CD31+, LCA-), and perfused chicken-derived ECs (CD31-, LCA+) can be distinguished (Figure 2A, panels III, IV, V).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65090/65090fig01.jpg" /> 
Figure 1: Intracelomic transplantation method. (A)&#160;Timeline of the differentiation of hiPSCs to kidney organoids, incubation of fertilized chicken eggs, and intracelomic transplantation of kidney organoids. Differentiation of hiPSCs to kidney organoids is initiated 15 days before incubation of the chicken eggs is started (differentiation day 0 = incubation day -15) to enable transplantation of the kidney organoids on day 7 + 12 of differentiation in chicken embryos on day 4 of incubation. Days of incubation: Day 0: Fertilized chicken eggs are positioned horizontally on holders (step 2.1.1), which are placed in an incubator at 38 &#186;C &#177; 1 &#176;C (protocol step 2.1.2). Day 3: A window is created in each egg by making a small hole in the upward-facing side of the egg with the sharp end of a pair of curved dissecting scissors (step 2.2.3) and cutting a circular window starting from this hole (step 2.2.4). The window is sealed with transparent tape before placing the egg back in the incubator. Day 4: Intracelomic transplantation of kidney organoids on day 7 + 12 of differentiation is performed. The embryo is in HH 23-24 and lying on its left side, with its right side facing the viewer (step 3.1.3). Between the wing and leg bud, an opening is made in the vitelline membrane, chorion, and amnion to obtain access to the celomic cavity, and the organoid is inserted through these openings into the celom. After transplantation, the organoid is visible as a white structure located just behind the wing bud. The edges of the opened vitelline and amnion membranes are visible (steps 3.2.3&#160;and 3.2.3-Zoom).&#160;In some cases, the embryos are rotated, lying on their right instead of left side (3.1.3 NOTE), and must be turned around prior to transplantation. Day 12: Fluorescently labeled lectin is injected intravenously. Veins are distinguished from arteries by their color; the blood in the veins is oxygen-rich, coming from the CAM, so they are a slightly brighter red than the arteries that are coming from the embryo (steps 4.2.2., 4.2.2-Zoom, and 4.2.3.). The embryo is sacrificed and the organoid retrieved. The organoid (circled) has become attached to the chicken liver and appears to be vascularized (step 5.2.3). Scale bar = 1 mm. The schematic image depicting the intracelomic transplantation image in the top panel was reprinted with permission from Koning et al.28. (B) Image of the assembled injection system, consisting of a mouthpiece, two pieces of 38 cm silicone tubing, a 0.2 &#181;m filter, a connector, and a glass microinjection needle that was generated by pulling glass microcapillaries in a micropipette puller. Abbreviations: A = allantois;&#160;Am = amnion; C = celom; CC = cut chorion membrane; CHIR = CHIR99021; FGF9 = fibroblast growth factor 9; hiPSCs = human induced pluripotent stem cells; IM = intermediate mesoderm; Lb = leg bud; O = organoid; PS = primitive streak; U = umbilical ring; V = vitelline membrane; Wb = wing bud; Y = yolk stalk. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65090/65090fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65090/65090fig02.jpg" /> 
Figure 2: Vascularized transplanted kidney organoids. (A) Immunofluorescent images of (I) an untransplanted kidney organoid and (II) a transplanted kidney organoid. In both conditions, glomerular (NPHS1+, cyan) and tubular (LTL+, yellow) structures are visible. In the untransplanted organoids, some human ECs (CD31+, green) are present. In the transplanted organoid, a perfused vascular network (CD31+, green; injected rhodamine-labeled LCA+, white) is visible throughout the organoid. In panels III, IV, and V, magnifications of the boxed areas in panel II are shown, to demonstrate the three types of ECs that can be distinguished in transplanted organoids. Panel III contains perfused human ECs (CD31+, LCA+), marked with arrowheads. Panel IV contains unperfused human ECs (CD31+, LCA-), marked with arrowheads. Panel V contains perfused chicken-derived ECs (CD31-, LCA+), marked with arrowheads. Scale bar = 200 &#181;m. (B) In untransplanted organoids (I), ECs (CD31+, green) surround glomerular structures (NPHS1+, cyan) but do not invade them. In transplanted organoids (II), glomerular structures (NPHS1+, blue) are vascularized by perfused capillaries (LCA+, white; CD31+, green). Scale bar = 50 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65090/65090fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Efficient Vascularization of Kidney Organoids through Intracelomic Transplantation in Chicken Embryos at JoVE.com

Efficient Vascularization of Kidney Organoids through Intracelomic Transplantation in Chicken Embryos

Here, we present a detailed protocol for the transplantation of kidney organoids in the celomic cavity of chicken embryos. This method induces vascularization and enhanced maturation of the organoids within 8 days and can be used to study these processes in an efficient manner.

