Dissecting Cell-Autonomous Function of Fragile X Mental Retardation Protein in an Auditory Circuit by In Ovo Electroporation

Eggs and chicken embryos were handled with care and respect in accordance with the animal protocols approved by the Jinan University Animal Care and Use Committee.

1. Egg and plasmid preparation

1. Egg preparation
   1. Obtain fresh fertilized chicken eggs (Gallus gallus) from the South China Agricultural University and store at 16 °C before incubation. For optimal viability, set all eggs for incubation within a week of arrival.
   2. Place eggs horizontally and incubate at 38 °C for 46-48 h until Hamburger and Hamilton (HH) stage 12²⁵ for neural tube transfection, or for 54-56 h until HH13 for otocyst transfection. Because the animal pole of the egg is lighter than the vegetal pole, horizontal placement positions the embryo on the top of the egg for easy manipulation. Use caution to keep the egg horizontal in this and all following steps.
   3. The location of the embryo appears as a dark area on the eggshell when casting light from the bottom of the egg using a flashlight. At the desired HH stages, use this flashlight method to mark the location of the embryo on the eggshell with pencil.
   4. Wipe the eggs with gauze containing 75% ethanol. Drill a hole in the pointy end of the eggs using the tip of scissors and remove 2 mL of albumin with an 18 G needle syringe. Ensure the hole is only large enough to allow needle insertion. Wipe away any leaking albumin with gauze and seal the hole with clear tape.
   5. To minimize cracks and to prevent falling shell debris during windowing, cover the top of eggs with clear tape, centering on the pencil-marked dark area.
2. Plasmid DNA extraction
   1. Clone chicken Fmr1 shRNA using a transposon-based Tet-on system as described previously²⁰. This system contains three plasmids: pCAGGS-T2TP, pT2K-CAGGS-rtTA-M2, and pT2K-BI-TRE-EGFP-Fmr1 shRNA, providing a drug-inducible vector system by which the expression of Fmr1 shRNA and EGFP is silenced after transfection and can be turned on by doxycycline (Dox) induction.
      ​NOTE: These plasmids are not currently available in a repository. The authors agree to provide the plasmids upon reasonable request.
   2. Extract plasmid DNAs using an endotoxin-free preparation kit according to manufacturer's instruction and precipitate by adding 1/15 volume of 7.5 M sodium acetate and 1 volume of 100% isopropanol.
   3. Precipitate plasmids at -20 °C overnight and centrifuge at 13,000 x g for 10 min. Dissolve the pellet with the Tris-EDTA buffer provided in the kit to a final concentration of 4-5 µg/µL. Plasmids can be stored at -20 °C for 12 months without reduced transfection efficiency.
   4. On the day of electroporation, thoroughly mix 2 µL of each plasmid in a sterile centrifuge tube using a pipette tip. The resultant 6 µL volume of the plasmid mixture is enough to electroporate three dozen eggs. To help visualize the plasmid during electroporation, add 1 µL of 0.1% fast green (prepared in sterile ddH₂O) for every 6 µL of DNA mixture²⁶, which turns the plasmid mixture blue.

