Technique to Target Microinjection to the Developing Xenopus Kidney

The following protocol has been approved by the University of Texas Health Science Center at Houston's Center for Laboratory Animal Medicine Animal Welfare Committee, which serves as the Institutional Care and Use Committee (protocol #: HSC-AWC-13-135).

1. Identification and Selection of Blastomeres for Kidney-targeted Injections 

1. Prior to generating embryos, use the Normal Table of Xenopus Development²¹ to understand the orientation of the early cell divisions in the embryo. Alternatively, access diagrams of the early developmental stages of Xenopus on Xenbase¹¹.
2. Access the interactive Xenopus cell fate maps on Xenbase¹¹ to select which blastomere will be targeted for microinjection.
3. Observe that the single cell embryo has a darkly pigmented animal pole and a vegetal pole, which is white and yolky. Note that a protective membrane, known as the vitelline envelope, covers the embryo.
4. Notice that the first cleavage typically occurs between the left and right sides of the embryo. These cells contribute equally to the pronephric lineage.
5. Note that the second cleavage divides the dorsal and ventral halves of the embryo, leading to a 4-cell embryo. The dorsal cells are smaller and have less pigment than the ventral cells (Figure 2A and Figure 3A).
   1. Identify the ventral blastomeres (V; the large, dark cells) on the left and right sides, which contribute more to the developing kidney than the dorsal (D; small, light cells) blastomeres (Figure 2A).
   2. If injecting into a 4-cell embryo, inject the left ventral blastomere to target the left kidney (Figure 3A and Section 3).
6. The third cleavage bisects the animal and vegetal sides, resulting in an eight-cell embryo. At this point, there are four animal blastomeres [left and right ventral (V1) and dorsal (D1)] and four vegetal blastomeres [left and right ventral (V2) and dorsal (D2)] (Figure 2B and Figure 3B).
   1. Locate the ventral, vegetal blastomeres (V2). These blastomeres contribute more to the developing kidney any other cells at this stage (Figure 2B). To target the left kidney of an 8-cell embryo, inject into the left V2 blastomere (Figure 3B and Section 3).
7. Notice that the fourth and fifth cleavages bisect the animal and vegetal blastomeres. Two progeny are generated from each blastomere, resulting in a 16-cell embryo. The cells are named after their predecessor. For example, the V2 blastomere from the 8-cell stage gives rise to V2.1 and V2.2 progeny at the 16-cell stage (Figure 2C). The V2.2 cell at the 16-cell stage provides the majority of the cells contributing to the developing kidney.
8. Note the sixth and seventh cleavages result in a 32-cell embryo. Again, two progeny are generated from each blastomere, which are named following their predecessor. For example, the V2.2 blastomere from the 16-cell stage gives rise to V2.2.1 and V2.2.2 at the 32-cell stage. There is an alternative naming system at the 32-cell stage in which cells are identified in four rows as A, B, C, and D (from animal to vegetal), and in four columns as 1, 2, 3, and 4 (from dorsal to ventral). Thus, the V2.2.2 blastomere, which contributes the most to the developing pronephros, is called C3 under this alternative naming system (Figure 2D).