Kidney organoids derived from human induced pluripotent stem cells contain nephron-like structures that resemble those in the adult kidney to a certain ...

efficient-vascularization-kidney-organoids-through-intracelomic

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

1.7K Views.  Leiden University Medical Center. In vitro gekweekte nierorganoïden blijven onvolgroeid, waardoor hun toepasbaarheid wordt beperkt. En deze methode van intracoelomale transplantatie induceert vascularisatie en verbetert de rijping van de organoïden. De procedure is zeer efficiënt, waardoor de transplantatie en analyse van een groot aantal organoïden in één experiment mogelijk is.Na het bereiden van HIPSC-afgeleide nierorganoïden en het incuberen van de bevruchte leghorn-eieren creëer je een venster in de eiersc...

Video: Efficient Vascularization of Kidney Organoids through Intracelomic Transplantation in Chicken Embryos

In Utero Intra-cardiac Tomato-lectin Injections on Mouse Embryos to Gauge Renal Blood Flow

The formation and perfusion of developing renal blood vessels (apart from glomeruli) are greatly understudied. As vasculature develops via angiogenesis (which is the branching off of major vessels) and vasculogenesis (de novo vessel formation), perfusion mapping techniques such as resin casts, in vivo ultrasound imaging, and micro-dissection have been limited in demonstrating the intimate relationships between these two processes and developing renal structures within the embryo. Here, we describe the procedure of in utero intra-cardiac ultrasound-guided FITC-labeled tomato lectin microinjections on mouse embryos to gauge the ontogeny of renal perfusion. Tomato lectin (TL) was perfused throughout the embryo and kidneys harvested. Tissues were co-stained for various kidney structures including: nephron progenitors, nephron structures, ureteric epithelium, and vasculature. Starting at E13.5 large caliber vessels were perfused, however peripheral vessels remained unperfused. By E15.5 and E17.5, small peripheral vessels as well as glomeruli started to become perfused. This experimental technique is critical for studying the role of vasculature and blood flow during embryonic development.

In Utero Intra-cardiac Tomato-lectin Injections on Mouse Embryos to Gauge Renal Blood Flow

This manuscript describes a technique for visualization of the developing vasculature. Here we utilized in utero intra-cardiac FITC-labeled tomato lectin microinjections on mouse embryos. Using this technique, we delineate the perfused and unperfused vessels throughout the embryonic kidney.

The formation and perfusion of developing renal blood vessels (apart from glomeruli) are greatly understudied. As vasculature develops via angiogenesis ...

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.

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.

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, ...

Comparing Metastatic Clear Cell Renal Cell Carcinoma Model Established in Mouse Kidney and on Chicken Chorioallantoic Membrane

Metastatic clear cell renal cell carcinoma (ccRCC) is the most common subtype of kidney cancer. Localized ccRCC has a favorable surgical outcome. However, one third of ccRCC patients will develop metastases to the lung, which is related to a very poor outcome for patients. Unfortunately, no therapy is available for this deadly stage, because the molecular mechanism of metastasis remains unknown. It has been known for 25 years that the loss of function of the von Hippel-Lindau (VHL) tumor suppressor gene is pathognomonic of ccRCC. However, no clinically relevant transgenic mouse model of ccRCC has been generated. The purpose of this protocol is to introduce and compare two newly established animal models for metastatic ccRCC. The first is renal implantation in the mouse model. In our laboratory, the CRISPR gene editing system was utilized to knock out the VHL gene in several RCC cell lines. Orthotopic implantation of heterogeneous ccRCC populations to the renal capsule created novel ccRCC models that develop robust lung metastases in immunocompetent mice. The second model is the chicken chorioallantoic membrane (CAM) system. In comparison to the mouse model, this model is more time, labor, and cost-efficient. This model also supported robust tumor formation and intravasation. Due to the short 10 day period of tumor growth in CAM, no overt metastasis was observed by immunohistochemistry (IHC) in the collected embryo tissues. However, when tumor growth was extended by two weeks in the hatched chicken, micrometastatic ccRCC lesions were observed by IHC in the lungs. These two novel preclinical models will be useful to further study the molecular mechanism behind metastasis, as well as to establish new, patient-derived xenografts (PDXs) toward the development of novel treatments for metastatic ccRCC.

Metastatic clear cell renal cell carcinoma is a disease without a comprehensive animal model for thorough preclinical investigation. This protocol illustrates two novel animal models for the disease: the orthotopically implanted mouse model and the chicken chorioallantoic membrane model, both of which demonstrate lung metastasis resembling clinical cases.

