Using Ex Vivo Upright Droplet Cultures of Whole Fetal Organs to Study Developmental Processes during Mouse Organogenesis

All mice used in these studies were CD-1 mice obtained from Charles River Laboratories. Previous culture experiments have also been performed on other strains, such as C57BL/6J (data not shown), but any strain can be used. Pregnant adult females were approximately 2-3 months old and were euthanized via CO₂ inhalation followed by cervical dislocation and bilateral thoracotomy prior to embryo removal. Mice were housed in accordance with NIH guidelines, and experimental protocols were approved by the Institutional Animal Care and Use Committee of Cincinnati Children’s Hospital Medical Center.

1. Preparation of Instruments, Culture Media, and Dishes

1. Prepare a 70% ethanol solution. Autoclave deionized water (for making humidified chambers and for making/diluting solutions). Sterilize dissection tools (forceps, scissors, and 27 G needle syringes) by spraying down with 70% ethanol.
2. Prepare 10X phosphate-buffered saline (PBS) containing calcium and magnesium (80 g NaCl, 2 g KCl, 14.4 g Na₂HPO₄, 2.4 g KH₂PO₄, 6.75 ml of 1 M CaCl₂, 3.75 ml of 1 M MgCl₂). Stir to dissolve in 1 L sterile autoclaved water and then filter with filter paper. Dilute 10-fold with water to make 1X PBS.
3. Heat inactivate fetal bovine serum (FBS) at 55 °C for 30 min and filter sterilize with a 0.22 µm syringe filter.
4. Prepare 38 ml 1X complete culture medium (called complete DMEM, cDMEM), consisting of Dulbecco’s Modified Eagle Medium (DMEM), 10% filtered heat-inactivated FBS and 1/100 dilution of penicillin-streptomycin stock. This usually should be used fresh, but can be stored at 4 °C for 1 month.
5. Reconstitute the VEGFR Tyrosine Kinase Inhibitor II (TKI II) in DMSO for a stock concentration of 5 mg/ml, and dilute stock with cDMEM to make a working solution. The final concentration for this study is 1.875 µg/ml in cDMEM. According to the company’s recommendations, stock solutions are stable for up to 6 months at -20 °C. As for working solutions, make fresh and use on the same day.
   Note: The concentration of this drug in the working solution should be two-fold higher than the desired final concentration (since it will be diluted two-fold in the droplet). At the same time, prepare the same volume of cDMEM containing the same volume of DMSO for the “control” treatment.
6. Prepare the tail digestion reagents (50 mM NaOH and 1 M Tris-HCl [pH 8.0]) separately in distilled water.
7. Make a 25 μM stock of the XY PCR primers (XY Forward 5'-TGA AGC TTT TGG CTT TGA G-3' and XY Reverse 5'-CCG CTG CCA AAT TCT TTG G-3') together in nuclease-free water.
8. Prepare 50X TAE buffer (242 g Trizma Base, 57.1 ml Glacial Acetic Acid, 100 ml 0.5 M EDTA, pH 8.5, and add water to 1 L final volume). Prepare 1X TAE Running buffer (20 ml 50X TAE diluted with water to 1 L total volume).
9. To create a 2.5% agarose solution (0.625 g agarose, 0.5 ml 50X TAE Buffer, 24.5 ml water), heat agarose solution in microwave until boiling, then let cool until about 55-65 °C (able to touch). Once cool, add 2 µl of 1% ethidium bromide solution, and pour gel.
   CAUTION! Ethidium bromide is a mutagen and teratogen; please use nitrile gloves and proper safety protocol when handling. Alternatively, other potentially less toxic gel resolving agents or dyes can be used.
10. Prepare Blocking Solution (PBS with 0.1% Triton X-100, 10% fetal bovine serum [FBS], and 3% bovine serum albumin [BSA]) for immunofluorescence.
11. Prepare Washing Solution (PBS with 0.1% Triton X-100, 1% FBS, and 3% BSA) for immunofluorescence.
12. Prepare PBTx (PBS with 0.1% Triton X-100) for immunofluorescence.
13. Prepare the incubation chambers for culture. Fill large (100 mm diameter) petri dishes with 35 ml autoclaved water. Place near where dissection and cultures will be performed.

