The pattern of expression of your gene can give a clue as to its function. Most testis transcripts are made in primary spermatocytes and stored until use.
A transcript that persists along the testis is more likely to encode a protein needed late in differentiation. Early degradation of a transcript suggests an earlier function (see Fig 4).
3.4.1. Preparation of Probe
Essentially follow the labeling protocol given with the Roche RNA labeling mix, outlined as follows.
1. Linearize plasmid with a suitable restriction enzyme (usually one that cuts in the polylinker at the 5' end of the gene).
2. Set up a 20-|L reaction using 1 |g of linear template and the relevant RNA polymerase. A typical reaction contains 2 |L of 10X RNA labeling mix, 2 |L of 10X RNA polymerase buffer, 1 |g of template DNA in 14 |L of RNAse-free water, and 2 | L of RNA polymerase.
3. Allow the transcription reaction to continue for 2 h at 37°C, then stop the reaction by adding 2 |L of 0.5 M EDTA.
4. Hydrolyze the probe by diluting with distilled water to 100 |L and adding 100 |L of 2X carbonate buffer; incubate at 60°C. Probe hydrolysis times vary according to transcript length. The aim is to generate short RNA fragments (approx 100 bp) that will penetrate the tissue more easily. I allow 15 min per 500 bp (i.e., incubate a 2-kb probe for 1 h).
5. Neutralize by adding 200 | L of 2X neutralization buffer.
6. Precipitate the RNA product by adding 3 vol of ethanol and incubating at -20°C for at least 30 min.
7. Spin in a microcentrifuge 15 min, wash with 70% ethanol, dry the pellet, and resuspend in 200 |L of RNAse-free water. Store the probe at -70°C. For hybridization, use 0.5-1 |L per 100 ||L of hybridization buffer.
1. Dissect testes from young adults (0-1 d old) in testis buffer and transfer to a 1.5 mL tube (see Notes 2, 6, and 19).
2. Fix in 4% paraformaldehyde in HEPES buffer for 20-60 min; agitation is not required—simply leave the tube on its side.
3. Wash three times in PBST, 5 min each.
4. Incubate in 50 |ig/mL of proteinase K in PBST for 5-7 min (see Note 20).
5. Remove the proteinase K solution and stop the digestion by incubating the testes in 2 mg/mL glycine in PBST for 2 min.
6. Wash twice in PBST, 5 min each.
7. Refix in 4% paraformaldehyde in HEPES buffer for 20 min.
8. Wash three times in PBST, 10 min each.
11. Transfer testes into tissue culture inserts in a 24-well tissue culture plate (see Note 7).
12. Prehybridize in HB at 65°C for at least 1 h (see Notes 21 and 22).
13. Dilute the RNA probe (from Subheading 3.4.1.) in HB, heat denature at 80°C for 10 min, and briefly chill on ice.
14. Hybridize at 65°C overnight.
15. Wash at least six times, 30 min each, in HB at 65°C (see Note 23).
16. Wash once in 4 : 1 HB : PBST for 15 min at room temperature.
17. Wash once in 3 : 2 HB : PBST for 15 min at room temperature.
18. Wash once in 2 : 3 HB : PBST for 15 min at room temperature.
19. Wash once in 1 : 4 HB : PBST for 15 min at room temperature.
20. Wash twice in PBST, 15 min each.
21. Incubate overnight at 4°C in preadsorbed (see Note 1) alkaline phosphatase-conjugated antidigoxygenin antibody diluted 1 : 2000 in PBST.
22. Wash four times in PBST, 20 min each.
23. Wash three times in freshly made buffer HP, 5 min each.
24. Make staining solution in HP buffer by adding 4.5 |L of NBT and 3.5 |L of X-phosphate per milliliter.
25. Add the color reaction solution to the testes and leave to develop in the dark. The signal typically takes 10 min to 1 h, although for some transcripts incubation for several hours may be required. The staining needs to look quite dark, and purple not pink, at this stage to get good pictures at high magnification.
26. Stop the reaction by washing three times in PBST, 5 min each.
27. Dehydrate through an ethanol series: 10 min in each of 30%, 50%, 70%, 90%, and 100% (twice) ethanol. Transfer testes into a glass staining block (see Note 24).