Metastatic clear cell renal cell carcinoma (ccRCC) is the most common subtype of kidney cancer. Localized ccRCC has a favorable surgical outcome. However, ...

Cancer Research

Dissection and Culture of Mouse Embryonic Kidney

The goal of this protocol is to describe a method for the dissection, isolation, and culture of mouse metanephric rudiments. 
During mammalian kidney development, the two progenitor tissues, the ureteric bud and the metanephric mesenchyme, communicate and reciprocally induce cellular mechanisms to eventually form the collecting system and the nephrons of the kidney. As mammalian embryos grow intrauterine and therefore are inaccessible to the observer, an organ culture has been developed. With this method, it is possible to study epithelial-mesenchymal interactions and cellular behavior during kidney organogenesis. Furthermore, the origin of congenital kidney and urogenital tract malformations can be investigated. After careful dissection, the metanephric rudiments are transferred onto a filter that floats on culture medium and can be kept in a cell culture incubator for several days. However, one must be aware that the conditions are artificial and could influence the metabolism in the tissue. Also, the penetration of test substances could be limited due to the extracellular matrix and basal membrane present in the explant.
One main advantage of organ culture is that the experimenter can gain direct access to the organ. This technology is cheap, simple, and allows a large number of modifications, such as the addition of biologically active substances, the study of genetic variants, and the application of advanced imaging techniques.

This protocol describes a method for isolating and culturing metanephric rudiments from mouse embryos.

The goal of this protocol is to describe a method for the dissection, isolation, and culture of mouse metanephric rudiments. 
During mammalian kidney ...

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 ...

Optimization of Renal Organoid and Organotypic Culture for Vascularization, Extended Development, and Improved Microscopy Imaging

Embryonic kidney organotypic cultures, and especially pluripotent stem cell-derived kidney organoids, are excellent tools for following developmental processes and modelling kidney disease. However, the models are limited by a lack of vascularization and functionality. To address this, an improved protocol for the method of xenografting cells and tissues to the chorioallantoic membrane (CAM) of an avian embryo to gain vascularization and restoration of blood flow was developed. The grafts are overlaid with custom-made minireservoirs that fix the samples to the CAM and supply them with culture medium that protects the grafts from drying. The improved culture method allows xenografts to grow for up to 9 days. The manuscript also describes how to provide optimal conditions for long-term confocal imaging of renal organoids and organotypic cultures using the previously published Fixed Z-Direction (FiZD) method. This method gently compresses an embryonic organ or organoid between a glass coverslip and membrane in a large amount of medium and provides excellent conditions for imaging for up to 12 days. Together, these methods allow vascularization and blood flow to renal organoids and organotypic kidney cultures with improved confocal imaging. The methods described here are highly beneficial for studying fundamental and applied functions of kidneys ex vivo. Both methods are applicable to various types of tissues and organoids.

This work describes two methods for studying organ development, an improved xenotransplantation setup on chorioallantoic membrane (CAM) from avian embryos that allows for vascularization of cultured embryonic organs and organoids and a novel fixed z-direction organ culture method with modified experimental conditions that allows for high-resolution time-lapse confocal imaging.

Embryonic kidney organotypic cultures, and especially pluripotent stem cell-derived kidney organoids, are excellent tools for following developmental ...

Xenotransplantation of Human Stem Cells into the Chicken Embryo

The chicken embryo is a classical animal model for studying normal embryonic and fetal development and for xenotransplantation experiments to study the behavior of cells in a standardized in vivo environment. The main advantages of the chicken embryo include low cost, high accessibility, ease of surgical manipulation and lack of a fully developed immune system. Xenotransplantation into chicken embryos can provide valuable information about cell proliferation, differentiation and behavior, the responses of cells to signals in defined embryonic tissue niches, and tumorigenic potential. Transplanting cells into chicken embryos can also be a step towards transplantation experiments in other animal models. Recently the chicken embryo has been used to evaluate the neurogenic potential of human stem and progenitor cells following implantation into neural anlage1-6. In this video we document the entire procedure for transplanting human stem cells into the developing central nervous system of the chicken embryo. The procedure starts with incubation of fertilized eggs until embryos of the desired age have developed. The eggshell is then opened, and the embryo contrasted by injecting dye between the embryo and the yolk. Small lesions are made in the neural tube using microsurgery, creating a regenerative site for cell deposition that promotes subsequent integration into the host tissue. We demonstrate injections of human stem cells into such lesions made in the part of the neural tube that forms the hindbrain and the spinal cord, and into the lumen of the part of the neural tube that forms the brain. Systemic injections into extraembryonic veins and arteries are also demonstrated as an alternative way to deliver cells to vascularized tissues including the central nervous system. Finally we show how to remove the embryo from the egg after several days of further development and how to dissect the spinal cord free for subsequent physiological, histological or biochemical analyses.