2. In ovo electroporation

1. Egg windowing and embryo identification
   1. On the day of electroporation, remove the eggs from the incubator, one dozen eggs at a time. Keeping eggs out of their incubator for more than 1 h introduces developmental variations and reduces viability.
   2. Wipe all surgery tools with gauze containing 75% ethanol.
   3. Place each egg in a custom-made foam holder. Wipe the tape-covered area with 75% ethanol and cut a window (1-2 cm²) around the circumference of the pencil mark using a small pair of scissors (Figure 1A). When cutting, hold the scissors flat to avoid damaging the embryo underneath.
   4. Place the windowed egg under a stereomicroscope with a 10x eyepiece and 2x zoom and identify the embryo. Adjust the angle and brightness of the light source for better visualization.
      NOTE: It takes practice and experience to visualize the embryo and identify the neural tube at this stage.
      1. For beginners, use the following optional ink injection to facilitate embryo identification. Prepare 10% Indian ink solution in 0.01 M phosphate-buffered saline (PBS) and autoclave in advance. Fill a 1 mL syringe with the ink solution and then fit a 27G needle. Bend the needle to a 45° angle with forceps.
      2. Under the microscope, carefully poke from the edge of the area opaca and insert the needle beneath the embryo. Inject ~50 µL of ink, which will diffuse below the area pellucida for embryo visualization. The ink will form a dark background for a clear visualization of the embryo.
         NOTE: Expel the air from the syringe before insertion and injection. When skilled at visualizing, avoid the ink injection step as it reduces the survival rate.
2. Neural tube injection and electroporation
   NOTE: This procedure is performed at HH12 for transfecting NM neurons in the brainstem.
   1. Pull glass capillaries (~1 mm in diameter and 100 mm long) into pipettes using a pipette puller. Under a dissecting microscope, carefully open the tip of the capillary needle to 10-20 µm in diameter with forceps. Store the pipettes (usually 5-10 at one time) in a storage box until use.
   2. Fill a capillary pipette with 0.5-1 µL of the plasmid mixture by applying negative pressure through a rubber tube at the end of the pipette. This can be achieved by a syringe or air pump.
   3. Place the egg under the microscope so that the embryo is vertically oriented with the tail near to you (or horizontally for injecting capillary pipettes using a three-axis manipulator). Hold the capillary pipette with one hand or with the three-axis manipulator and drive the tip of the pipette to the r5/6 in a tail-to-head direction.
      NOTE: The most anterior hindbrain area is r1 and each subsequent bulge along the neural tube is an individual rhombomere. R5/6 is at the same anteroposterior level as the otocyst, which is a readily recognizable cup-like structure (Figure 1B-C).
   4. Gently poke the tip through the vitelline membrane and into the dorsal neural tube and then withdraw the pipette a bit so the tip is in the lumen of the neural tube. Inject the plasmid mixture by applying air pressure until the tinted plasmid diffuses fully into r5/6 and extends into r3 and r4.
      NOTE: It is not necessary to make a slot on the vitelline membrane for injection.
   5. Check for a successful injection, which is achieved when the blue plasmid solution rapidly diffuses down the neural tube without leaking (Figure 1B-C). When leaking occurs, the blue quickly fades.
   6. Immediately after the injection, place a platinum bipolar electrode on either side of the neural tube (Figure 1D). Deliver two pulses of 12 V and 50 ms duration with an electroporator. Observe air bubbles at the ends of the bipolar electrode, with more on the negative side.
   7. Check for successful electroporation, which is achieved when the tinted plasmid mixture enters the neural tube tissue near the positive side of the electrode. After electroporation, carefully remove the bipolar electrode.
   8. Cover the window on the eggshell with a piece of transparent film that is pre-cut into 2 in squares and sprayed with 75% ethanol. Place the egg back into its incubator.
   9. Clean the bipolar electrode by delivering 10-20 pulses of 12 V and 50 ms duration in saline before proceeding to the next egg.
3. Otocyst injection and electroporation
   NOTE: This procedure is performed at HH13 for transfecting hair cells and AG neurons in the inner ear.
   1. At HH13 (which is ~embryonic day 2.5), the embryo has turned so the right side of the head faces up. Under the microscope, place the egg so that the embryo is vertical, with the tail near to you. Hold the capillary pipette and gently poke the right otocyst in a dorsolateral direction (Figure 2A).
      NOTE: At this stage, the otocyst appears as a small circular structure on the top of the body at the same anteroposterior level as r5/6.
   2. Inject the plasmid mixture with air pressure until the otocyst is filled with blue solution²⁷. Check for a successful injection, which is achieved when the blue plasmid mixture is confined within the otocyst and doesn't leak.
   3. Immediately after the injection, place the bipolar electrode on the otocyst as depicted in Figure 2B. Position the positive and negative sides anterior and posterior to the otocyst, respectively. Deliver two pulses of 12 V and 50 ms duration with the electroporator.
   4. Check for successful electroporation, which is achieved when the blue plasmid mixture enters the tissue of the otocyst near the positive side of the electrode. After electroporation, carefully remove the bipolar electrode. Cover the window on the eggshell with a transparent film and return the egg to the incubator.

