An Antibody Uptake Assay for Imaging Notch/DeltaD Signaling in Zebrafish Radial Glia Progenitors

1. Preparation of zebrafish embryos

1. Set up fish crossing tanks in the afternoon before 5:00 p.m., with one female wild-type fish and one male Tg [ef1a:Myr-Tdtomato] fish by using dividers to separate them in each tank.
2. Remove the dividers before 11:00 a.m. from all crossing tanks the next morning. Keep quiet and do not disturb the fish while they are mating. Spawning typically occurs within 30 min after removing the dividers. The fertilized eggs remain on the bottom of the tank.
   1. Collect fertilized eggs from the tanks with a mesh filter. Transfer the eggs by washing them off into a Petri dish full of egg water and examine the embryos under a dissecting microscope at a 10x to 20x magnification.
3. Transfer the fertilized embryos to a clean Petridish containing approximately 20 mL of embryo medium (100 mL of 1,000x stock solution contains 29.4 g of sodium chloride (NaCl), 1.27 g of potassium chloride (KCl), 4.85 g of calcium chloride, dihydrate (CaCl₂.2H₂O), and 8.13 g of magnesium sulfate heptahydrate (MgSO₄.7H₂O)) with large-bore glass Pasteur pipettes. Keep the embryos at 28 °C. Keep 50 fertilized embryos per Petri dish with 30 mL of embryo medium at 28.5 °C.  
   ​NOTE: One pair of fish normally produces 100-300 embryos, and the fertilization and survival rate of healthy embryos is over 95% for each mating.
4. The next morning, take the embryos out of the incubator at 8:00 a.m., when the embryos have reached the developmental stage of ~18-20 h post-fertilization (hpf). Observe them under an epifluorescence microscope at a 20x magnification, using white light at first. At that time, the zebrafish embryos are at the 20-somite stage.
   1. Discard dead embryos that are cloudy or ruptured under the microscope using a glass pipette.
5. Turn on the fluorescent lamp and choose the red fluorescent protein (RFP) filter setting of the microscope. Then, select the embryos with strong red fluorescence and transfer them to a new dish with egg water.
   1. Dechorionate the embryos manually with two fine forceps (the tips of forceps should be sharp and undamaged) under white light (Table of Materials).
   2. Hold the chorion with one pair of forceps and make a tear in the chorion with the other forceps. Open the tear carefully using the forceps, and make it large enough for the embryo to pass through by gently pushing the embryo with the tips of the other forceps.
6. Transfer the dechorionated embryos to a new Petri dish with fresh embryonic medium for a quick rinse before microinjection.

2. Preparation of microinjection

1. Use the capillaries (1.2 mm outer diameter (OD), 0.9 mm inner diameter (ID), with filament) for pulling fine injection needles on a puller. Design and optimize the pulling program according to the handbook.
2. Open the tip of the needle with forceps under a stereo-dissection microscope. Make the diameter of the tip around 10 µm and the taper angle around 30°.
3. Conjugate the mouse monoclonal anti-Dld antibody with the anti-Mouse-IgG-Atto647N before injection.
   1. For each injection experiment, mix 0.5 µL of the anti-Dld antibody (0.5 mg/mL) with 2 µL of the anti-Mouse-IgG-Atto647N antibody (1 mg/mL) by pipetting 5-10 times. Then, incubate at room temperature for at least 30 min (or on ice for 2-3 hours).
   2. After incubation, add 2.5 µL of blocking buffer (10 mg/mL mouse IgG) and 0.5 µL of 0.5% phenol red to the antibody mixture and pipette 10x to mix thoroughly to block any unconjugated antibodies remaining in the mixture.
      NOTE: For each trial, prepare one extra mixture without the anti-Dld antibody and use it as a control.
4. Prepare 1% low-melting point agarose in the embryo medium. Heat the agarose-containing mixture at 70 °C till the mixture turns transparent.
   1. Aliquot the agarose solution in 2 mL microcentrifuge tubes. Keep the aliquots in a heat block at ~40 °C.
5. Rinse the embryo in the tube containing 1% low-melting point agarose for 3 s.
6. Place the embryo on an inverted plastic Petri dish lid together with individual drops of agarose (~30-40 µL) to mount each embryo separately on the lid as shown in Figure 1. Lay the embryos flat laterally in the agarose and keep this position until the agarose has solidified at room temperature.
7. Mount 12 embryos in three rows one by one as above. Cover all the mounted embryos in the agarose with egg medium.