2. Isolation of Fetal Testes from Mus musculus

1. Check the female mouse for vaginal plugs each morning. Noon on the day at which the vaginal plug is detected is considered E0.5. Euthanize the pregnant female mouse at E11.5, in a manner consistent with approved animal protocols.
2. Spray down mouse abdomen with 70% ethanol.
3. Open the belly skin and peritoneum with a V-shaped incision using fine scissors and pull back flap of tissue to expose interior organs.
4. Locate the ovaries at both ends of the uterine horn. Using scissors, cut at the ovary and the connective tissue to separate the uterus containing the embryos from the mother’s body.
5. Open the uterine wall by gently cutting the side opposite the placentae, exposing the yolk sacs. Be careful not to puncture yolk sac to avoid damaging or losing embryos.
6. Cut near the placentae to remove the embryo-containing yolk sacs and place them into a dish containing PBS with calcium and magnesium. The presence of calcium and magnesium ions will help support cell adhesion molecules to maintain tissue architecture and prevent the tissue from sticking to the dissection tools.
7. With forceps, remove the yolk sac and amnion from the embryo by carefully puncturing the yolk sac to create a hole in which the embryo can slide out. Once the embryo is outside of the yolk sac, cut the umbilical cord.
8. From this point forward, use a microscope located in a sterile tissue culture hood. Remove the head of embryo and discard by pinching with forceps on either side of neck.
9. Remove the tail and place into new microcentrifuge tube for XY PCR analysis to determine sex of embryo.
   Note: While it is optimal to culture gonads as soon as possible after harvest, it is also possible to remove tail DNA and perform XX/XY genotyping prior to culture, so that the sex of each embryo is known and only the desired sex needs to be cultured. While the genotyping is being done, embryos can be kept separated (so that they can be tracked) at 4 °C or on ice until ready for culture. One caveat is that this period outside in utero or culture conditions may potentially affect viability for culture or subsequent development during culture.
10. Secure the embryo in a supine position against the bottom of the culture dish by pinning the armpits of embryo with one pair of forceps (usually the forceps held in the weak hand). Maintain these forceps in place to hold the embryo still during the entire dissection process.
11. With other pair of forceps, remove skin covering abdomen and gently remove liver, intestines, and other organs to expose back body wall.
12. As the gonads will be located on the back body wall on either side of the dorsal aorta, a large blood vessel running along the midline of the body, scoop underneath the urogenital ridge with closed forceps and remove the urogenital ridge by opening the forceps and lifting up. As the gonads will be loosely attached to the wall, be careful not to pull up too fast without removing these connections or the gonads may stretch, causing damage to the organs.
13. Move the urogenital ridge into cDMEM in a dish to acclimate tissue to media.
14. Separate the gonad-mesonephros complex from the rest of the urogenital ridge using 27 G needles. Use one needle to cut by pressing down, and use the other to guide the tissue correctly to allow optimal separation. Avoid using a sawing motion against the dish bottom, as sharp needles will create shards of plastic that can stick to the tissue. Make sure to keep the mesonephros completely attached to the gonad.

3. Culturing of Gonad with a Small-molecule Inhibitor

1. Set two 20 µl pipettes to 15 µl each. Cut about 1-2 mm off one of the barrier pipette tips using a clean, sterilized razor blade for transfer of the gonads and addition of control cDMEM, while the other pipette will be used solely for drug-treated cDMEM.
2. Line up gonads with their long axis parallel to the pipette tip so they can easily be pipetted using the cut-off tip. Be careful of gonads sticking to the inside of the pipette tip.
3. Label one side as the control droplet and the other as the drug-containing droplet. Only two droplets can fit on a 35 mm dish lid. Make sure that labels on lids match the labels on tail-tissue-containing microcentrifuge tubes.
4. Pipette 15 µl of cDMEM containing a single gonad into a droplet in the lid of a small (35 mm) culture dish (use only the lids, as the bottoms are too tall to fit within a humidified petri dish chamber). Place two separate gonad-containing droplets on either side of the dish, well-separated from one another. Check under the microscope to ensure that the gonads have been transferred to the droplets.
5. To the droplet designated as the control, add an additional 15 µl of cDMEM containing DMSO (made in step 1.5) to make a droplet with a 30 µl total volume. Use only the “control” pipette that will not contact the drug.
6. To the other droplet (drug-treated sample), add 15 µl of TKI-II-containing cDMEM to make a droplet with a 30 µl total volume. Use only the pipette designated for drug-treated samples.
7. Using the pipette, spread out the droplets in an expanding circular pattern until they are about 15-18 mm in diameter and the gonad is located roughly in the middle. Orient the gonad so that it lies on its side and the gonad and mesonephros are easily distinguishable. Orient the lung so that the two lobes lay flat.
   1. Although droplet diameter may vary slightly, ensure that gonads are not floating and are held in place by surface tension. Make sure that droplets do not touch each other or the side of the lid. Without the surface tension, the organs will grow onto the lid during the culture and flatten, thus distorting organ morphology.
8. Carefully place the culture dish lid containing the droplets upright onto the surface of the water in the large (100 mm) humidifier dish. Ensure there are no bubbles trapped underneath the bottom of the lid and that it is lying flat on the surface of the water. Do not invert the lid and be careful not to let any water touch the top of the lid where the droplets are located.
9. To create a small, humidified chamber, immediately cover with the lid of the larger humidifier dish to enclose the gonad cultures. Act as quickly as possible to minimize any evaporation of media. Ensure that the smaller dish has room between it and the lid of the larger dish to move around freely; otherwise, condensation may make a seal and block air exchange to the tissue.
10. Once two small lids are placed into the chamber, immediately place the chamber into the incubator. Incubate the droplet cultures for 48 hr in a humidified CO₂ incubator at 37 °C.

