1. Prepare humidity chamber: line large square plate with humid paper towels and place 4 5ml plastic pipets across chamber to support slides.

Place aluminum block in dry ice. Fill coplin jar (if you have only 1-10 slides) or Wheaton dish (if you have 10-20 slides) with acetone, and place in non-frost free -20oC freezer.

2. Polylysine coat slides as follows:

- Use all 3-well of 3-well brown slides from cel-line.

- Put 50uls. of polylysine solution onto each well. Be sure to cover the entire well. Let sit for 5-10minutes.

- Wipe excess liquid off the well with a KimWipe.

- Bake slides in a 60C oven for about 15 minutes.

3. Wash worms off the plate in EN Buffer (0.1M NaCl, 10mM EDTA) into a 15ml conical plastic tube.

4. Spin the worms in clinical centrifuge up to 1700 rpm. Do not use brake. Aspirate supernatant. The worm pellet may be very loose so be careful not to disturb it.

If you are staining more than one set of worms at a time, be sure to rinse out the aspirator between each set of worms by aspirating clean water through the system between each set .

5. Wash worms in EN buffer.

6. Spin and aspirate as in step 4.

7. Pipet 20-40 worms onto the well of the slide with a 50ul. capillary pipet or a yellow tip. Be sure to cover the entire surface of the well.

8. Allow the worms to settle and stick to the slide for about 1 minute.

9. Pipet off excess liquid, you should leave only about 5ul of liquid.

10. Put a coverslip (24mm X 50mm) on each well at right angle to the slide, so that the edge of the coverslip extends over the edge of the slide.

11. Put pressure on the coverslip with a pair of forceps to just burst the adults. Do this while watching through the microscope. Try not to completely squish the worms and break open the embryos.

12. Put the slide on the prefrozen aluminum block on dry ice and put a little pressure on the coverslip while the slide freezes. The slide can be left on dry ice at this point until all slides have been frozen.

13. Pop the coverslip off the slide and immediately put the slide in prechilled (-20C) 100% acetone. Put in -20oC freezer for 5'.

14. Put the slide through an acetone dilution series as follows:

- 75% acetone (RT) 1'

- 50% acetone (RT) 1'

- 25% acetone (RT) 1'

These washes can be done in plastic coplin jars which hold 40-60mls or in wheaton jars which hold 150 mls.

15. Put the slide in PM Buffer (50mM NaPi pH7.5, 1mM EDTA). The slide can stay in this buffer until you are ready to put on the stain mix.

16. Make up the stain mix (50uls for each well).

general formula / stocks / amount for 100ul final volume

0.2M NaPi pH7.5 0. / 5M NaPi pH7.5 / 40uls.

2mM MgCl2 / 1M MgCl2 / 0.2uls.

5mM KFe/5mM KFo / 50mM KFe/50mM KFo / 10uls.

75ug/ml Kanamycin / 3mg/ml Kanamycin / 2.5uls.

1ug/ml DAPI/ 0.1mg/ml DAPI / 1ul.

0.004% SDS / 1% SDS (freshly diluted) / 0.4uls.

0.024% X-gal / 3% X-gal (in DMF) / 0.8uls.

dH2O / to 100uls.

17. Wipe off the back of the slide and around the well carefully with a Kimwipe.

18. Put 50uls of stain mix on each well. Be sure to completely cover the well.

19. Let develop for 12 hours. Store the slide in a humidity chamber at RT or in the refrigerator.

20. When ready to view slides under the light microscope, remove excess staining solution and place long coverslip (22 x 70) on slide so as to cover all three wells.

For additional information on this protocol see Histochemical Techniques for Locating E. coli ß-galactosidase Activity in Transgenic Organisms, a paper by A. Fire.