Materials

1. Wild or mutant strains of Drosophila.

2. Egg collection plates (per 1 L): Dissolve 22.5 g of Bacto-agar (Difco) and 25 g of sucrose in 750 mL of boiling water. Add 250 mL of apple juice. Cool to approx 60°C and pour into Petri dishes (50 x 9 mm) halfway without air bubbles. Let the contents harden, cover and store at 4°C. Bring to room temperature before use.

Fig. 8. Feulgen-stained whole-mount preparation from the EGFR mutant (torpedo) animal 18 h after pupariation showing a conspicuous reduction in the number of cells in the (A) anterior dorsal (ADN) and posterior dorsal (PDN), (B) ventral (VN), and (C) spiracular (SN) nests. Compare the appearance of these nests to those of the wildtype animal of the same stage of development in Fig. 3. (From ref. 9, reprinted with permission of Springer-Verlag.)

Fig. 8. Feulgen-stained whole-mount preparation from the EGFR mutant (torpedo) animal 18 h after pupariation showing a conspicuous reduction in the number of cells in the (A) anterior dorsal (ADN) and posterior dorsal (PDN), (B) ventral (VN), and (C) spiracular (SN) nests. Compare the appearance of these nests to those of the wildtype animal of the same stage of development in Fig. 3. (From ref. 9, reprinted with permission of Springer-Verlag.)

3. Drosophila Ringer's solution (27): 130 mM NaCl, 4.7 mM KCl, 1.9 mM CaCl2. Dissolve 7.5 g NaCl, 0.35 g KCl, and 0.21 g CaCl22H2O in 1000 mL of distilled water. Store at 4°C in a stoppered bottle.

4. Kahle's fixative: Mix 12 mL of filtered formalin, 32 mL of absolute ethanol, 2 mL of glacial acetic acid, and 60 mL of water. Store in a stoppered bottle at room temperature.

5. Pasteur pipets, small paintbrushes, Sharpie pens.

7. Straight iridectomy scissors.

Fig. 9. Feulgen-stained whole-mount preparation of the abdominal integument of an EGFR mutant (torpedo) pharate adult, 25 h after pupariation. The photo montage of the right dorsal histoblast nests of the second to sixth segments shows a range of defects as a result of the mutation. The anterior dorsal and posterior dorsal nests of segments 2 and 3 have fused together, show a normal number of cells, and have started to spread in all directions, replacing the larval epidermal cells, as in the wild type at this stage of development. However, the dorsal nests of segments 4, 5, and 6 contain fewer cells, still remain apart, and are yet to spread actively. (From ref. 9, reproduced with permission of Springer-Verlag.)

Fig. 9. Feulgen-stained whole-mount preparation of the abdominal integument of an EGFR mutant (torpedo) pharate adult, 25 h after pupariation. The photo montage of the right dorsal histoblast nests of the second to sixth segments shows a range of defects as a result of the mutation. The anterior dorsal and posterior dorsal nests of segments 2 and 3 have fused together, show a normal number of cells, and have started to spread in all directions, replacing the larval epidermal cells, as in the wild type at this stage of development. However, the dorsal nests of segments 4, 5, and 6 contain fewer cells, still remain apart, and are yet to spread actively. (From ref. 9, reproduced with permission of Springer-Verlag.)

8. Sharp, steel dissecting needles.

9. Small Stender dishes (36 x 19 mm).

10. Glass depression slides.

11. Precleaned RITE ON (frosted at one end) microslides (Fisher Scientific).

12. No. 1 Glass cover slips (22 x 22 mm2) (Fisher Scientific).

13. Small plastic Petri dishes (50 x 9 mm)

14. Filter paper (42.5 mm in diameter) to line the Petri dishes.

16. Schiff's Feulgen reagent (28) (see also Chapter 7): Put 1 g of basic fucshin in a 500-mL Erlenmeyer flask. Add 200 mL of boiling distilled water to dissolve the stain (see Note 1). Stir for 5 min, cap with aluminum foil, and cool to exactly 50°C. Filter and add 20 mL of 1 N HCl to the filtrate. Cool to 25°C and add 1 g of sodium metabisulfite (Na2S2O5). Stir well. Keep the solution in a stoppered and aluminum-foil-covered bottle in the dark for 20 h at room temperature. The solution should appear straw colored after this period. Add 1 g of activated charcoal and stir for 2 min. Filter; the filtrate should be clear (see Note 2). Store the clear filtrate in another stoppered bottle covered with aluminum foil at 4°C. This can be used as long as it remains colorless. Bring the solution to room temperature before use.

17. 6 NHCl: Add 50 mL of concentrated (12 N) HCl to 50 mL of distilled water. Stir.

18. Bleaching solution: Make stock solutions of 100 mL each of 10% potassium bisulfite (K2S2O5) and 1 N HCl in distilled water. Just before use, add 5 mL K2S2O5 to 90 mL of distilled water and stir well. To this, add 5 mL of 1 N HCl and mix well. Keep in a stoppered bottle.

20. Xylene.

21. Permount mounting medium (Fisher Scientific).

22. Water mounting medium (Fisher Scientific).

24. 1 N NaOH: Dissolve 4 g NaOH in 100 mL of distilled water. Store in a bottle with a nonglass stopper.

25. Small hot plate (10 cm in diameter).

3. Methods

1. Keep fly stocks in an incubator maintained at 25 ± 1°C and 65% relative humidity.

2. Transfer well-fed adults from a stock culture food bottle to a fresh food bottle (without live yeast grains), cap it with an egg collection plate, and make two precollections of eggs of 1 h each using separate plates. This procedure will remove older eggs retained in the oviducts of the flies. Replace the second precollection plate with a fresh one and collect eggs for 30 min. This will allow emergence of a synchronized population of larvae from these eggs. Repeat this step until one gets enough eggs.

