C. elegans DAPI staining

(This protocol was adapted for students in my Developmental Biology course from methods worked out and detailed by Ross Francis in the Schedl lab at the University of Washington, MO)

Gonad dissections

1. Pick adults to an "unseeded" plate. Alternatively, wash worms off plate with phosphate buffered saline (PBS) and spin in a clinical centrifuge (3000 rpm) for 1 minute to pellet worms. Aspirate supernatant carefully.

2. Resuspend worms in 1-2 ml of PBS containing 0.2 mM Levamisole (this will paralyze worms) and transfer to the well of a depression slide.

3. As paralysis sets in, begin cutting off heads at level of pharynx: Place head between two 25 gauge syringe needles and decapitate by moving needles in a scissors motion. The release from the internal hydrostatic pressure of the worm should result in at least one gonad arm extruding completely.

4. Remove excess liquid with a drawn-out pasteur pipette.


Use one of the following methods:

A. Methanol: Add 3 ml of cold (-20C) methanol for 5 min. Using a pasteur pipette, transfer to a glass conical tube and add 2 ml PBTw (phosphate buffered saline containing 0.1 % tween-20). Spin 1 min in clinical centrifuge (3000 rpm). Carefully aspirate the supernatant and wash 2x in several ml of PBTw.

B. Formaldehyde: Fix in 3-4 ml of 3% formaldehyde in 0.1 M K2HPO4 (pH 7.2) for 2 hours. After fixation, transfer to a 10 ml glass conical tube, add a few mls of PBTw and spin 1 min in clinical centrifuge (3000 rpm). Remove supernatant, wash 1x in PBT, an then post-fix in 4 ml of ­20C methanol for 5 min. Fill tube with PBTw, spin, and wash 1x in 4 ml PBTw.

C. Methanol/Formaldehye: To make fixative, mix 10 mls of 16% formaldehye, 3.3 mls of 0.1 M K2HPO4 (pH 7.2), and 40 ml methanol. This solution is stored at ­20C and is used as described above for "methanol" (but increase fixation time to 10 min).

DAPI staining

1. CAUTION: Be very careful with DAPI. Always wear gloves. Throw all waste in special "DAPI waste" containers in hood. Remember, anything that binds to DNA can bind to YOUR DNA, and can thus potentially act as a carcinogen.

2. After last rinse of fixation protocol (above) pellet worms by either spining in clinical centrifuge or simply let worms settle by gravity to bottom of glass tube. Aspirate off supernatant and add 1 ml or less of DAPI solution (1:1000 dilution of stock in PBS).

3. Using a drawn capillary pipette, transfer settled worms onto a large 2% agarose pad** that covers most of a slide. After drawing off excess liquid with a capillary, an eyelash hair can be used to manipulate gonads for better viewing. Cover with a large (24 x 40 mm) coverslip, taking care not to move the coverslip once in place.

4. Slides can be stored at 4C for a week or more, particularly if sealed with nail polish around the periphery of the coverslip.



PBS: Dilute to 1x from 10x stock.

10x PBS: 80 g NaCl, 2 g KCl, 6.1 g anhydrous Na2HPO4, 2 g KH2PO4, H2O to 1 liter. Autoclave and store at room temperature.

PBTw: 1x PBS with 0.1% Tween 20

3% formaldehye/0.1 M K2HPO4(pH 7.2): Prepared from sealed ampoules of 16% EM grade formaldehyde. Freeze any excess.

DAPI (4',6-diamidino-2-phenylindole): Make stock solution by dissolving powder in 70% ethanol to a concentration of 100 µg/ml. To use, dilute stock in buffer 1:1000 (final concentration is 100 ng/ml).

Methanol: 100% stock kept at ­20C

**Agarose pads can be made by the following method: Boil 1g of agarose in 50 ml of H2O. Transfer solution to borosilicate tubes and place in heating block set at 70C. Using a pasteur pipette, add a drop of agarose to a glass slide and then immediately place another glass slide as if it were a coverslip at a perpendicular angle over the drop, spreading the agarose out thinly. When you want to add a sample to the slide, remove the top slide by carefully sliding it off.

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