Dissection and Culture of Mouse Embryonic Kidney

Mice were maintained according to Swedish regulations and European Union legislation (2010/63/EU). All procedures were performed following the guidelines of the Swedish Ethics Committee (permits C79/9, C248/11, and C135/14). Procedures at Heidelberg University involving animal subjects have been approved by the Regierungspräsidium Karlsruhe and the Animal Welfare Officers at the University of Heidelberg.

1. Preparation of Reagents and Materials for Culture

NOTE : Use a laminar flow hood to minimize contamination.

1. On the day of dissection, pre-cluster human Fc alone or recombinant chimeric ephrin protein fused to human Fc by mixing it with anti-human Fc goat antibodies at a molar ratio of 1:5 in sterile phosphate-buffered saline (PBS). Incubate for 1 h at 37 °C⁵.
2. Prepare the culture medium by supplementing Dulbecco's Modified Eagle Medium: Nutrient Mixture F-12 (DMEM/F12) with 1% penicillin/streptomycin (v/v), 1% glutamine (v/v), 1% insulin-transferrin-selenium (v/v), 50 µg/mL human holo-transferrin, and 1.5 µg/mL amphotericin.
   NOTE: 10 mL of medium is sufficient for 16 embryos.
3. Add clustered recombinant ephrin solution at a final concentration of 20 µg/mL to the culture medium. Use clustered human Fc as control.
4. Prepare culture plates by adding 500 µL of culture medium to each well of a sterile, uncoated, flat-bottom polystyrene 4-well plate.
5. Using sterile forceps, carefully place one polycarbonate membrane filter with a pore size of 5 µm per well on top of the medium. Transfer the prepared plate to the incubator (37 °C/5% CO₂). Ensure that the filter stays dry on the upper surface and floats.
   NOTE: One filter is sufficient for two kidney anlagen.
6. Using a razor blade, cut approximately 30 pipette tips (volume: 100 µL) approximately 1 mm from the tip to result in a wider opening. Place the tips back into a pipette-tip rack.

2. Dissection of Metanephric Rudiments at E11.5

1. Sacrifice a timed-pregnant mouse at E11.5 by cervical dislocation (or using a method approved by the local ethics committee).
2. Remove the uterus and the embryos, as described by Brown et al ⁶. From this point forward, work under a dissection microscope.
3. Using No. 5 watchmaker forceps, cut the embryo directly under the forelegs. Use a fresh petri dishfor each embryo. Remove the legs and the tail of the embryo. To do this, grab the tissue to be removed with one pair of No. 5 watchmaker forceps and use the tips of a second pair to cut.
4. To remove the ventral organ mass, use the tips of a pair of forceps to laterally open the body wall of the embryo in a rostral-to-caudal direction.
   1. Cut superficially, parallel to the dorsal aorta. Use the blood-filled and therefore visible dorsal aorta for orientation. Carefully separate the ventral part of the body wall from the dorsal part using the tips of closed forceps and flip the embryo over. Cut the body wall laterally in a rostral-to-caudal direction and using the dorsal aorta for orientation, as for the previous side.
5. Once the ventral body wall has been separated from the dorsal body, grab it with one pair of forceps and carefully pull to remove it, together with the organ mass.
   NOTE: The metanephric anlagen usually stay attached to the dorsal body wall. They are oval, ~ 200 µm long structures located at the level of the hind limbs.
6. Hold the dorsal body wall with one pair of forceps, with the ventral side facing up. Using the other pair of forceps, grab the dorsal aorta at its rostral end and carefully peel the dorsal aorta from the dorsal body wall.
   NOTE: Both metanephric kidneys usually stay attached to the dorsal aorta.
7. Using one pair of forceps, cut rostrally to the kidney anlagen to remove the aorta. Then, carefully separate the two kidney anlagen using the tips of the forceps.
8. Carefully peel off unwanted tissue from the kidney anlagen using the tips of the forceps. Take care not to damage the mesenchyme, as damage can result in poor growth and exclusion of the explant from further analysis.
9. Transfer the kidney anlagen separately using a microliter pipette and a wide-open pipette tip in 10 µL of PBS to the polycarbonate membrane filter in a 4-well-plate.
   NOTE: Due to surface tension, the explant will be covered with a thin layer of medium. One filter is sufficient for two kidney anlagen.
   1. Place each of the two kidney anlagen at the center of each respective half of the filter. Transfer the plate back to the incubator at 37 °C/5% CO₂. Leave the plate in the incubator until the metanephric rudiments from the next embryo are ready to be placed on the filter.
10. Repeat steps 2.3-2.9.1 for each embryo.
    NOTE: With some practice, the dissection procedure will take about 5 min per embryo.
11. Incubate the kidney anlagen for 3 days at 37 °C/5% CO₂.

