Analysis of Transforming Growth Factor ß Family Cleavage Products Secreted Into the Blastocoele of Xenopus laevis Embryos

All procedures described are approved by the Institutional Animal Care and Use Committee (IACUC) of the University of Utah. Frogs are housed in an IACUC approved facility. Male frogs are euthanized by immersion in tricaine following by clipping the ventricle of the heart. Female frogs are housed in the laboratory for a maximum of 24 h following hormonal induction of spawning to allow for egg collection and then returned to the care facility.

1. Collection of X. laevis Testes

1. Anesthetize a sexually mature male in 0.2% Tricaine (Table 1) for 30-40 min until the animal is unresponsive to claw pinch.
2. Use blunt forceps to pull the skin away from the body wall. Make a horizontal incision in the skin across the lower abdomen using surgical scissors. Make a parallel incision in the underlying body wall, and then a third, perpendicular incision through the body wall up through the rib cage.
   1. Fold back the resulting flaps, locate the ventricle of the heart and clip the base to exsanguinate and ensure death.
3. Pull the yellow fat bodies out of the abdomen with blunt forceps and locate the testes on each side, underlying the base of the fat bodies. Remove each testis by clipping it away from the fat bodies and any attached viscera using surgical scissors.
4. Transfer testes to a Petri dish containing 1x Modified Barth's Solution (MBS) (Table 1). Rinse and remove any adherent blood vessels or attached viscera.
   1. Transfer one testis to a 35 mm Petri dish containing testis buffer (Table 1) for immediate use and store the other in a 10 mL conical tube filled with testis buffer for later use. Testis should be stored at 4 °C when not used and will remain viable for at least two weeks.
      ​NOTE: Photographs of testes isolation can be found in Reference 15.

2. Collection and Fertilization of X. laevis Eggs

NOTE: Frogs should be handled with vinyl gloves or hands washed before and after handling the frogs. Latex gloves, lotions and detergent residues on human hands will damage the frogs' fragile skin.

1. The evening before spawning, inject 0.3 mL (1200 IU) of human chorionic gonadotropin (Table 1) into the dorsal lymph sac of two or three mature X. laevis females using a 1 mL syringe fitted with a 26 G needle. Use a new needle for each injection.
2. After injection, place the females in a sealed container (not air-tight) in an 18 °C biological incubator. Spawning generally begins 14-16 h later.
   NOTE: Plastic drawers or plastic containers with air holes cut into the lid work well for holding females overnight. Be sure the lid is on firmly to prevent frogs from escaping and desiccating in the incubator.
3. Hold the female over a 100 mm Petri dish containing 1x MBS and apply gentle pressure to the frog's abdomen to expel eggs into the dish.
4. Cut a small piece of the testis (~1/5) using a razor blade and transfer it into a 1.5 mL tube containing ~500 µL of 1x MBS. Crush with a pellet pestle or tissue homogenizer. Store the resulting sperm suspension on ice or at 4 °C for subsequent fertilization on the same day.
5. Tilt the 100 mm Petri dish to pour off as much of the 1x MBS as possible, and remove the remainder using a plastic transfer pipette. Apply approximately 100 µL of sperm suspension, add about 1 mL of 0.1x MBS and swirl to mix.
   1. After 5 min, flood the dish with dechlorinated tap water or 0.1x MBS and let stand for 30 min.
      NOTE: Water can be dechlorinated using a filter or municipal tap water can be left standing uncovered for at least 24 h to allow the chlorine to dissipate in regions where conventional chlorine is added to the water. In areas where chloramine is added to disinfect drinking water, the water must instead be treated with a commercially available conditioner that can break down chloramine.
      NOTE: Eggs are collected in a high salt solution (1x MBS) to prevent the formation of a jelly coat, which would provide a barrier to fertilization. After fertilization, the low salt solution (dechlorinated water or 0.1x MBS) allows for jelly coat formation and the eggs will adhere to each other and the dish. Eggs that have been fertilized will rotate such that the pigmented animal pole faces upward within 30 min. If this does not happen, and the eggs look healthy under the microscope, sperm viability is suspect and it may be necessary to harvest new testes.
6. Pour off the solution covering the eggs and fill the Petri dish with freshly prepared 2% cysteine solution (Table 1). Swirl the solution in the dish for 3-5 min until the jelly coat is dissolved and the eggs are no longer sticking to each other or the dish.
   1. Tilt the dish and carefully pour off the cysteine solution, or remove with a plastic pipette. Replace with 1x DeBoer's pond water (Table 1). Swirl to rinse and decant.
   2. Repeat one wash with DeBoer's solution, one with 0.1x MBS, and refill the dish with 0.1x MBS. After removing the jelly coat, the eggs become more fragile and must not be exposed to the air-liquid interface.
7. Move the fertilized eggs to a 16 °C biological incubator or leave them on the bench at room temperature. If left at room temperature, the first cleavage will occur 1-1.5 h after fertilization, and subsequent cleavages will occur approximately every 30 min. By contrast, if embryos are incubated at 15-16 °C, cleavages will occur roughly every hour and it will be possible to inject a larger number of embryos from a given spawn.