3. Administration of Dox to initiate and maintain plasmid transcription

1. Preparation of the Dox solution
   1. Measure 100 mg of Dox powder under a chemical hood and dissolve it in 100 mL of sterile PBS to make a 1 mg/mL of working solution. Filter the solution with a 0.22 µm filter and store 1 mL aliquots at -20 °C. Protect from light^20,28.
2. Administration of Dox
   1. For full-strength gene editing, at 24 h before the desired age, thaw a Dox aliquot on ice. Drop 50 µL of Dox directly on the chorioallantoic membrane of the egg using a syringe penetrating the transparent film. Seal the needle hole with transparent film or tape after injection.
   2. To maintain the knockdown effect, administer Dox every other day until tissue harvest at the desired developmental stage.

4. Tissue dissection and sectioning

1. Brainstem
   1. For embryos at embryonic stage 3 (E3) and E6, open the eggshell and cut the associated membrane with scissors. Spoon the embryo out and immerse it in 4% paraformaldehyde (PFA) in 0.1 M phosphate buffer.
   2. For E9 embryos, open the eggshell, decapitate the embryo with scissors, and transfer the head to a silicon-bottomed plate filled with PBS. Pin the head down through the orbits and cut the top of the skull apart. Carefully place the forceps beneath the brain from both sides and elevate the entire brain from the skull with the shoulders of the forceps. Immerse the brain in 4% PFA.
   3. For E15 and E19 embryos, open the eggshell, decapitate the embryo with scissors, and place the head on a silicon-bottomed plate filled with PBS. Cut open the skin on top of the head to expose the skull.
   4. Incise through the skull vertically with a razor to separate the forebrain and the caudal brain. Immerse the caudal block in PBS, remove the top of the skull, and then take the brain off the skull.
   5. Pin the brain down through the optic lobe and separate the cerebellum and brainstem. Take care not to damage the auditory brainstem, which is located right below the cerebellum. Remove as much dura as possible from the brainstem and then immerse it in 4% PFA.
   6. Keep embryos and brain tissues in 4% PFA at 4 °C overnight except for E19 brainstems, which should be kept under the same condition for 24 h.
   7. Following fixation, dehydrate the tissue in PBS containing 30% sucrose until it settles, which usually takes 1-3 days, depending on the age.
   8. Section the brainstem (E15 and E19) at 30 µm at the coronal plane using a sliding microtome. Collect sections in PBS and store at 4 °C before immunostaining.
   9. For E3 and E6 embryos and E9 brainstems, section at 50 µm transversely. Collect sections in PBS and store at 4 °C. Mount the sections on gelatin-coated microscope slides before immunostaining. Collecting thicker sections for these ages facilitates tissue handling.
2. Auditory duct
   1. After removing the brainstems from E9, E15, and E19 embryos, the auditory duct, which is embedded in the temporal bone, is readily identifiable lying beneath the skull, and the temporal bone is easily separatable from the surrounding skull. For E9 embryos, fix the whole temporal bone in 4% PFA. For older embryos, remove the bony structure of the temporal bone to isolate the auditory duct and fix the auditory duct in 4% PFA.
   2. Keep all temporal bones and auditory ducts in 4% PFA at 4 °C overnight before tissue embedding.
   3. Perform tissue embedding as described. Prepare the embedding solution as follows: Soak 10% gelatin (from bovine skin) in cold water until the gelatin granules swell and settle (about 30 min). Heat the mixture to ~50 °C to dissolve the gelatin and add 20% sucrose into the gelatin solution and stir to dissolve. Store the gelatin-sucrose solution at 4 °C for up to a month.
   4. For use, warm the gelatin-sucrose solution at 37 °C. Soak the temporal bones/auditory ducts in the warm gelatin-sucrose solution on a 96-well plate at 37 °C until they settle, usually 30-60 min.
   5. Line the bottom of the wells of a 12-well plate with a strip of transparent film. Add a layer of warm gelatin-sucrose solution and wait until it solidifies. Transfer a temporal bone/auditory duct to each well.
   6. Implant the tissue with a second layer of warm gelatin-sucrose solution. Adjust the tissue's position so that it is in the center of the gel and the auditory duct is oriented horizontally. Carefully move the plate into a 4 °C refrigerator and wait until the gel is firm.
   7. Pull the transparent film strip to move the gel-tissue block out of the well. Trim extra gel into a square block and cut a corner of the block to identify the orientation. Wrap the gelatin block with a piece of foil, freeze on dry ice, and then store at -80 °C until sectioning.
   8. Section the block at 20 µm along the longitude of the auditory duct with a cryostat. Mount the sections directly on gelatin-coated microscope slides. Store the slides at -80 °C before immunostaining.
   9. Prior to immunostaining, immerse the slides in pre-warmed PBS (45 °C) for 5 min to dissolve the gelatin, and then wash the slides 3x with room temperature PBS to remove residual gelatin.

