C. elegans P-granule Antibody Staining
(Freeze crack protocol)

(This protocol was adapted from methods published by Susan Strome at Indiana U., Bloomington, IN and a protocol made available by the Seydoux Lab, Johns Hopkins Medical School, Baltimore, MD)


1. Prepare humidity chamber: Line large circular petri plate with damp paper towels.

2. Place an aluminum block on dry ice.

3. Fill 1 coplin jar with acetone and another jar with methanol. Place both at ­20C.

4. Polylysine coat slides as follows:

- use outer 2 wells of 3-well slides

- Put 20 µl of polylysine solution onto each well. Be sure to cover entire well.

- Let sit for 5-10 minutes

- Wipe excess liquid off well with a KimWipe

- Bake slides in a 60C over for approximately 15 minutes


Freeze-crack protocol, fixation, and staining

5. Wash worms off the plate in PBS into a 15 ml conical plastic or glass tube.

6. Spin the worms in clinical centrifuge up to 3000 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.

7. If necessary, wash worms in PBS buffer. Spin and aspirate as in step 6.

8. Pipet 20-40 worms onto the well of the slide with a 50µl capillary pipet or a drawn out glass pipet. Be sure to cover the entire surface of the well.

9. All the worms to settle an d stick to the slide for approximately 1 minute.

10. Put a coverslip (24 mm x 40 mm) 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 or the blunt end of a dissecting needle. Apply just enough pressure to 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 coverslips while the slide freezes (which should only take 1-5 seconds). 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% methanol. Put in ­20C freezer for 15 minutes.

14. Put the slides in prechilled (-20C) 100% acetone for 10 minutes.

15. Wash the slides in PBT Buffer (1X PBS with 0.1% Triton X-100 and 0.1% BSA)

3 x 10 minutes.

16. Incubate slides in PBT for 30 minutes.

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

18. Put 30 µl of OIC1D4 (1:3 dilution in PBT) on each well. Cover well with a parafilm coverslip.

19. Incubate at 4C overnight in sealed humidity chamber.

20. Wash the slides in PBT 3 x 10 minutes.

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

22. Put 30 µl of FITC- or CY2-GOAT ANTI-MOUSE (1:50 dilution in PBT) with DAPI (1:1000 dilution of 1 mg/ml) on each well.

23. Incubate at room temperature for 2 hours OR 4C overnight (or longer).

24. Wash the slides in PBT 3 x 10 minutes.

25. Wipe off the back of the slide and around the well carefully with a Kimwipe. Add one drop of "anti-fade" (DO NOT USE ANTIFADE WITH CY2-conjugated Abs) to each well and cover with a coverslip.

26. Observe stained worms with epifluorescent microscope.


This monoclonal antibody can be obtained as a cell supernatant from the Developmental Studies Hybridoma Bank at the University of Iowa (phone: (319) 335-3826, FAX: (319) 335-2077).

Secondary Antibodies

A Goat Anti-Mouse secondary antibody conjugated to FITC or Cy2 works well. These antibodies can be obtained from a variety of commercial sources, but I have had good success with Cy2-GAM from Jackson ImmunoResearch Laboratories.