Western Blot Analysis of RNAiTreated Cells

Western blotting, like that shown for Cnn in Fig. 2, is recommended to assay the efficiency and time-course of decay for each target protein.

1. Remove aliquots of S2 cells at different time points and place in 1.5-mL tubes. Remove an aliquot of the cells at time zero (500 |L; see Note 11), before dsRNA addition, and remove aliquots every day (or other time increments thereafter), for several days.

2. Pellet cells by centrifugation in a microfuge. Discard the supernatant and suspend in 50 |L 1X SDS-PAGE loading dye. Load 10 |L of the sample onto SDS-PAGE gel following heating to 95-100°C for 5 min. Samples can be stored at -20°C or -70°C.

3. Pour an SDS-PAGE minigel (see Note 12). For the resolving gel, mix acrylamide (7-15% final, depending on the size of your protein), 375 mM Tris-HCl, pH 8.8, 0.1% SDS, 1/1000 volume of 25% APS, 1/1000 volume TEMED. Pour gel immediately after the addition of TEMED, leaving about a 3-cm space for the stacking gel. Overlay with approx 100 |L water. Let polymerize for 1 h. For the stacking gel, mix acrylamide (4%), 125 mM Tris-HCl, pH 6.8, 0.1% SDS, 1/1000 volume of 25% APS, 1/1000 volume TEMED. Remove the overlay solution, then pour immediately and insert the comb. Let polymerize at least 30 min.

4. Separate the proteins by electrophoresis on an SDS-PAGE minigel.

5. Transfer to nitrocellulose membrane in gel transfer buffer using a cooled transfer chamber at 100 V for 1 h.

6. Place the membrane in 20 mL of blocking solution in a 9 x 9 cm square Petri dish or similar chamber. Incubate for 1 h at room temperature with gentle shaking.

7. Remove the blocking solution. Add primary antibody in 10 mL TBS-T and incubate for 1 h at room temperature (or overnight at 4°C).

8. Remove the antibody solution and wash the blot three times with 20 mL TBS-T for 5 min each.

9. Incubate with HRP-conjugated secondary antibody (1 : 10,000) in 10 mL TBS-T for 30 min.

10. Repeat step 8.

11. Treat the blot with chemiluminescence substrate reagent and expose to X-ray film for various times.

3.5. Staining of Cells

1. Treat the slides with poly-L-lysine as follows: Wash glass slides in water and wipe dry with a Kimwipe. Apply 50 |L of 1 mg/mL poly-L-lysine into each well on the slide and let sit for 45 min. Wash slides with water three times in Coplin jars. Let slides dry (see Note 13).

2. Apply 50 |L of cells to each well and let sit for 30 min.

3. Rinse cells briefly (2 s) in PBS and then place directly into -20°C methanol. For this, dip the slides into a Coplin jar containing PBS and place them into a Coplin jar with methanol that has been kept in the freezer. Incubate the slides in -20°C methanol for 10 min.

4. Remove the slides from -20°C and place into a Coplin jar with PBS. Rinse once with fresh PBS. The cells should appear as a film in the well. The cells should not be permitted to dry in any of the subsequent procedures.

5. Using a Kimwipe twisted into the shape of a probe, or using a cotton swab, blot the PBS from the region of the slide surrounding the well dry. This will prevent the antibody solution from spreading out from the well in subsequent procedures.

6. Apply the primary antibodies, diluted in PBS + 0.1% saponin + 5 mg/mL BSA, 50 |L per well. If DNA dyes such as propidium iodide or TOTO-3 are to be used, RNase A can be added at this step at a concentration of 50 |g/mL (see Note 14). We recommend using one combination of antibodies for all the samples on the same slide to prevent cross-contamination. For different antibody mixtures, use additional slides.

7. Incubate slides in a humid chamber for 1 h at room temperature, or overnight at 4°C. A simple humid chamber can be made by taking an empty pipet tip box, adding water into the box, and placing the slides onto the slotted tip holder.

8. Wash slides in a Coplin jar with three changes of PBS, 5 min each.

9. Apply secondary antibodies to the slides (see Note 15). First, blot the area around the wells dry as described in step 5. Add 50 ||L of secondary antibodies, diluted in PBS + 0.1% saponin + 5 mg/mL BSA, into the wells. Incubate for 1 h at room temperature in the dark.

10. Wash as in step 8; then, blot the slides dry as in step 5.

11. Apply 4 |L of Mountant to each well. Overlay a cover slip slowly and at an angle to prevent the inclusion of air bubbles under the cover slip (see Note 16). Fix coverslip to the slide with clear nail polish.

3.6. Imaging Cells by Confocal Microscopy

1. High magnification with a 60x or higher objective is required to image S2 cells effectively. These objectives require immersion in oil or water (see Note 17).

2. For confocal microscopy, use multiple excitation lasers to image multiple fluorophors. There are a variety of configurations available; some include lasers that produce lines typically at 488, 568, and 647 nm (argon-krypton), or 488, 568, and 633 nm (argon, krypton, and helium-neon (RedHeNe), or 488, 543, and 633 nm (argon, GreenHeNe, RedHeNe). These should all be compatible with three-color imaging like that shown in Figs. 3 and 4, where the (excitation peak wavelength/emission peak wavelength [in nm]) for FITC (490/ 520), TRITC (541/572), and TOTO-3 (642/660) allowed separation of all three emission signals.

3. S2 cells are small, approx 10 |im thick. Therefore, when a z-series is collected, a large stack of images will not need to be produced. Steps of 0.5-1.0 |im may be adequate for most purposes.

4. If bleaching becomes a problem, one method is to set up the imaging using only one of the fluorescent signals (the more robust) to view the cell. Then, turn on the other lasers when the images are being captured. This strategy reduces the bleaching of weaker signals or sensitive fluorophors.

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