A. Embryo Dissection
1. Under a dissecting microscope, in a 35-mm culture dish containing grasshopper saline (150 mM NaCl, 10 mM KCl, 4mM CaCl2, 2mM MgSO4, 5mM TES, and 140 mM sucrose, pH 7.1), embryos are removed from their egg cases by puncturing the opposite end of the egg that contains the embryo to relieve pressure (the dark pigment at one end of the egg indicates the presence of an embryo); some yolk will be expelled from the egg.
2. Using micro scissors, the very tip of the end of the egg containing the embryo is cut off and the embryo is removed by applying gentle pressure to the center of the egg case. The embryo and some yolk will be expelled. Separate the embryo from the yolk and move embryo to a yolk-free area of the dish.
3. To allow access of culture media and reagents, the amnionic membrane is removed on the dorsal and ventral sides. This membrane is easiest removed by pinning the head or abdomen of the embryo with one set of forceps while running a second set of forcepts along the midline of the animal to tear and remove the amnion.
4. Whenever embryos need to be transferred, the best method is to use a Pasteur pipette and gently pipette the embryos to their new location. Because the embryos will stick to the inside of the glass pipette, it is best to coat the inside of the pipette with grasshopper saline [containing 0.3% bovine serum albumen (BSA) or some egg yolk from the dissection dish] by pipetting up and down a few times before pipetting the embryos. This should be done every time embryos are pipetted.
1. Prior to dissection, eggs are brushed clean in grasshopper saline and sterilized in 70% ethanol for 1 min followed by rinsing twice in sterile grasshopper culture medium [RPMI (Gibco-BRL), 4 yM 20-hydroxyecdysone, 0.4 mM CaCl2, 0.4 mM MgSO4, 2 units/ml insulin, 100 units/ml penicillin, 100 yg/ml streptomycin, 2mM L-glutamine, 0.45 mM sodium pyruvate, 1 mM oxaloacetic acid, 0.45% D-glucose, 0.12 M sucrose, and 10 mM TES, pH 6.9].
2. Place eggs in fresh sterile culture media for dissection under a dissecting microscope in a laminar flow hood.
3. Using sterile technique, embryos are dissected as described earlier, except that when culturing, embryos should be dissected in grasshopper culture medium instead of grasshopper saline.
4. After dissection, embryos can be placed in a well of a 96-well culture dish. To assess the Ti1 pathway, embryos are cultured for 24 h, but can be cultured for as long as 48 h, with a change of culture medium after 24 h. Embryos are cultured in a 30° C incubator (without CO2).
1. Embryos are dissected in grasshopper saline as described earlier.
2. Embryos are fixed by immersion in PEM-FA [0.1 M PIPES (pH 6.95) 2.0 mM EGTA, 1.0 mM MgSO4, and 3.7% formaldehyde] for 30min to 1 h.
3. Embryos are washed three times for 1 min and three times for 10 min in PBT [1x phosphate-buffeted saline (PBS), 0.1% Triton X-100, 0.1% BSA].
4. Embryos are incubated overnight at 4°C in primary antibody. Dilutions ranging from 1:500 to 1:1200 are appropriate for both rabbit and goat anti-HRP (horseradish peroxidase) (depending on individual lots; Jackson Immunoresearch).
5. After primary antibody incubation, embryos are again washed three times for 1 min and three times for 10 min in PBT.
6. Secondary antibodies are incubated for 1 h at room temperature, followed by three times for 1 min and three times for 10 min in PBT.
7. For mounting, we typically use the antifade reagent SlowFade (Molecular Probes).
8. In order to visualize the Ti1 pathway and the limb bud properly, further dissection of the embryos after labeling is employed. Embryos are pipetted gently on glass slides with a minimal amount of SlowFade antifade. Using forceps, separate the three pairs of limb buds gently by making incisions under each pair of limb buds across the width of the body and spreading out the pieces on the slide.