3. Microinjection of X. laevis Embryos

1. Heat and pull glass capillaries to a fine point using a micropipette puller with the following settings: Two-step pull; Heat 1: 67.4 °C; Heat 2: 62 °C.
   NOTE: These settings are specific for the micropipette puller and glass capillaries used here (Table of Materials).
2. Under a dissecting microscope, clip off the tip of a pulled needle close to the end using Dumont #5 forceps. The needle should be sharp but not so thin that the tip bends when it contacts the embryo. Figure 1 shows an example of an appropriately pulled needle that has not been clipped for microinjection (A) and one that has been clipped (B).
3. Insert the needle into a needle holder connected to a microinjector. Attach the needle holder to a micromanipulator. Place 2-4 µL of capped synthetic RNA encoding the Tgfß precursor protein to be analyzed on a small piece of parafilm under a dissecting microscope.
   1. Using the micromanipulator, lower the needle into the drop and use the fill function on the microinjector to aspirate RNA into the needle. Be sure to keep the needle below the surface of the drop, and watch progress under the microscope to avoid aspirating air into the needle.
      NOTE: If antibodies that recognize the prodomain and/or ligand domain of the Tgfß precursor to be analyzed are not available, sequence encoding an epitope tag (e.g., V5, myc, Flag) can be inserted in-frame into the cDNA that will be used as a template for RNA transcription. The in vivo bioactivity of the tagged protein should be compared with that of untagged protein to ensure that the epitope does not interfere with proper folding, cleavage or activity.
4. Eject a drop of the RNA solution into the air and measure the drop size using a micrometer mounted on the injection stage or in the eyepiece of the microscope. Adjust the drop size to approximately 10 nL utilizing the time and pressure controls on the microinjector.
   NOTE: RNA concentrations in the range of 5-20 ng/µL are appropriate to deliver 50-200 pg of RNA into each embryo. This amount of RNA is generally sufficient to detect cleavage products in aspirates from 10-20 embryos. It is best to aim for the lowest dose of RNA that gives a detectable signal on immunoblots. Higher amounts of RNA can cause gastrulation defects, which interfere with the expansion of the blastocoele.
5. Move embryos to an injection dish or tray containing 5% Ficoll in 0.1x MBS (Table 1) when they reach the 2-4 cell stage. A 35 mm Petri dish fitted with a piece of nylon mesh in the bottom can serve as an inexpensive injection dish that will keep the embryos immobilized. Insert the needle near the animal pole and inject 10 nL of RNA into one cell of each embryo.
   NOTE: Embryos can easily be injected at any time between the one and 16 cell stages. Injecting the embryo after it has cleaved ensures that the egg was fertilized and that the embryo is healthy. The exact location of the injection is not essential. A more detailed description of spawning, culture and microinjection of RNAs into X. laevis embryos can be found in Reference 15.
6. Transfer the injected embryos into a 35 mm Petri dish filled with 5% Ficoll in 0.1x MBS (Table 1). Place the small dishes inside a 150 mm Petri dish and cover loosely with a lid to prevent the culture media from evaporating. Culture the embryos overnight at 15-16 °C.
   ​NOTE: Culturing embryos in Ficoll helps to heal the injection site. If fine needles are used, culturing in 2-3% Ficoll is sufficient, whereas 5% Ficoll is preferred if apparent damage is observed. Keeping the embryos in Ficoll solution for 1-2 h after injection is enough to aid healing, but overnight incubation does not damage the embryo as long as the solution is changed before the onset of gastrulation.