3. Keep the eggs in the plates for 21 h. Remove any prehatched larvae and collect freshly hatched ones for the next 10 min and transfer them to a new vial of food to continue further development. Under these conditions, the first, second, and third instars last 0-24, 24-48, and 48-96 h after hatching from the egg respectively (see Note 3).

4. Collect freshly pupariated animals (white puparia) from the wall of the culture vial, 96 h after hatching of first instar larvae.

5. Transfer white puparia with a wet paintbrush to Petri dishes lined with moistened filter paper and allow to develop to desired stages of pharate pupal and pharate adult development. The pupal molt and adult emergence occur 12 and 96 h after puparium formation, respectively.

3.1. Whole-Mount Preparation of Larval Integument

1. To make whole-mount preparations of the larval integument, collect cleaned larvae of appropriate age (see Note 4) and place them in a Petri dish lined with moist filter paper.

2. Keep a Stender dish half-filled (2 mL) with distilled water and allow the water to reach 50°C on a hot plate.

3. Transfer, using watchmaker's forceps, a larva to this hot water and leave it there for 5-10 s so that it is killed and straightened.

4. Immediately, transfer this larva to a depression slide containing a few drops of Ringer's solution.

5. Under the dissecting microscope (100 x) keep the larva on its back and hold its anterior end (with the black mouth hooks) with forceps and make a transverse cut with iridectomy scissors immediately posterior to the holding point. Similarly, make a transverse cut at the posterior end, anterior to the posterior paired spiracles.

6. Hold the anterior cut end of the larva with forceps firmly and make a clean mid-ventral longitudinal slit starting from the posterior to the anterior end.

7. Continue to hold the anterior end of the larva with forceps, and using another forcep carefully remove all the major internal organs (digestive, nervous, and reproductive systems and the extensive sheets of fat body) as much as possible without disturbing and damaging the integument and its attached muscles.

8. Transfer the carcass to a Stender dish half-filled with Ringer's solution. Holding the carcass to the bottom of the dish, flush out the remaining tissue debris by carefully squirting Ringer's solution several times at the inner surface of the integument.

9. Remove the medium with the floating debris and repeat the cleaning process outlined above (step 8) with fresh Ringer's solution. Repeat this step six to eight times so that all possible loose debris is removed from the integument, resulting in a clean filet containing the cuticle, LECs, histoblasts, and muscle fibers attached to the body wall.

10. Transfer quickly the clean filet to another Stender dish containing a drop of Kahle's fixative with fast green (see Note 5). Hold the filet to the bottom for 2 min so that it remains flattened (see Fig. 1 for the appearance of a representative extended segment) and submerged while being fixed.

11. Gently add more fixative to the dish and cover it and leave it for 12-24 h at room temperature. Make sure the filet is immersed completely in the fixative.

12. Carefully remove the fixative and replace it with decreasing concentrations of ethanol, starting with 70% and finally with distilled water. Keep the tissue in each of these solutions for 5 min.

13. Remove the distilled water and replace it with 2 mL of 6 N HCl for 10 min for hydrolysis. Replace the acid and rinse briefly with distilled water once.

14. Remove the water immediately and add 2 mL of Schiff's reagent and cover the dish. Keep this in a lightproof box for 90 min at room temperature.

15. Remove the Schiff's reagent and quickly add the bleaching solution. After 2 min, replace with fresh bleaching solution and repeat twice more.

16. Replace the bleaching solution with filtered tap water. Remove the water after 2 min and repeat this 10 times. After the last rinse, keep the tissue in filtered tap water for 30 min. Make sure that during all of these steps the tissue remains submerged in the medium.

17. At the end of this period, check nuclear staining in the tissues of the integument under the dissecting microscope. The nuclei of LECs, histoblasts (see Fig. 2A,B), and muscles should appear as dark magenta dots.

18. Replace tap water with 30%, 50%, and 70% ethanol and keep the stained tissue in each of the above for 3 min.

19. If counterstaining of the cytoplasm of the tissue is needed, replace the 70% ethanol with fast green stain for 1-2 min.

20. Transfer the integument to a fresh dish containing 90% ethanol and process it through 100% ethanol twice, keeping it for 5 min in each of these solutions.

21. While it is in the last change of 100% ethanol, add an equal amount of xylene and swirl the solutions to mix well. Leave the tissue in it for 5 min.

22. Replace the ethanol-xylene mixture with fresh xylene and leave the tissue for 5 min; repeat this twice.

23. Place a small drop of Permount mounting medium in the center of a microslide. Gently and carefully introduce the dehydrated and stained integument into this drop by holding the integument on one end and slowly inserting it into the medium at a slant. Gently sway the tissue two or three times in the medium so that the carried-over xylene mixes with the Permount.

24. After a minute, transfer the integument to a drop of Permount on a labeled slide. Gently push down the specimen to the bottom of the medium with the cuticular surface facing upward (see Note 6).

25. Hold the cover glass on its edge and bring it down slowly so that its middle area touches the mounting medium first, and taking care not to trap air bubbles.

26. Press the top of the cover glass gently with the tip of a dissecting needle, so that the integument remains flat at the bottom and allows the medium to fill under the surface of the cover glass up to its edges.

27. Keep the slide on a flat surface to dry. Carefully place a brass bar in the center of the cover glass for 24 h. This allows the specimen to remain flat.

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