D. Preparation of Glass Bottom Culture Dishes for Microinjection and Time-Lapse Imaging
In order to culture and image filleted embryos, specialized culture dishes must be prepared in advance.
1. A 15-mm hole is bored into a 35-mm sterile polystyrene tissue culture dish using a drill press.
2. To ensure proper contact between the coverslip and the petri dish plastic, the edges of the hole are smoothed using sandpaper. The petri dish is then soaked briefly with 70% ethanol, rinsed with distilled water, and allowed to dry. Dishes can be sterilized further by overnight ultraviolet light exposure.
3. With the petri dish upside down, the outside of the hole is ringed with a melted mixture of beeswax and petroleum jelly (80:20), and then a rinsed round (22-mm diameter, No. 1) coverslip that has been pretreated with 5mg/ml poly-L-lysine (see Section V,E) is placed on top of the wax so that the coverslip covers the hole.
4. Because the wax hardens rapidly, it is typically necessary to place the upside-down petri dish in an 80° C oven for 3-5 min to allow the wax to melt and seal the coverslip. Dishes prepared in this manner allow for visualization of the Ti1 neurons at high magnification on a compound microscope.
E. Preparation of Poly-L-Lysine-Coated Coverslips
1. Using sterile conditions, one drop of filter sterilized 5 mg/ml poly-L-lysine is placed into a 35-mm tissue culture dish.
2. A sterile 22-mm-round coverslip (number 1) is placed on top of the drop of poly-L-lysine.
3. Another drop of poly-L-lysine is placed on top of the coverslip, followed by another coverslip, and this process is repeated until the desired number of coated coverslips is achieved.
4. To avoid evaporation, the bottom of the tissue culture dish is filled with poly-L-lysine and is wrapped in Parafilm.
5. The coverslips are incubated at 37° C overnight followed by storage at 4° C. In general, the longer the coverslips are incubated at 4° C, the more adherent the coverslips become.
F. Filleting Grasshopper Limb Buds to Gain Access to the Til Pathway
1. Sterilize and dissect embryos as described earlier and prepare a glass bottom dish.
2. Transect the embryo between the second and third limb pairs with forceps, retaining the third limb pair and tail for fillet. The transection can be made more anteriorly if it is desired to fillet the T1 or T2 limbs.
3. Using a glass pipette, transfer the embryo to the edge of the glass bottom dish filled with hopper saline.
4. At low magnification, pick up the embryo by the tail with forceps ventral side down and transfer onto the poly-L-lysine coverslip.
5. Let the tips of the limbs touch and attach, and then flip the abdomen 180° so that the ventral side is up. This ensures that the anterior surface of the limbs, and consequently the Ti1 neurons, is adjacent to the coverslip.
6. Dissection of the limb is carried out at high magnification using dark-field optics.
7. Using a tamp [a fire-polished pulled solid glass tube (1 mm diameter) that has been passed through the flame of a match to give it a smooth tip], ensure that the limbs are attached firmly to the coverslip by pressing them against the coverslip gently.
8. Using a sharp dissecting tip (a solid 1-mm-diameter glass needle that has been pulled on a gravity micropipette puller), a cut is made along the length of the limb down the center of the posterior epithelium (Fig. 5), being careful only to pierce the top epithelium. Each freshly pulled glass needle is sharp enough to finely cut two limbs. Pipettes should be repulled for additional use.
9. Using the tamp, roll out the epithelium flat on the coverslip and apply gentle pressure to the edges of the limb to promote adhesion to the coverslip
10. Changing to bright field, the mesoderm is discernible as an irregular mass of cells on top of a homogeneous layer of epithelium.
11. Using a micromanipulator, position a thin-walled, pulled micropipette (with a broken tip) over the limb and aspirate the mesoderm off using gentle suction with a mouth pipette (Fig. 5). Removal of all the mesoderm may require several passes.
12. Replace hopper saline with grasshopper culture medium.
Roll out Epithelium
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