3. Microinjection

1. Put the embedded embryos under the stereomicroscope and cover the agarose with egg water.
2. Set the air pressure injector together with micromanipulators, placing them close to the microscopes as shown in Figure 1. Use the steel gas cylinder containing gaseous nitrogen (N₂) under high pressure as the air resource.
   1. Open the gas valve only after the embryos have been mounted in the agarose. Then, front load the prepared glass needle with 2 µL of antibody mixture on the micromanipulator, as shown in Figure 1, when using the front fill module of the microinjector.
   2. Tune the input pressure to ~80-90 psi, and the injection pressure to ~20 psi.
3. Calibrate the injection volume by using a micrometer under the microscope, as described previously. Set the tune time duration to be from 10 ms to 120 ms, according to the size of needle opening. Deliver each pulse of injection by tapping the paddle. Tune the injection volume of each delivery to ~4-5 nL.
4. Poke the tip of the microinjection needle through the dorsal roof plate of the hindbrain posterior to the r0/r1 hinge point, and inject about 10 nL (two or three pulses) of antibody mixture without hitting the brain tissue. Observe the flowing of red fluids in the brain ventricle.       
   NOTE: The hindbrain ventricle is posterior to the midbrain hindbrain boundary. The injected phenol red-containing antibody mixture fills up the brain ventricle from the hindbrain to the forebrain immediately by diffusing with cerebrospinal fluid.
5. After injection, remove the tip of the needle from the embryo swiftly by rotating the knob of the micromanipulator. For a successful injection, the red dye of the injected mixture remains in the brain ventricle stably without leaking into the surrounded agarose.
6. Move the mounting plate under the microscope to locate another mounted embryo at a suitable position for repeats.
   1. After injecting six to eight embryos, peel the agarose with a microsurgical knife to release the embryos from the embedded agarose. Transfer the microinjected embryos to a fresh dish with 30 mL of embryo medium, and place them at room temperature for the next steps.

4. Mounting and time-lapse live imaging

1. After 30 min, transfer the selected embryos to 10 mL of embryo medium. Add 420 µL of tricaine stock (4 mg/mL) to 10 mL of embryo medium to anesthetize the embryos.
2. To mount the embryos, prepare 0.8% low melting point agarose containing the same concentration of tricaine in the tube. Keep the aliquots in a heat block at ~40 °C.
3. Use a glass pipette to immerse the injected embryos in the warm agarose for 3 s. Then, immediately remove the embryos from the agarose with the same glass pipette and place the embryos on the center of 35 mm glass bottom culture dishes with a drop of agarose from the tube. Place only one embryo per drop on the glass.
4. Orientate the embryos gently with a fiber probe or loading tip to keep the dorsal side of the embryonic brain as close to the glass bottom as possible. Tune the embryo position gently to extend the embryo without curling as the agarose solidifies gradually.
5. Afterward, check the embryo position by flipping the glass bottom dish over. Ensure that the whole dorsal forebrain with the correctly mounted embryos can be seen under the microscope.
6. Add 2-3 mL of 28.5 °C preheated embryonic medium containing tricaine to cover the embryo. Place the dish properly on the temperature-controlled stage of the confocal microscope, as shown in Figure 1. Adjust the temperature of the imaging chamber to be at 28.5 °C. The embryo is now ready for imaging.