28. Incubate 15 min in 1 : 1 ethanol : methyl salicylate, then in 100% methyl salicy-late (see Note 25).
29. Mount in GMM and observe with Nomarski optics.
1. To preadsorb the antibody, fix embryos using a protocol suitable for immuno-staining (see Chapter 9). Dilute the antibody 1 : 20 in PBST and incubate with the rehydrated fixed embryos for 2 h. Remove the antibody solution from the embryos and store at 4°C.
2. Newly eclosed males are used for all of these protocols, as they show the best morphology. To dissect testes from flies, place an anesthetized male next to a drop of testis buffer on the dissecting dish. Hold near the top of the abdomen with a pair of fine forceps in your left hand (or your right hand if you are left-handed). Grasp the external genitalia with the other pair of forceps and pull into the drop of buffer. The male genital tract, including testes, should come clear of the carcass. If it does not come clear, you will have to "fish" for the testes in the abdomen. Transfer the genital tract into a fresh drop of testis buffer and dissect the coiled testes and attached seminal vesicle from the rest of the tissues. If scoring for sperm motility, it is important to note that many males have no mature sperm for about 12 h after eclosion. To be sure of the absence of motile sperm from a mutant, keep males isolated from females for 3 d, then dissect their seminal vesicles.
3. Generally, it is best to cut halfway along the straight portion of the testis. The contents should partially spill out, and this can be encouraged by gently tapping on the slide. The diameter of the drop of buffer should be approx 7 mm to get good preparations under a 22 x 22-mm2 cover slip. Cysts of spermatocytes or spermatids should stay intact. The seminal vesicle from males that produce normal motile sperm will normally be slightly opaque. When it is opened, the sperm spill out. Motile sperm will show a shimmering effect visible even under the dissecting microscope.
4. Take pictures first, ask questions later. Photography can be done with either a digital camera or using black-and-white film. Squash preparations are only good for approx 20 min. After that, the cells are usually too flat and dead to observe. Figure 6 shows some typical squash artifacts.
5. Alternatively, preparations can be made for visualization with an inverted microscope by using slides with a hole cut in them, sealing a cover slip over the bottom of the hole with vaseline, and dissecting the male into halocarbon oil in the chamber thus generated. Under these conditions, the cells remain viable for at least 3 h (40,41). (See Chapter 3.)
6. To transfer testes, place a small drop of testis buffer in the lid, put the testes into this, add 600 |L of fix to the tube, close the lid, and mix. Testes stick to tweezers if you try to put them directly into fix.
7. Add testes and washes into inserts; remove by lifting up the insert and aspirating solution from the well (see Fig. 9). Be careful; sometimes the mesh at the bottom detaches. Check for loose testes before aspirating. Do not fix testes in these dishes, as they may stick to the mesh.
8. Blocking does not seem to be essential for whole-mount immunohistochemistry, but it may improve the staining with certain antibodies.
9. The optimal dilution for each antibody needs to be empirically determined. As a first guess try using it two to four times more concentrated than gives an acceptable signal on Western blots. Immunohistochemistry often works with the antibody more dilute than is needed for immunofluorescence. For example, if an antibody works well at 1 : 4000 on Western blots, try 1 : 2000 for whole-mount immunohistochemistry and 1 : 1000 for immunofluorescence on squashed preparations. Antibody incubations can generally be done for a few hours at room temperature or overnight at 4°C. The choice is usually governed by convenience, although some primary antibodies work better overnight at 4°C.
10. The Vectastain ABC kit gives a slightly stronger signal, which decreases the development time, and may be useful for some proteins with relatively low levels of expression. Substitute the following for step 9 of Subheading 3.2.1.: Incubate testes in 0.5X ABC reagent (50 |L solution A, 50 |L solution B, 5 mL PBS mixed 30 min before use) for 30 min. Go to step 10 of Subheading 3.2.1.
11. DAB is a potent carcinogen; wear gloves and inactivate DAB with bleach after use. Tablets from Sigma are 0.7 mg and should be dissolved in 1 mL of PBS.
Lowering the concentration of DAB to 0.35 mg/mL (i.e., one tablet in 2 mL) does not significantly compromise the staining.