3. Preparation of Reagents for Fixation and Staining

1. To prepare 4% paraformaldehyde (PFA) in PBS without calcium and magnesium (w/v), dissolve the PFA at 60 °C in PBS, continuously stirring. Cool to room temperature and use a paper filter to remove the remaining particles. Aliquot and keep at 4 °C for immediate use or freeze for long-term storage.
   CAUTION: PFA is toxic. Wear the personal protective equipment specified in the local safety standards and work in a fume hood. Discard waste following institutional guidelines and using designated waste containers.
2. To prepare permeabilization solution, dilute Triton X-100 in PBS without calcium and magnesium to 0.3% (v/v).
3. To prepare blocking solution, add 5% (v/v) goat serum and 0.1% Triton X-100 to PBS without calcium and magnesium. Add sodium azide to a final concentration of 0.02% (w/v). Store at 4 °C.
   CAUTION: Sodium azide is toxic. Handle sodium azide in accordance with the local safety and environmental protection regulations.
   NOTE: When using antibodies for staining, use 5% serum from the species the secondary antibody was derived from to make blocking solution.
4. Dilute biotinylated Dolichorus biflorus agglutinin 1:200 in blocking solution. Dilute Alexa488-conjugated streptavidin 1:200 in blocking solution.
5. To make ready-to-use embedding medium (see the table of materials), add 0.1 g of water-soluble polyvinyl alcohol-based mucoadhesive/mL to 25% (w/v) glycerol in 0.1 M Tris-HCl (pH 8.5). Heat to 50 °C under continuous agitation for 1 h, cool down, and adjust the pH to 8.0-8.5 using 1 M NaOH. Avoid higher NaOH concentrations. Add 100 µg/mL N-propylgallate and thimerosal to a final concentration of 0.02% (w/v). Stir for 30 min at room temperature.
   CAUTION: Thimerosal is toxic. Handle it in accordance with the local safety and environmental protection regulations.
   1. Fill the embedding medium in 50 mL tubes, balance the rotor, and centrifuge at 3,200 x g and room temperature for 10 min. Decant the clear solution into 15 mL tubes and discard the pellet. Aliquot and store at -20 °C for several weeks. Once thawed, keep the solution at 4 °C. Warm to ~30 °C immediately before use.
6. Prepare as many glass slides as filters with explants are to be mounted. For this, glue two small coverslips, 18 x 18 mm, on either side of the glass slide using instant adhesive, Leave a ~15 mm space for the filter in between.

4. Fixation and staining

1. After a culturing period of 3 days in vitro (div), remove the plates from the incubator and carefully aspirate the culture medium from each well using a micropipette. Make sure not to touch the filter and keep the tip opening towards the wall of the well.
2. Add 500 µL of 4% PFA/PBS to each well. The filters will float on the fixative. Using a micropipette, carefully add the PFA fixative dropwise to submerge the filters. Incubate at room temperature for 30 min.
   NOTE: For all following steps, care must be taken not to wash off the explants from the filter. Keep the opening of the pipette tip towards the wall of the well.
3. Remove the PFA and rinse twice with 500 µL of PBS, submerging the filter. Add 500 µL of permeabilization solution to each well and incubate at room temperature for 1 h.
4. Rinse twice with 500 µL of PBS and add 500 µL of blocking solution to each well. Incubate at room temperature for 1 h.
5. Replace the blocking solution with 250 µL of biotinylated Dolichorus biflorus agglutinin diluted 1:200 in blocking solution and incubate at 4 °C for 24 h.
6. Remove the Dolichorus biflorus agglutinin solution and rinse twice with 500 µL of PBS per well.
7. Add 250 µL of Alexa488-conjugated streptavidin diluted 1:200 in blocking solution and incubate at 4 °C for 24 h. Alternatively, incubate at room temperature for 2 h.
   NOTE: A better signal-to-noise ratio is achieved when incubated at 4 °C.
8. Rinse twice with 500 µL of PBS per well. Using forceps, transfer the filters to the prepared glass slides, with the explants facing up. Mount with ~50-100 µL of embedding medium. Cover with 60 mm-long No. 1.5 coverslips. Allow the embedding medium solution to solidify for 2 h at room temperature in the dark. Proceed to imaging or store the slides, wrapped in foil, at 4 °C until ready to image.
9. Image with a widefield epifluorescence microscope⁷ coupled to an Hg-lamp and use a mirror unit for blue excitation (excitation bandpass, 460 – 495 nm; dichromatic mirror, 505 nm; and emission bandpass, 510 – 550 nm) and 20X Plan-Apochromat lenses, 0.75 numerical aperture.