Table of contents



1.1 Mounting animals for observation with Nomarski DIC optics by Monica Driscoll

1.2 Freeze-crack and staining protocol by Janet Duerr

1.3 FMRFamide staining protocol by Chris Li

1.4 Gonad-Intestine staining protocol by Barth Grant

1.5 Antibody staining of formaldehyde-fixed animals by Gary Ruvkun and Michael Finney

1.6 Protocol for staining early embryos by Bruce Bowerman

1.7 Formaldehyde fixation and cytokeletal staining by Raffi V. Aroian


2.1 FITC (fluorescein isothiocyanate) staining of amphid (ADF, ASH, ASI, ASJ, ASK, ADL) and phasmid (PHA and PHB) neurons :

A stock dye solution containing 20 mg/ml 5-fluorescein isothiocyanate in dimethylformamide can be stored at -20°C indefinitely. 50 ul of this stock solution is then mixed with 200 ul of M9 buffer and applied evenly to the surface of a 10 ml NGM plate preseeded with a lawn of E. coli. After 2 hr to allow the dye to diffuse into the agar, (final concentration 0.1 mg/ml), live animals are transferred to the plate. After staining for 2 hr to overnight, the animals are transferred to a plate devoid of the dye for at least 10 min to remove free FITC from the intestine before viewing under the microscope (Hedgecock et al, 1985)

2.2 DiI staining of amphid (ASI, ADL, ASK, AWB, ASH, ASJ), phasmid (PHA and PHB), IL1 and IL2 neurons and IL sheath and socket cells:

A stock dye solution containing 2mg/ml DiI (Molecular Probes, catalog # D-282) in dimethyl formamide can be stored at -20°C in a tube wrapped in foil. Transfer well-fed worms from a plate into an eppendorf tube with 1 ml M9, spin worms down at 2000-3000 rev/min, take out supernatant leaving (loose) worm pellet. Resuspend worms in1 ml M9 and add 5 microliter DiI stock sol (1:200 dilution) and incubate on a slow shaker for 3 hr-overnight (some dye may precipitate). Spin and wash worms with M9 twice before transferring them onto agar pads with sodium azide to visualize by fluorescence using the appropriate filters (DiI fluoresces red, therefore use the Texas red filters on the fluorescence scope). To additionally stain inner labial neurons and inner labial socket and sheath cells with DiI, wash the worm plate with 1ml H2O/50 mM Calcium acetate, wash once with 50 mM Calcium acetate and add 50 mM Calcium acetate into DiI staining solution and incubate for overnight. At the end of staining, wash with H2O before transferring worms onto agar pads with azide.
Amphids in D-V view
Amphids in lat view
Phasmids in D-V view

Inner labial neurons and inner labial socket and sheath cells

2.3 DiO staining of amphid (ASI, ADL, ASK, AWB, ASH, ASJ) and phasmid (PHA and PHB) neurons .

2.4 DAPI (4'-6-Diamidino-2-phenylindole) staining of nuclei: See example image

2.5 FITC staining of sensory neurons (Hedgecock et al., 1985)

2.6 Fluorescent Staining of Live Worms Using SYTO Nucleic Acid-Binding Dyes


3.2 Electron Microscopy Methods

3.2.1Conventional two-step fixation by David H. Hall

3.1.2 High pressure freeze fixation (HPF) by David H. Hall

3.1.3 Microwave aldehyde fixation followed by normal osmium fixation by David H. Hall

3.1.4 Osmium and aldehyde in one-step fixation by David H. Hall

3.1.5 Osmium-only fixation by David H. Hall

3.1.6 Metal mirror(slam-freeze) fixation (MMF) by David H. Hall

3.1.7 Flat Embedding of Worms (Slam Freezing) by Steve Fields (Rand Lab)

3.1.8 SEM preparation of worms by David Greenstein

3.1.9 Laser Hole Fixation by Carolyn Norris (Hedgecock Lab)

3.2 Experimental EM Methods

3.2.1 Freeze substitution in 1% Osmium by Robby Weimer -pdf file

3.2.2 Freeze substitution with tannic acid/osmium_long incubation by Robby Weimer-pdf file

3.2.3 Freeze substitution with tannic acid/osmium_short incubation by Robby Weimer-pdf file

3.2.4 High pressure freezing/freeze-substitution of C. elegans embryos and L1 worms by Richard Fetter

3.2.5 Freeze substitution in Potassium Permanganate by Robby Weimer-pdf file