2. Preparation of Embryos

1. Prepare 50 ml of Dejelly Solution (2% cysteine, NaOH to pH 8.0).
2. Isolate both testes from a single male frog according to standard protocols¹⁴. Place the testes in a 60 mm Petri dish filled with 10 ml Testes Storage Solution (1x Marc's Modified Ringers [MMR; 0.1 M NaCl, 2 mM KCl, 1 mM MgSO₄, 2 mM CaCl₂, 5 mM HEPES pH 8, 0.1 mM EDTA]¹², 1% bovine serum albumin, 50 µg/ml gentamycin). Store the testes at 4 °C.
   Note: Testes can be stored for approximately 7 – 10 days at 4 °C, but fertilization efficiency will decrease the longer the testes have been stored.
3. Squeeze a female frog to obtain eggs according to standard protocols¹⁴. Collect the eggs in a 100 mm Petri dish. Pour off any excess water.
4. Cut off ¼ of a testis while it is in Testes Storage Solution using forceps and a razor blade. Transfer the piece of testis to the Petri dish containing eggs. Adjust the size of the testis portion used to account for the size of the testis, how long the testis has been stored, and how many eggs are to be fertilized. Generally use ¼ to 1/3 of a freshly dissected testis to fertilize one clutch of eggs.
5. Cut the testis portion into small pieces using forceps and a razor blade. Add enough 0.3x MMR + 30 mg/ml gentamycin to the Petri dish to cover the eggs. Swirl the MMR in the dish to mix.
6. Wait approximately 30 min for fertilization to take place at room temperature. Note that the animal hemisphere (the pigmented side of the embryo) will sit on top of the embryo upon effective fertilization. Then, remove the MMR from the Petri dish using a transfer pipette. Add enough Dejelly Solution to the dish to cover the embryos.
7. Over the next few minutes, gently swirl the dish intermittently. Vigorous shaking of the dish at this time can cause axis defects. The jelly coat on the embryos will dissolve, and the embryos will congregate in the center of the dish during swirling. Once the embryos are closely touching each other in the center of the dish, remove the Dejelly Solution with a transfer pipette. Do not leave embryos in the Dejelly Solution for longer than 5 minutes, or the embryos may be damaged.
8. Wash the dejellied embryos 3 – 5 times in 0.3x MMR + 30 mg/ml gentamycin by carefully pouring or pipetting off the MMR and filling the dish with new MMR. Do not remove all of the MMR from the dish, or the embryos may be damaged.
9. Remove any unfertilized eggs or pieces of testis from the Petri dish using a transfer pipette.
10. Incubate embryos between 14 and 22 °C.
    Note: Embryos grown at lower temperatures develop more slowly than embryos grown at higher temperatures. Timing of developmental stages can be found on Xenbase²².
    1. To space out their development, place half of the embryos from a single fertilization in a Petri dish kept at 14 °C, and the other half of the embryos in a Petri dish kept at 18 °C. This allows for two sets of injections into 4-cell or 8-cell embryos from a single fertilization.

3. Preparation of Injection Solutions and Microinjection of Embryos

1. Prepare the injection solution containing 0.01 ng/nl membrane-bound red fluorescent protein (MEM-RFP) mRNA⁹ while the embryos are developing to the 4-cell or 8-cell stage. Store the injection solution on ice until ready to inject.
2. Load a 7" replacement glass capillary tube into a needle puller, with the top of the replacement tube aligned with the top of the needle puller case. Set the heat #2 value to 800, and the pull value to 650. Press the "pull" button to pull the needle. This will create 2 needles from a single 7" glass capillary tube.
3. Snip off the tip of a pulled needle with a pair of Dumont forceps.
   Note: After pulling the needle, the tip is sealed shut and must be cut open. The closer to the point of the needle that it is cut, the smaller the diameter the needle tip will be. Although the diameter of the tip will not affect the injection volume with the microinjection system used here, a tip with a larger diameter is more likely to damage the embryo.
4. Slip the micropipette collet onto the back of the needle. Next, slip the large hole O-ring onto the back of the needle behind the collet.
5. Fill the needle with mineral oil using a 27 gauge hypodermic needle, being careful not to get air bubbles in the needle.
6. Slip the needle onto the plunger of the microinjector, seating the needle into the large hole of the white plastic spacer installed on the plunger. The plunger should have a small O-ring nearest to the body of the microinjector, followed by the white spacer, large hole O-ring, and collet. Secure the needle by tightening the collet. Pull gently on the needle to make sure that it is properly secured.
7. Press and hold the "empty" button on the microinjector control box until there are two beeps.
8. Pipette 3 µl of injection solution onto a piece of Parafilm. Insert the tip of the needle into the bead of injection solution on the Parafilm. Press and hold the "fill" button on the microinjector control box to draw the injection solution into the needle.
9. Fill a 60 mm Petri dish lined with 500 micron polyester mesh with 5% Ficoll in 0.3x MMR + 30 mg/ml gentamycin. Carefully pipette 20 – 30 4-cell or 8-cell embryos into the dish.
10. Using a hair loop¹⁴, manipulate the embryos so that the blastomere to be injected is facing the needle. To target the left kidney, line up the embryos so that the left ventral blastomeres of 4-cell embryos or the left V2 blastomeres of 8-cell embryos face the needle.
11. Inject 10 nl of injection solution into the selected blastomere of each embryo in the dish.
    Note: The mesh at the bottom of the Petri dish stabilizes the embryos and prevents them from rolling, allowing them to be injected without the use of a hair loop for stabilization.
12. Transfer injected embryos into wells of a culture plate that have been filled with 5% Ficoll in 0.3x MMR + 30 mg/ml gentamycin. Incubate the injected embryos at 16 °C for at least one hour to allow the injected blastomeres to heal.
13. Transfer the healed embryos into wells of a new culture plate that have been filled with 0.3x MMR + 30 mg/ml gentamycin by stage 9 (prior to gastrulation).
14. Incubate the embryos at 14 – 22 °C until the embryos reach stage 38 – 40²¹.