In this paper we present a method for transplanting human stem cells into various regions of the central nervous system of the chicken embryo. This provides an in vivo model for assessing the proliferation and differentiation of various types of human stem cells in embryonic tissue environments.

The chicken embryo is a classical animal model for studying normal embryonic and fetal development and for xenotransplantation experiments to study the ...

Biology

An ex-ovo Chicken Embryo Culture System Suitable for Imaging and Microsurgery Applications

Understanding the relationships between genetic and microenvironmental factors that drive normal and malformed embryonic development is fundamental for discovering new therapeutic strategies. Advancements in imaging technology have enabled quantitative investigation of the organization and maturing of the body plan, but later stage embryonic morphogenesis is less clear. Chicken embryos are an attractive vertebrate animal model system for this application because of its ease of culture and surgical manipulation. Early embryos can be cultured for a short time on filter paper rings, which enables complete optical access for cell patterning and fate studies1,2. Studying advanced developmental processes such as cardiac morphogenesis are traditionally performed through a window of the eggshell3-5, but this technique limits optical access due to window size. We previously developed a simple method to culture whole embryos ex-ovo on hexagonal weigh boats for up to 10 days, which enabled high resolution imaging via ultrasonography6,7. These cultures were difficult to transport, limiting the types of imaging tools available for live experiments. We here present an improved shell-less culture system with a cost-effective, portable environmental chamber. Eggs were cracked onto a hammock created by a polyurethane membrane (cling wrap) affixed circumferentially to a plastic cup partially filled with sterile water. The dimensions of the circumference and depth of the hammock were both critical to maintain surface tension, while the mechanics of the hammock and water beneath helped dampen vibrations induced by transportation. A small footprint circulating water bath was also developed to enable continuous temperature control during experimentation. We demonstrate the ability to culture embryos in this way for at least 14 days without morphogenic defect or delay and employ this system in several microsurgical and imaging applications.

An ex-ovo Chicken Embryo Culture System Suitable for Imaging and Microsurgery Applications

In this article, we present a simple methodology to enable long-term ex-ovo avian embryo culture. This technique is ideal for longitudinal experimentation requiring complete optical accessibility and/or sterile transportation in avian embryos.

Understanding the relationships between genetic and microenvironmental factors that drive normal and malformed embryonic development is fundamental for ...

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 ...

Grafting Human Brain Organoids into the Chorioallantoic Membrane of the Chick Embryo

Early Unguided Human Brain Organoid Neurovascular Niche Modeling into the Permissive Chick Embryo Chorioallantoic Membrane

Engrafting organoids into vascularized tissues in model animals, such as the immunodeficient mouse or chick embryo chorioallantoic membrane (CAM), has proven efficient for neovascularization modeling. The CAM is a richly vascularized extraembryonic membrane, which shows limited immunoreactivity, thus becoming an excellent hosting model for human origin cell transplants.
This paper describes the strategy to engraft human brain organoids differentiated at multiple maturation stages into the CAM. The cellular composition of brain organoids changes with time, reflecting the milestones of human brain development. We grafted brain organoids at relevant maturation stages: neuroepithelial expansion (18 DIV), early neurogenesis (60 DIV), and early gliogenesis (180 DIV) into the CAM of embryonic day (E)7 chicken embryos. Engrafted brain organoids were harvested 5 days later and their histological features were analyzed.
No histological signs of neovascularization in the grafted organoids or abnormal blood vessels adjacent to the graftings were detected. Moreover, remarkable changes were observed in the cellular composition of the grafted organoids, namely, an increase in the number of glial fibrillary acidic protein-positive-reactive astrocytes. However, the cytoarchitectural changes were dependent on the organoid maturation stage. Altogether, these results suggest that brain organoids can grow in the CAM, and they show differences in the cytoarchitecture depending on their maturation stage at grafting.

Author Spotlight: Optimizing the Neurovascular Development of Human Brain Organoid in Chick Embryo

Here, we present a protocol to engraft human brain organoids at multiple maturation stages into the chick chorioallantoic membrane (CAM). Brain organoids were grown following unguided standardized protocols.

Engrafting organoids into vascularized tissues in model animals, such as the immunodeficient mouse or chick embryo chorioallantoic membrane (CAM), has ...

Efficient Vascularization of Kidney Organoids through Intracelomic Transplantation in Chicken Embryos

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