5. Immunostaining and microscope imaging

NOTE: Two types of immunostaining are performed depending on whether sections are mounted on slides or free-floating in PBS.

1. Immunostaining on slides
   1. Wash the slides 3x for 10 min each with PBS. Circle the region containing the sections with an oil pen to prevent liquid leaking during staining. Cover the sections with a blocking solution containing 5% normal goat serum in PBS and incubate for 30 min at room temperature.
   2. Dilute the primary antibodies (for the final concentration used refer to Table of Materials) in PBS containing 0.3% TritonX-100 and 5% normal goat serum. Remove the blocking solution and then cover the sections with the primary antibody solution. Incubate the slides at 4 °C overnight in a box containing a layer of distilled water at the bottom.
   3. Rinse the slides 3x for 10 min with PBS. Apply fluorescent secondary antibodies diluted at 1:500 in PBS containing 0.3% TritonX-100 on slides. Incubate the slides at room temperature for 4 h, shielded from light.
   4. Wash the slides 3x with PBS. Gently drop two drops of mounting medium on the slides and cover the sections with coverslips. Take care not to generate air bubbles between the coverslip and the slide.
2. Immunostaining for whole-mount embryos and free-floating sections
   1. Wash the embryos/sections with PBS 3x for 10 min each in a well plate. Dilute the primary antibodies in PBS containing 0.3% TritonX-100 and 5% normal goat serum in centrifugal tubes. Transfer each embryo/section into a tube with a glass hook and incubate on a shaker at 60 rpm overnight at 4 °C.
   2. Wash embryos/sections 3x with PBS for 10 min each in a well plate. Transfer each embryo/section into a dark centrifugal tube filled with fluorescent secondary antibody solution and incubate at room temperature for 4 h on the shaker.
   3. Wash the embryos/sections 3x with PBS in a well plate. Use a brush to mount the sections on gelatin-coated microscope slides in a 15 cm dish containing PBS. After the slides are dried slightly (~5 min), apply two drops of mounting medium and place a coverslip. Take care not to generate air bubbles between the coverslip and the slide. Keep the whole mount tissues in PBS for imaging.
3. Imaging
   1. Image the whole mount tissues in PBS in a dish with a silicon bottom containing carbon powder, which provides a black background. Capture and process images using a commercial image processing Olympus software package, as described previously²⁹.
   2. Image the sections on the slides with a confocal microscope as described previously²⁰. Image the sections at a single focal plane with 10x, 20x, and 63x objectives. Capture all images from the same animal using the same parameters. Take care to adjust the laser level and imaging acquisition settings to avoid signal saturation.
   3. Perform image processing using the Fiji software. For illustration, adjust image brightness, contrast, and gamma using a professional image editing software.