4. Fixation and Immunostaining of Embryos

1. Prepare 50 ml of MOPS/EGTA/Magnesium sulfate/Formaldehyde Buffer [MEMFA: 100 mM MOPS (pH 7.4), 2 mM EGTA, 1 mM MgSO₄, 3.7% (v/v) formaldehyde].
2. Using a transfer pipette, put 10 – 20 stage 38 – 40 embryos in a glass vial. Add 10 µl 5% benzocaine in 100% ethanol to the vial and invert vial to mix. Wait 10 min to anesthetize the embryos.
3. Remove the MMR from the vial using a glass pipette. If processing multiple vials at the same time, the vials can be held upright in a 24-well cell culture plate.
4. With a glass pipette, fill the vial with MEMFA. Place the vial on a three-dimensional rocking platform for 1 hr at room temperature.
5. Remove the MEMFA from the vial using a glass pipette. Fill the vial with 100% methanol. Place the vial on a three-dimensional rocking platform for 10 min at room temperature. Repeat this wash step one more time, and store the embryos in 100% methanol overnight at -20 °C.
6. Prepare 1x Phosphate Buffered Saline-Bovine Serum Albumin-Triton (PBT): 1x PBS, 2 mg/ml bovine serum albumen, 0.1% Triton X-100.
7. Prepare the primary antibody solution: 1x PBT with 10% goat serum with a 1:5 dilution of mouse monoclonal 4A6 antibody (to label the membranes of the intermediate, distal and connecting tubules²⁰), a 1:30 dilution of mouse monoclonal antibody 3G8 (to label lumen of the proximal tubules²⁰), and a 1:250 dilution of rabbit polyclonal RFP antibody (to label the MEM-RFP tracer). Store at 4 °C.
8. Prepare the secondary antibody solution: 1x PBT with 10% goat serum, 1:500 Alexa 488 goat anti-mouse IgG (stock concentration 2 mg/ml; to label 4A6 and 3G8), and 1:500 Alexa 555 goat anti-rabbit IgG (stock concentration 2 mg/ml; to label the MEM-RFP tracer). Store at 4 °C, covering the tube in foil to protect from light.
9. Alternatively, collect the primary and secondary antibodies after staining, and save at 4 °C for reuse in future experiments. If the antibody is to be saved for reuse, add 0.01% sodium azide.
10. Immunostain the embryos using established protocols¹⁸.

5. Visualization of Embryos and Analysis of Targeted Pronephric Tissue 

1. Screen the immunostained embryos to verify that the correct blastomere was injected by viewing the fluorescence of the tracer under a fluorescent stereomicroscope at 1X (to view whole embryo) and 5X (to view kidney) magnification. Place embryos in a multi-well glass plate with the wells filled with 1x PBT using a transfer pipette with the tip cut off. Manipulate the embryos with a hair loop. Use only embryos which have the co-injected tracer present in the pronephros on the left side of the embryo (Figure 3C, F and Figure 4C, F) for gene overexpression or knockdown analysis.
2. Alternatively, clear the embryos in Murray's Clear (2 parts benzyl benzoate: 1 part benzyl alcohol) by placing the embryos in a glass vial and filling the vial with Murray's Clear. Visualize the embryos using a glass well plate.
   Note: Murray's Clear is an organic solvent, and should be handled with care. Wear gloves, and only use glass vials and pipettes with Murray's Clear.
3. Store embryos at 4 °C in 1x PBT for 2 – 3 weeks. For long-term storage of embryos, dehydrate the embryos by washing two times in 100% methanol at room temperature for 10 min. Store the embryos at -20 °C in 100% methanol.