Worm Breeder's Gazette 14(4): 21 (October 1, 1996)

These abstracts should not be cited in bibliographies. Material contained herein should be treated as personal communication and should be cited as such only with the consent of the author.

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

Katherine L. Hill, Steven W. L'Hernault

Emory University Dept. of Biology, 1510 Clifton Rd., Atlanta GA 30322

    We have been studying spermatogenesis-defective (Spe) worms that make
spermatozoa and are defective in some aspect of fertilization.  To
understand the exact nature of the defect, we needed to develop tools that
would allow us to study the behavior of these mutant spermatozoa.  One
tool we found to be especially useful is the SYTO vital nucleic acid dyes
from Molecular Probes.   Using these dyes, we have been able to label male
sperm in vivo and subsequently observe them within the reproductive tracts
of mated hermaphrodites after insemination.  This has revealed that
certain mutants contain spermatozoa that can migrate to the spermatheca
and maintain their position as oocytes pass through  to the uterus.

    This technique maybe of general use, for the following reasons:  The
dyes penetrate readily (with some variation between dyes) to label entire
worms, including internal structures.  There appear to be no serious
effects on viability.  Labelled males are able to mate efficiently, and
their sperm are able to fertilize oocytes and produce apparently
normal progeny.  Labelled hermaphrodites are fertile as well, and their
eggs hatch and produce healthy larvae.  The dyes come in a range of
excitation and emission maxima and therefore should be amenable to
a variety of applications.

    We have had particularly good results with SYTO 17, which
achieved the best penetration of the dyes we examined, and gives a
brilliant red signal in the rhodamine channel.  SYTO 25 behaves similarly,
with an intense orange-red signal.  Of the green dyes, (visualized in the
fluorescein channel) SYTO 11-16 and 20-24, we found #13 to be most
effective.  To date, there are fourteen dyes, #11-18 and #20-25 from which
to choose.  Listed below is a protocol for using these dyes that has
yielded consistent results.

1) Wash worms off plate with TBS using a cotton-plugged Pasteur pipette.
   Transfer to a 1.5 ml microcentrifuge tube.  (TBS:  10mMTris, 1mM EDTA,
   5mM NaCl, pH 7.4)
2) Spin worms to pellet.  Remove supernatant rapidly to prevent
   redistribution of worms.
3) Prepare dye solution:  3 ul of 5 mM  stock in 1497 ul TBS makes a 10
   uM solution (be sure to check the stock concentration; some are 1 mM).
   Add dye to worms and mix well.
4) Transfer worms and dye to a clean, empty 3.5cm plate.  Cover, seal with
   parafilm, wrap with aluminum foil and place in the appropriate
5) Incubate worms in dye for 3 hours.  This is a general guideline;
   different dyes will need more or less time.
6) After incubation, transfer worms to a fresh microcentrifuge tube, spin
   to pellet and remove dye solution.  This should be poured through
   activated charcoal to inactivate it for disposal.   Resuspend in a few
   drops TBS and spot onto plates with bacteria for recovery.  Once the
   plates are dry the worms can be picked.

Because of the rapid photobleaching of these dyes, we have found it
best to keep plates containing dyed worms shielded from the light as
much as possible.  These dyes can also be viewed by SIT camera
technology, which allows at least 15 minutes of continuous (dim)
illumination.  In addition, we recommend double gloves when handling
the dyes.