Kidney Capsule Grafting

The kidney capsule is well established as an ectopic site able to provide a permissive environment for thymus development (see Note 4). Described below is one application of this technique to study early thymus organogenesis. Enzyme digestion and manual dissection are used to isolate the pharyngeal endoderm from the overlying ectoderm and mesoderm of E9.0 embryos. The resulting tissue explants containing the third pharyngeal pouches are grafted under the kidney capsule of an adult mouse and the ability to form a functional organ is assessed. All steps of this procedure should be carried out under sterile conditions as far as possible.

3.2.1. Isolation of the Pharyngeal Endoderm

1. Remove E9.0 embryos (10-15 somites) from the uterus into serum-free culture medium and remove all extra-embryonic tissues. M2 medium works well as the pH does not alter when exposed to air for long periods of time.

2. Remove all structures caudal to the heart. Digest the remaining tissue in a trypsin/pancre-atin enzyme mixture for 15 to 20 min on ice, then transfer to serum-containing medium and wash thoroughly (see Note 5).

3. Using an eyelash tool and a pair of fine forceps, remove the head and first pharyngeal arches. Carefully peel away the surface ectoderm, and then remove the heart and neural tube (see Note 6).

4. Carefully strip away the remaining mesenchyme using the eyelash tool, to leave a clean endodermal gut tube (see Note 6). Trim this down to include only the second and third pharyngeal pouches. Explants should be stored in M2 medium on ice until ready for grafting (see Note 7).

3.2.2. Anesthesia

1. Anesthetize mice with an intraperitoneal injection of Avertin (Tribromoethanol). For an average 40-g adult mouse, 5 to 10 mg of Avertin is required to allow approx 30 min to perform the procedure (see Note 8).

2. Administer an intraperitoneal injection of Torbugesic for pain relief. Use 0.2 mL of a 1% solution for an average 40-g adult mouse.

3.2.3. Preparation of the Tissue for Transplantation

1. Prepare a mouth-controlled micropipet: heat the shaft of a Pasteur pipet over a flame, and when molten pull into a capillary. Break the capillary about 2 cm from the shoulder of the pipet and connect the micropipet to a length of flexible rubber tubing and a mouthpiece.

2. Fill the mouth-controlled micropipet by dipping the tip into medium and allowing it to fill up by capillary action. Then draw a small column of air into the pipet to create an air bubble.

3. Pick up the tissue fragments in a small volume of fluid and position them about 5 mm from the tip of micropipet (see Note 9). Multiple tissue explants can be transplanted simultaneously (see Note 10).

4. Place the loaded micropipet on the microscope stage until required.

3.2.4. Exposure of the Kidney

1. Lay the anesthetized mouse on its side, with the head pointing either to the left or right, and sterilize the flank skin with 70% ethanol.

2. Make a 1-cm incision through the skin about 1 cm from the spine and immediately caudal to and parallel with the last rib.

3. Make a second incision of about 0.7 mm through the muscle layers to expose the kidney. Avoid blood vessels and nerves as much as possible.

4. Hold the incision open with an open pair of blunt, angled forceps and grasp the fat pad at the cranial pole of the kidney with a second pair. Pull the kidney out of the peritoneal cavity. Applying a slight pressure to the flank of body may also help ease the kidney through the incision. Use of a small incision will help prevent the kidney from sliding back into abdominal cavity.

5. Let the exposed kidney stand for about 30 s to dry the capsule slightly. This makes it easier to grasp with forceps.

3.2.5. Transplantation of the Tissue (Fig. 1A-C)

1. Pick up the kidney capsule (the thin, transparent covering of the kidney) with a pair of fine forceps and use a second pair to make a small hole in the capsule about 2 to 3 mm away from the holding forceps. Lift the holding forceps slightly to create a space between the capsule and the kidney surface.

2. Insert the micropipet containing the explants into the opening in the capsule until it is about 5 mm into the space created by the raised holding forceps (see Note 9). Blow gently into the mouthpiece to slowly expel the tissue. Use the air bubble as a guide and stop blowing just before it reaches the end of the micropipet. Carefully withdraw the pipet, maintaining a positive pressure until it is completely out of the capsule, to prevent the tissue being drawn back into the pipet. (see Notes 9-11 for alternative methods.)

3. Release the holding forceps to secure the grafts between the capsule and the kidney surface.

4. Check the micropipet to ensure that the tissue has been transplanted, and examine the kidney to ascertain the position of the grafts. Depending on the size of the grafts, a microscope may be required.

3.2.6. Suture and Recovery

1. Allow the kidney to slide back into the peritoneal cavity by manipulating the wound opening.

2. Stitch the muscle layers with suture thread to seal the peritoneal cavity and close the skin with a wound clip.

3. Warm the mice slightly on a standard small animal heat pad to aide recovery from anesthesia. A slide warmer (Fisher, cat. no. 12-594) set to 37°C is also effective.

3.2.7. Graft Recovery and Analysis

After the desired period of time (see Note 12), sacrifice the mouse and remove the kidney by cutting through the skin and muscle layers at the original wound site. The grafted tissue should be evident under the kidney capsule (Fig. 1D). The graft can then either be freed completely, or removed together with a small piece of the underlying kidney. The grafted tissue can now be analyzed as appropriate, e.g., using standard histological or immunohistological techniques. (also see Note 13).

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    How to implant thymus tissue under the kidney capsule of a mousde?
    10 days ago

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