4. Blastocoele Extraction and Analysis of Tgfß Cleavage Products

1. The following morning after injection, remove the Ficoll solution and remove any dead or dying embryos. Rinse the embryos once or twice with 0.1x MBS and culture the embryos in 0.1x MBS on the bench at room temperature.
   NOTE: Embryos can also be returned to the 16 °C biological incubator to slow down development.
2. Heat and pull glass capillaries to a fine point using a micropipette puller with the following settings: Two step pull; Heat 1: 67.4 °C; Heat 2: 62 °C.
   NOTE: These settings are specific for the puller and glass capillary used here (Table of Materials).
3. Under a dissecting microscope, clip off the tip of a pulled needle using forceps. To prevent clogging, the opening of a needle used for blastocoele aspiration should be larger than that in a needle used for microinjection. An example of a good pulled and clipped needle to be used for blastocoele aspiration is shown in Figure 1C. Insert the aspiration needle into needle holder connected to a microinjector and attach the needle holder to a micromanipulator.
   NOTE: The optimal needle diameter is determined empirically by trial and error. Needles with a large diameter fill too quickly and make it more likely that cellular debris will enter the needle. By contrast, very small diameter needles are likely to become clogged. Once an appropriate needle is generated, it's useful to generate multiple needles clipped at the same position.
4. Place embryos in an injection tray or dish filled with 0.1x MBS when they reach the early to the mid-gastrula stage (stage 10-11).
5. Insert the needle below the surface of the embryo near the animal pole. Press the fill button on the microinjector while looking through the dissecting microscope to keep an eye on the needle and the embryo. Over a few seconds, the level of clear fluid will rise in the needle and the embryo will collapse and become concave.
   1. If cloudy white matter enters the needle, let go of the fill button and pulse the inject button one or more times to eject any cloudy matter, which consists of cellular debris that is likely to contain proteases.
      NOTE: If the blastocoele collapses and cell debris enters the needle as soon as the fill button is pressed, this indicates that the opening in the needle is too large. If the blastocoele fails to collapse but white matter enters the needle immediately, this shows that the needle was inserted too deeply and is touching the blastocoele floor. Eject the white matter, and move to the next embryo. The aspiration rate can be more carefully controlled by pulsing the fill button rather than pressing it continuously. This is particularly important if the opening in the needle is larger than optimal.
6. Repeat step 4.5 until the fluid has been aspirated from the blastocoele of 10-20 embryos.
   NOTE: Generally, blastocoele fluid aspirated from 10-20 embryos is sufficient to detect cleavage products on immunoblots. However, this can vary depending on antibody quality and must be determined empirically.
7. Place a piece of paraffin on the injection tray and pipette 1 µL of nuclease-free water onto it. Submerge the needle under the drop of water and press the inject button to dispel the blastocoele fluid into the water.
   1. Alternatively, eject the fluid directly onto the parafilm, but in this case, pulse the inject button rather than holding it down. This expels the fluid under lower pressure, preventing it from flattening out on the parafilm, making it difficult to draw up in a pipette.
8. Transfer the blastocoele fluid into a sterile microcentrifuge tube on ice. Expect to harvest approximately 0.3-0.5 µL of blastocoele fluid per embryo. Add nuclease-free water to adjust the final volume to 30 µL.
9. Transfer the blastocoele fluid-depleted embryos to a separate tube on ice. Remove excess 0.1x MBS and add 200 µL of chilled (4 °C) embryo lysate buffer (10 µL per embryo) (Table 1). Pipette the embryos up and down 10-20 times using a micropipette until they are fully homogenized and no clumps remain.
