Efficient Vascularization of Kidney Organoids through Intracelomic Transplantation in Chicken Embryos

Summary

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.

Abstract

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.

Introduction

Human induced pluripotent stem cell (hiPSC)-derived kidney organoids have been shown to have potential for developmental studies^1,2,3,4, toxicity screening^5,6, and disease modeling^{5,7,8,9,10,11,12,13}. 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 properly^14,15,16. 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 organoids^18,19,20,21. 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 eggshell^23,24,25,26. 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 tissues^29,30. 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.

Protocol

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

1. 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 µm pores (cell culture inserts) until day 7 + 12 of differentiation. Each cell culture insert will contain three organoids.
2. Remove the organoids from the cell culture insert.
   1. 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.
   2. 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's phosphate-buffered saline (DPBS) without calcium and magnesium (DPBS^-/-) to the organoid to avoid dehydration. Repeat for the other two organoids.
3. 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.

2. Preparing chicken embryos for transplantation

1. 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.
   1. 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.
   2. Place the holder with the eggs in a humidified incubator at 38 ± 1 ºC (Figure 1A; day 0).
   3. Incubate the eggs for 3 days, keeping the water basin in the incubator filled.
2. Create a window in the eggshell on day 3 of incubation.
   1. 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.
   2. Insert a 19 G needle on a 5 mL syringe into the hole at a 45° 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.
   3. 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).
   4. 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).
   5. 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.
   6. Add three drops of DPBS^+/+ supplemented with 0.5% penicillin/streptomycin to the egg using a plastic transfer pipette.
   7. 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.
   8. 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.
   9. Keep the water basin in the incubator filled.

3. Intracelomic transplantation on day 4 of incubation

1. Gaining access to the celomic cavity
   1. Cut the tape from the window with curved dissecting scissors.
   2. Place the egg under a dissecting microscope on a rubber holder or egg carton.
   3. 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.
   4. 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.
2. Transplantation
   1. Place half an organoid inside the egg on top of the allantois using a dura dissector.
   2. 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.
   3. 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).
   4. Add three drops of DPBS^+/+ to the egg using a plastic transfer pipette.
   5. 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).
   6. 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.
   7. Check the water basin in the incubator daily and keep it filled.

4. Injection of fluorescently labeled lectin

1. Preparing for injection
   1. 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.
   2. Assemble the injection system. Take two pieces of 38 cm silicone tubing and connect them to each other by placing a 0.2 µm 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).
   3. 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.
   4. Pipette 20 µL of LCA onto a piece of parafilm.
   5. Aspirate the 20 µL of LCA from the parafilm into the microcapillary needle of the assembled injection system.
2. Injection
   1. Cut the tape from the window with curved dissecting scissors. Place the egg under a dissecting microscope in a rubber holder.
   2. 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.
   3. Insert the tip of the microcapillary needle into the selected vein at a 0°-20º 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.
   4. Place the egg back in the incubator for 10 min to let the LCA circulate.
      ​NOTE: It is not necessary to seal the egg with tape during this time.

5. Collecting transplanted organoids on day 12 of incubation

1. Sacrificing the chicken embryo
   1. 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.
   2. 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.
2. Locating and collecting organoids
   1. Place the embryo on its back in the Petri dish and spread its limbs.
   2. Carefully open the abdominal wall of the embryo along the longitudinal axis using forceps.
   3. 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.
   4. 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.
   5. Process the organoid depending on the desired analysis.

6. Whole-mount immunofluorescence staining

1. Place a transplanted organoid in a 24-well plate and fix in 500 µL of 4% paraformaldehyde (PFA) at 4 °C for 24 h. Wash 3x with DPBS^-/-.
2. Permeabilize and block the organoid in 300 µL of blocking solution (0.3% Triton-X in DPBS^-/- containing 10% donkey serum) for 2 h at room temperature.
3. Prepare the primary antibody mix: for one organoid, dilute primary antibodies NPHS1 (sheep-α-human, dilution of 1:100), CD31 (mouse-α-human, dilution of 1:100), and LTL (biotin-conjugated, dilution of 1:300) in 300 µL of blocking solution. Add the antibody mix to the organoid and incubate for 72 h at 4 °C.
4. Wash 3x with 0.3% TritonX in DPBS^-/-.
5. Prepare the secondary antibody mix: for one organoid, dilute secondary antibodies donkey-α-sheep Alexa Fluor 647 (dilution of 1:500), donkey-α-mouse Alexa Fluor 488 (dilution of 1:500), and streptavidin Alexa Fluor 405 (dilution of 1:200) in 300 µL 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.
6. Wash 3x with DPBS^-/-.
7. Embed the organoid in ~30 µL 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 °C.
8. Image using a confocal microscope.

Results

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. The actions on day 0, 3, 4, and 12 of incubation are illustrated by photographs below the timeline. Organoid...

Discussion

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 str...

Materials

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