Hybridization

1. 20X SSC: 3 MNaCl, 0.3 MNa-citrate. Adjust to pH 7.0 with NaOH and autoclave.

2. Humid chambers: Plastic boxes with close-fitting lids are lined with damp filter paper. Slides are propped off the bottom with broken plastic pipets.

3. 1 M Triethanolamine-HCl (TEA): Mix 132.7 mL of triethanolamine (Sigma-Aldrich, T-1377) in approx 800 mL dH2O. Bring pH to 8.0 with concentrated HCl and adjust the volume to 1 L. Store in the dark.

4. Acetic anhydride.

6. Hybridization mix: In a microfuge tube, mix 210 | L of formamide, 63 | L of 20X SSC, 77 |L dH2O, 40 mg dextran sulfate (MW 500K). Thoroughly mix and dissolve at 65°C for about 30 min. This can be stored at -20°C for 3 mo.

7. Probes: Probe DNA can be a synthesized oligonucleotide of a repetitive sequence, a polymerase chain reaction (PCR)-amplified repetitive sequence or a cloned stretch of unique or repetitive genomic sequence. The synthetic oligonucleotide should be 45-70 bp in length. PCR amplification has worked for the rDNA repeats and the 359-bp repeat from D. melanogaster using divergent primers. For a unique genomic region, the best probes are the cosmids and P1s from the Drosophila Genome Project (www.fruitfly.org). In determining the probe to use for specific genomic regions, one must consider size and uniqueness. I have found that as little as 15 kb can be detected using our microscope, but this can be unreliable. Fifty kilobases is good and 100 kb is very good. However, it is often difficult to find a larger clone that is unique for the region of interest. Each P1 or cosmid must be tested for uniqueness on mitotic diploid spreads and location accuracy on polytenes. For the P1s, I have found that even if one avoids the clones described as repetitive or chromocentral on polytenes, only about one in five are unique on mitotics. In general, it is probably better to go for something a bit smaller, like a cosmid, if there is a well-annotated one in your region of interest.

Probe DNA can be labeled by a number of methods, including (1) oligonucle-otide-primed incorporation using the Klenow fragment of DNA polymerase I, (2) incorporation of label during PCR to genomic DNA for repetitive sequences, and (3) terminal transferase. Oligonucleotides must be labeled by terminal trans-ferase. In order to use terminal transferase to label large genomic clones, the DNA must first cut down to 50- to 150-bp fragments using a 4-bp restriction enzyme mix (5).

There are two types of label: direct and indirect. Direct has the fluorochrome on the nucleotide. These are sold by Amersham (Fluoro Red or Green) and are available as rhodamine or fluorescein isothiocyanate (FITC) conjugates. These are very convenient, but although I have not seen a difference with sensitivity, they can have greater background artifacts. Also, one must be careful of light exposure in earlier steps of the hybridization procedure. The indirect method incorporates biotin-dUTP or digoxygenin (DIG)-dUTP and then detects it with fluorochrome-conjugated streptavidin or anti-DIG antibodies, respectively.

In considering the fluor choice, you must take into account the capabilities of your microscope system. For the basic BrdU detection, you will use the FITC and DAPI filters sets for BrdU and total DNA detection. This then leaves you with the rhodamine and Cy5 for probes, or if you do not have a scope with far red abilities, only rhodamine.

8. Formamide: Formamide must be of very high quality. Unfortunately, this is the major expense associated with this procedure. We use formamide from Fluka (47671) that is packed under nitrogen. It can be stored at 4°C until opened. After opening, formamide should be stored at -20°C. It can be aliquoted or defrosted at each use and then refrozen. When defrosting, minimize exposure to oxygen by not stirring or shaking it. If it does not freeze solid when it is returned to -20°C, it's no good. Formamide is a hazardous chemical and should be handled and disposed of properly.

9. Fluorochromes for detection of biotin- and digoxygenin-labeled probes: Streptavidin-Cy5 is from Jackson ImmunoResearch and made up to a concentration of 0.83 mg/mL in 50% glycerol. Anti-DIG Fab conjugated to FITC or rhodamine is from Roche and is made up at 0.2 mg/mL in 50% glycerol. All are aliquoted and stored at -20°C. Aliquots should not be refrozen, but stored at 4°C and will last 2 mo at this temperature. The aliquots stored at -20°C are good for at least 2 yr. Final dilution should be in PBS with 1% BSA (Sigma-Aldrich, A-7638).

10. Anti-BrdU-FITC antibodies: These can be obtained from a number of companies, including Roche and BioMeda. Anti-BrdU that is not conjugated to a fluor can also be purchased and a secondary antibody with your fluor of choice can be used. Follow the manufacturer's recommendations for preparation, storage and dilution. Final dilution should be in PBS with 1% BSA (Sigma-Aldrich, A-7638).