10. Centrifuge the homogenized embryos in a refrigerated microcentrifuge at 10,000 x g for 10 min. Remove 160 µL of the supernatant using a P200 pipette and transfer to a new tube on ice, being careful to avoid the white yolk proteins and other cellular debris in the bottom half of the tube.
    1. Repeat the microcentrifugation once and transfer 128 µL of the clear supernatant to a new tube on ice. At this point, cleared embryo lysates and blastocoele fluid can be stored at -80 °C for as long as desired.
       NOTE: With a few exceptions, Tgfß precursor proteins are not secreted into the blastocoele and will only be detected in the depleted embryo lysates. By contrast, Tgfß family cleavage products are primarily seen in the blastocoele fluid. It is helpful to harvest and analyze both the embryo lysates and the blastocoele fluid for most purposes. At a minimum, this provides a positive control to ensure that the precursor was robustly translated and stable. In addition, it allows one to compare relative ratios of cleavage products to precursors among different wild type or mutant proteins, to identify mutations that enable intact precursors to be secreted into blastocoele fluid (e.g., mutations within the PC consensus motif that allow for proper folding and secretion without cleavage, in which case the intact precursor is detected in both the embryo and the blastocoele fluid) or mutations that generate misfolded, non-cleavable proteins (in which case the precursor will be detected in the embryo lysate, but cleavage products will be absent in the blastocoele fluid).
11. Deglycosylate proteins present in blastocoele fluid with PNGase F at this point by following the manufacturer's instructions.
    NOTE There is no need to deglycosylate precursor proteins present in the embryo lysate since these represent endoplasmic reticulum (ER)–resident forms of the precursor that lack the complex N-linked oligosaccharides removed by PNGase F. By contrast, cleavage products present in blastocoele fluid have moved through the trans-Golgi network (TGN) where carbohydrates are further modified and can now be removed by PNGase F. Following deglycosylation, cleavage products migrate as a more tightly condensed band on SDS gels, which can aid in visualizing, quantitating and accurately identifying cleavage products. This is not strictly required, but it can be helpful, for example, if you are trying to distinguish between homodimers and heterodimers based on migration patterns or if a cleavage product contains heterogeneous carbohydrates that cause it to migrate as a broad fuzzy band.
12. Add 5 µL of reducing 4x sample buffer (Table 1) to 15 µL of blastocoele fluid and 15 µL of clarified embryo lysate. Add 5 µL of non-reducing 4x sample buffer (Table 1) to the remaining 15 µL of blastocoele fluid. Heat to 100 °C for 5 min and place on ice.
    NOTE: Analyzing proteins under non-reducing conditions is essential to analyze the formation of cleaved homodimeric or heterodimeric ligands but is not needed if only analyzing cleavage sites or levels of cleaved proteins. With a few exceptions, prodomain fragments are secreted as monomers. Furthermore, precursor proteins present in the embryo lysate are primarily unfolded, ER resident monomers. Thus, there is no need to analyze these products under non-reducing conditions.
13. Resolve precursor proteins and cleavage products by electrophoresis on 15% SDS/polyacrylamide gels.
14. Transfer proteins from the gel onto a PVDF membrane using electrophoretic transfer.
15. Incubate membranes with appropriate antibodies that recognize the prodomain and the ligand domain (or epitope tags inserted into those domains), wash and incubate with proper secondary antibodies. Visualize proteins with chemiluminescence or fluorescent imaging.