1) If possible, grow larvae at 18°C in an uncrowded vial.  I only leave the adults in the vial for 3 or 4 days.

2) On a silicone plate, dissect larvae in Drosophila Ringers solutions and collect 1 to 3 pairs of glands in a single drop.  Place a small drop of 45% acetic acid on the silicone plate and 10 ul of 45% acetic acid on a siliconized coverslip.  With a single prong of the tweezer, scoop up each gland and briefly dip it into acetic acid on the silicone plate.  Then transfer the gland to the acetic acid on the coverslip.  You will probably have to push the gland off the tweezer with the prong of another tweezer.

3) Incubate the glands in acetic acid for 4 minutes.  Overlay the drop on the coverslip with a glass slide and press the slide against the coverslip.

4) Turn the slide over so the coverslip is up.  Tap the coverslip in the region where the glands are located.  The coverslip should slip around in order to break open cells.  Inspect the slides under the scope and continue to tap on the glands until they are adequately spread.

5) Place the slide with the coverslip down on a paper towel.  Cover the slide with a kimwipe and hold the slide steady with one hand.  With you thumb or fingers, press the slide over the region where the coverslip is in such a way so that you are pushing fluid from one side of the coverslip out the other side.   Examine the chromosomes in the scope - they should appear flattened.

6) Flip the slide over and hold the coverslip in place with a kimwipe.  Gently scratch over the surface of the coverslip with a metal probe.  Don't allow the coverslip to slide.  The chromosomes should be well flattened.

7) Slowly lower the slide into liquid nitrogen and allow it to freeze for 3 to 5 minutes.

8) Lift the slide out of the liquid nitrogen and blow briefly on the surface of the coverslip.  Quickly flip the coverslip off the slide with a razor blade and mark the region where the coverslip was located with a diamond scribe.  Quickly submerge the slide in 95% ethanol.  After 30 minutes, allow the slide to air dry and store in the refrigerator.  These slides supposedly last for months.

1) Place 100 ng of DNA in 16 ul of water.  Boil for 5 minutes, centrifuge briefly and place on ice.

2) Add 4 ul of vial 1.

3) Incubate overnight at 37°C.

4) Check for DNA synthesis by measuring 0.5 ul with the fluorometer.  I synthesized 700 ng on my first attempt.

5) Add 20 ug of carrier DNA (salmon sperm or calf thymus).

6) Add 1/10 volume 3M NaOAc. Precipitate with 2.5 volumes EtOH for 1 hr @ -70°C or overnight @ -20°C.

7) Spin DNA in microfuge 15 min. Remove EtOH, rinse the pellet with 75% EtOH and finally SpeedVac dry for 2 min.

8) Resuspend pellet in 150 ul hybridization buffer and store in the freezer.

Hybridization Buffer

0.6 M NaCl
50 mM Sodium phosphate buffer, pH 6.8
1X Denhardt's (0.02% BSA / 0.02% Ficoll / 0.02% polyvinylpyrrolidone) {I have a 10X stock in the freezer behind Cathy)
5 mM MgCl₂

The chromosomes used for hybridization must be non-refractile after ethanol dehydration. The chromosomes on the slide should look the same at this point as they did when the slide was made or it is not worth using. In general one slide is enough for a single probe if the chromosomes have good morphology. After the pretreatment the slides should be hybridized within 24 hours.  All of the pretreatments are done by placing the slides in coplin jars.

1) Place the slides in 2X SSC at 65°C for 30 minutes.  I do this in a Coplin Jar and prewarm the 2X SSC to 65°C in the microwave (check temp. with thermometer).

2) Wash for 2 minutes in 2X SSC at room temperature.

3) Acetylate the slides as follows: Prepare 50 ml of 0.1 M triethanolamine-HCl from a 1M stock that has been adjusted to pH 8.0. Add 62.5 ul of acetic anhydride and mix thoroughly. Pour into a Coplin jar and immerse slides for 10 minutes.

4) Wash the slides in 2X SSC for 5 minutes.

5) Denature the chromosomes by incubation in freshly prepared 0.1 N NaOH for 3 minutes.

6) Wash the slides in 2X SSC two times for 5 minutes each.

7) Ethanol dehydrate two times 5 minutes in 70% ethanol and 5 minutes in 95% ethanol. Air dry.

Hybridization and Washes:

1) Denature the desired amount of hybridization probe by boiling for 3 minutes and chill on ice.  Denature 12 ul per slide.

2) Apply 10 µl of hybridization solution on the slides and distribute the solution over the chromosomes with a 22 x 22 mm coverslip. Try to avoid trapping air bubbles.

3) To prevent evaporation of the probe seal the edges of the coverslip with rubber cement.

4) Incubate in a chamber containing 10 ml of 4X SSC at 60°C for 12-18 hours.  Use the hybridization oven in 451 N.Frear.  Carefully remove the rotating shaft by loosening the hex nuts.  Take care not to loose any parts.

5) Peel off the rubber cement and float the coverslip off in 2X SSC.
 

6) Wash the slides 3 times 15 minutes in 2X SSC at 65°C. Proceed directly with the signal detection. It is important not to let the slide dry out during the signal detection procedure.

Signal Detection (these manipulations are similar to what is done for immunofluorescence analysis of proteins):

1) Incubate slides for 3 minutes in TBS.

2) Block the slides with 15 ul of 10% FBS (in TBS) for 30 minutes at room temperature - cover with coverslip and place in humid chamber.

3) Float off coverslip in TBS and overlay glands with 15 ul of a 1/50 dilution of rhodamine tagged anti-digoxigenin antibody.  Dilute antibody in 10% FBS (in TBS).  Cover with coverslip and incubate in humid chamber at room temperature for one hour.

4) Float off coverslip and incubate slides for 10 minutes in TBS containing 1/100,000 Hoechst.

5) Incubate slides for 10 minutes in TBS lacking Hoechst.

6) Mount each slide with 100 mM Tris-Cl pH 8.5, 80% glycerol, 2% n-propylgallate and a coverslip.  Store in refrigerator until ready for microscopy.