4. Polymerase Chain Reaction for Determining the Sex of Embryos

1. Add the embryonic tails to bottom of a 1.5 ml microcentrifuge tube.
2. Add to each tube 200 µl of 50 mM NaOH.
3. Place at 95 °C for 15 min or until tissue is completely dissolved.
4. Add 50 µl of 1M Tris-HCl and vortex lightly and briefly.
5. In a 1.5 ml microcentrifuge tube, make a fresh PCR master mix by multiplying each volume by the total number of samples: 0.5 µl 25 µM Primer solution, 2.5 µl 10X Taq Buffer, 0.5 µl 10 mM dNTPs (nucleotides), 19.3 µl nuclease-free water, 0.2 µl DNA Taq polymerase. Mix well.
6. Add 23 µl of PCR master mix to PCR tubes containing 3 μl lysate of digested tails.
7. Flick tubes to mix them, then perform a short spin to bring the samples to the bottom of the tube.
8. Run XY (Short) PCR program (Table 1).
9. Load the samples (with 5x dye) and ladder in a 2.5% agarose gel in 1X TAE buffer.
10. Run the agarose gel electrophoresis until XX and XY bands can be resolved.
11. Image the gel with UV light and capture image.
12. Analyze the X-chromosome-specific vs Y chromosome-specific bands (X chromosome=331 base pairs [bp] and XY=302 bp)¹³; XY (male) samples will have two bands of 331 and 302 bp, whereas XX (female) samples will appear as a single 331 bp band.

5. Whole Mount Organ Immunofluorescence

Day 1:

1. Remove the gonads from the culture using a 1,000 µl pipette with a cut-off tip. If desired, they may be placed in a dish of PBS to wash off media and reagents. Place into PBS in a 0.5 ml microcentrifuge tube. The smaller tube size will conserve reagents and make it easier to keep track of samples in the tube.
   1. Be sure to keep control and treated samples, as well as XX and XY samples, in separate tubes and remove them from culture with separate sets of pipettes to avoid potential drug contamination.
2. Wash the gonads twice with PBS. From this point on, this protocol can use 1X PBS lacking calcium and magnesium. To wash them, allow the gonads to sink to the bottom of the tube (by gravity) and then remove the liquid above. Be careful not lose gonads by pipetting too close to them.
3. Remove as much PBS as possible from the tube. Add 250 µl of 4% paraformaldehyde in PBS containing 0.1% Triton X-100, and let incubate O/N at 4 °C on a rocker. Alternatively, this fixation can be done for 2 hr at 4 °C on a rocker. CAUTION! Paraformaldehyde is a toxic substance, so follow safety protocol when handling.

Day 2:

4. Rinse the gonads twice in 250 µl PBTx at RT.
5. Wash the gonads 3 times with 250 µl PBTx for 10 min each at RT on a rocker.
6. Block the gonads at least 1 hr in 250 µl Blocking solution at RT on a rocker.
7. Stain the gonads O/N at 4 °C on a rocker in 250 µl primary antibody diluted in Blocking Solution.

Day 3:

8. Rinse the gonads twice in 250 µl Washing Solution.
9. Wash the gonads 3 times with 250 µl Washing Solution for 10 min each at RT on a rocker.
10. Block the gonads 1 hr in 250 µl Blocking Solution at RT on a rocker.
11. Cover the microcentrifuge tube with aluminum foil to protect fluorescent secondary antibodies from light.
12. Incubate the gonads 2-4 hr at RT on a rocker in 250 µl secondary antibody diluted in Blocking Solution containing Hoechst 33342. Alternatively, perform secondary antibody and Hoechst 33342 incubation O/N at 4 °C on rocker.
13. Rinse the gonads twice in 250 µl PBTx.
14. Wash the gonads 3 times in 250 µl PBTx for 10 min each at RT on a rocker.
15. Using a cut-off pipette tip, transfer gonads onto a slide with minimal liquid. Remove excess liquid with pipette, but make sure that tissue does not dry out.
16. Orient the gonads in desired fashion with forceps, and quickly add a drop of mounting media to mount the gonads on the slide. Make sure the mounting media coats all samples completely; it is critical to avoid letting samples dry out.
17. Use coverslips (number 1.5 style) of appropriate size to gently cover samples. Let mounting media spread to contact the entire surface of coverslip. If necessary, very lightly press on the corners of the coverslip so that there are no large air bubbles.
18. Seal the slides with clear nail polish around the edges to reduce evaporation of the mounting media. Store mounted slides in the dark at 4 °C.