2.6. BrdU Detection Without FISH

3. Methods

3.1. Larvae Collection and Prelabeling Treatment

1. Place 50-80 females and at least 10 males on white Carolina instant food with about 1 mL of yeast paste of peanut butter consistency. Grow for 2-3 d, changing to new food (with yeast paste) every day.

2. In the morning of the third or fourth day, move the flies to a plastic bottle (without food) with approx 20 air holes poked in it. Tape a grape plate with large dollop of yeast paste to the opening of the bottle. Allow flies to "dump" eggs for 2-3 h. This will help get rid of eggs that are being held and make the subsequent collections more synchronous. Discard this grape plate and replace it with a new one having approx 50 |L of yeast paste. Collect embryos on grape plates at 2-h intervals. Maintain the collection plates at 25 °C in a larger Petri dish. Collections can be done similarly during succeeding days for at least a week. Allow flies to dump eggs each morning.

3. After 24 h, move larvae from grape plates to vials (30-50 larvae/vial) containing white instant food and a dollop of yeast paste.

4. Allow the larvae to develop at 25°C. At 95-100 h after egg deposition (AED), pick larvae out of the food. This can be done by scraping out the top layer of food into a watch glass containing Ringer's and picking out the larvae with a forceps. Larvae should be crawling in the food. Never take larvae crawling up the sides, as they have finished eating. (Of course, if they are properly synchronized, they should not be at that stage yet.) Rinse the larvae in Ringer's to remove adhered food. Place larvae on sucrose food for 2 h. Proceed with BrdU labeling.

3.2. BrdU Feeding

1. Pick larvae off the sucrose food, give them one final rinse with Ringer's, and place them on the culture plates containing BrdU-laced food. Let them eat for the desired time interval at 25°C (see Note 1). Place the BrdU culture plates in larger Petri dishes.

2. Remove the larvae from the food and rinse them in Ringer's or PBS. If you have labeled for less than about 3 h, make sure the larvae have blue food in their guts.

3.3. Pulse Feeding

1. Take larvae from the sucrose food as described in Subheading 3.2. Place them on dark blue food mixture for 15-20 min. The BrdU concentration should be from 1 to 2 mg/mL.

3. Remove the larvae to a dish of PBS and rinse. Select only those larvae with blue food visible in the crop, but not in the gut. Place them on crushed white instant food.

4. After 2 h from the beginning of BrdU feeding, transfer the larvae to a dish of PBS, and rinse. Select only those larvae with no blue food visible in their bodies. Place on crushed blue instant food (with no BPB or BrdU). If necessary, the larvae can then be examined for light blue food in the gut to ensure that they have continued eating.

5. Dissect after the desired "chase" time.

3.4. Dissection

1. If BrdU labeling is not being done, wandering third instar larvae can simply be collected from any food cultures. In either case, the larvae should be rinsed in Ringer's.

2. Use a multiwell depression plate. All solutions should fill the well about two-thirds. Put 0.7% NaCl in a well and either 1% (for interphase nuclei) or 0.5% (for mitotic spreads) sodium citrate solution in the well next to it. Dissect the larval CNS (brain lobes and ventral ganglion) in NaCl by pulling on the mouth-hooks. If you will be looking at interphase nuclei to examine nuclear structure, make sure that you remove the imaginal discs, because you will not be able to tell what tissue you are examining once you squash it. This precaution is not necessary if you will be looking at mitotic spreads.

3. Move the CNS into sodium citrate solution. For interphase cells, incubate the CNS for 5 min (in 1% sodium citrate); for mitotic spreads, incubate for 10 min (in 0.5% sodium citrate).

4. Clean a slide with 95% ethanol. Wipe with lint-free lens paper. Place 7 |L of 45% acetic acid on a siliconized 18 x 18-mm2 cover slip. Place fresh (less than 3 h old) MAW fixative in a well. Move brains to MAW for 30-60 s. If carryover of sodium citrate into the fixative is significant, decant the fixative using a micro-pipet and replace with additional fixative.

5. Move the CNS to acetic acid on a cover slip. It does not matter if the tissue falls apart at this point. If the squash is for mitotic figures, leave the CNS in the acetic acid for 1 min before squashing.

6. Pick up the cover slip with the slide so that the sample is in the middle of the slide. Move the cover slip slightly to spread the cells and wick out excess acetic acid with the edge of bibulous paper. Squash between folds of bibulous paper. Squash slightly for interphase and harder for mitotics.

7. Immerse the slide in liquid nitrogen until it stops sizzling. Remove the cover slip with a razor blade. Air-dry the slide.

8. Examine slides under phase contrast to determine if the squash was successful. Store in the dark until ready to hybridize.

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