12. To mount stained testes in 85% glycerol, first transfer to a glass staining block. Remove the PBS and replace with glycerol. Mix well with a tungsten mounted needle or tweezers. Leave the testes in glycerol for 15-30 min before transferring to a cover slip, cutting off accessory gland and picking up with a clean slide. Imaging stained tissue under Nomarski optics usually requires that the optics be somewhat compromised, because the stain rarely shows up well when the Nomarski is set up to give the most structural information. Photograph using a tungsten balanced color slide film or a daylight correction filter and normal color slide film.
13. Write the genotype on the slide with pencil or a diamond pen; marker dissolves in ethanol! Use at least two slides per genotype, per antibody combination.
14. For phalloidin staining, testes do not need to be permeabilized with DOC. Instead, wash in PBST, then block in PBST-FCS. Dry the required amount of labeled phal-loidin (stored at -20°C in methanol) in a Speed Vac just before use, and resuspend in the appropriate volume of PBST. Incubate for 2 h at room temperature, wash three times in PBST for 15 min each, then counterstain and mount. If costaining with an antibody, the phalloidin can be included in the secondary antibody incubation. Caution: Phalloidin is extremely hazardous! Wear protective clothing and be aware of the risks and safety procedures before using this compound.
15. To make Evostick wells, dry off the surface of the slide around the testes, taking care not to let the testes dry out. Paint a ring of Evostick or rubber cement around the area containing the testes and leave to dry for about 1 min (see Fig. 10). This forms a barrier, keeping a good depth of antibody solution above the tissue. Use 100 |L of diluted antibody per slide. Some antibodies penetrate the tissue much better under these conditions. Additionally, the testes remain stuck more firmly to the slide than if they are covered with a cover slip during antibody incubations. Evostick can be removed from the slide with tweezers.
16. If propidium iodide will be used to stain DNA, then RNAse A (0.5-1 mg/mL) must be added to one of the antibody incubations to reduce background fluorescence. I generally add it to whichever antibody incubation is being carried out at room temperature.
17. Do not fix in the tissue culture inserts used for other protocols, because glutaral-dehyde makes testes stick to the mesh. Testes can be transferred to the inserts during the washes after fixation, but because the whole protocol is short, little time is saved.
18. Four percent formaldehyde may be substituted for glutaraldehyde. It is less toxic, but the staining intensity will be lower, so it is only suitable for reporters that express well.
19. Prepare about 10 males per probe. If many probes are to be used, process the testes together until the transfer into tissue culture inserts step. Removal of the accessory glands and other bits of genital tract is not essential; they can provide a nice in-sample negative control. If any mutant testes being used are different enough to be easily told apart from wild-type by Nomarski optics, then an excel-
lent control is to mix wild type and mutant testes together after fixation. This allows a somewhat more accurate assessment of relative transcript levels, as the treatment of both genotypes will have been identical.
20. New batches of proteinase K should be checked. Overdigestion results in testes that are extra fragile and sticky, with a tendency to clump together.
21. To incubate at 65°C, the tissue culture dish can be left floating in a water bath.
22. Testes may be stored in HB at -20°C for up to 1 wk (in Eppendorf tubes).
23. The numbers of washes given here is a minimum. Increasing the number and total time of washing, especially in HB, can improve the signal-to-noise ratio.
24. To transfer stained testes, use a 200-|L pipet tip with the end cut off. Watch under the dissecting microscope to ensure complete transfer, especially of lightly stained tissue. Transfer of the testes at this stage is essential because methyl sali-cylate dissolves the tissue culture plastic.
25. Methyl salicylate clears the testes, but also dissolves some of the color product. Therefore, this incubation will make background staining disappear, but care must be taken to prevent the real staining from disappearing too. Canada balsam stabilizes the color, so remove the methyl salicylate and add GMM to the staining block when ready. The color should look quite intense under the dissecting microscope, in order to obtain good higher-magnification pictures. To mount the stained testes, transfer them in GMM onto a 22 x 22-mm2 cover slip with a cutoff pipet tip. Methyl salicylate also makes the testes very brittle so that any dissection (e.g., separation of pairs of testes) can be done by prodding or cutting with a tungsten needle.
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