Dros. Inf. Serv. 96 (2013) Technique Notes 199
were not different among any of the four interactions, it is not known whether the larger tube will
alter other activity parameters, such as average bout length, counts per bout, bouts per day, or
Light:Dark activity ratio.
References: Ahmad, S.T., S.B. Steinmetz, H.M. Bussey, B. Possidente, and J.A. Seggio
2013, Behav. Brain Res. 241: 50-55; Chiu, J.C., K.H. Low, D.H. Pike, E. Yildirim, and I. Edery
2010, J. Vis. Exp. (43); Helfrich-Forster, C., 2004, J. Comp. Physiol. A Neuroethol. Sens. Neural
Behav. Physiol. 190(8): 601-613; Klarsfeld, A., J.C. Leloup, and F. Rouyer 2003, Behav. Processes
64(2): 161-175; Konopka, R.J., and S. Benzer 1971, Proc. Natl. Acad. Sci. USA 68(9): 2112-2116;
Lone, S.R., and V.K. Sharma 2012, J. Biol. Rhythms 27(2): 107-116; Oh, Y., D. Jang, J.Y. Sonn,
and J. Choe 2013, PLoS ONE 8(7): e68269; Pfeiffenberger, C., B.C. Lear, K.P. Keegan, and R.
Allada 2010, Cold Spring Harb. Protoc. 2010(11): pdb prot5518; Rosato, E., and C.P. Kyriacou
2006, Nat. Protoc. 1(2): 559-568; Seggio, J.A., 2011, Dros. Inf. Serv. 94: 170-173; Seggio, J.A., B.
Possidente, and S.T. Ahmad 2012, Chronobiol. Int. 29(1): 75-81; Zordan, M.A., C. Benna, and G.
Mazzotta 2007, Methods Mol. Biol. 362: 67-81.
An efficient, practical, and reliable Drosophila trap.
Freda, P.J.1, and J.M. Braverman1*. 1Department of Biology, Saint Joseph's
University, Philadelphia, PA, USA; *corresponding author (E-mail: email@example.com).
A good Drosophila trap should be made of materials that are inexpensive and readily
available. Also, the materials should be sturdy enough to be used outdoors. Additionally, a trap
should be simple enough that anyone can assemble it quickly. The trap of Medeiros and Klaczko
(1999) is well designed, but improvements and simplifications are possible. Using their work as a
foundation, an efficient, practical, and reliable trap for live Drosophila specimen collection was
Table 1. Average activity (number of beam crossings per 10 min bin ± SEM) in LD and DD, as well
as free-running period (hours ±SEM) for the two wild-type flies (CS and Or-R) and the period
mutants (perS and perL).
Activity in LD
Activity in DD
5.00 ± .31
5.13 ± .31
24.34 ± .03
4.28 ± .30
5.14 ± .35
24.33 ± .02
6.35 ± .37
6.56 ± .31
24.05 ± .04
5.92 ± .44
6.47 ± .48
24.04 ± .04
6.87 ± .60
7.25 ± .57
19.26 ± .12
6.79 ± .58
7.65 ± .81
19.29 ± .10
4.82 ± .37
4.77 ± .42
28.37 ± .11
5.29 ± .52
4.00 ± .39
28.44 ± .18
200 Technique Notes Dros. Inf. Serv. 96 (2013)
Figure 1. Bottle cap
The first step is to modify a bottle cap for attachment of a plastic
culture vial (1¼’’ diameter 4’’ high; Carolina Biological Supply item
#173120). Using a rotary tool or sharp blade, make a hole approximately
½-inch in diameter in the middle of the bottle cap so flies can travel
through it (Figure 1a). Make sure not to damage the threading on the inside of the cap so it can still
be tightened onto the bottle. Next, add masking tape to the outside of the bottle cap (Figure 1b).
Circle the side of the cap with enough tape so that a culture vial fits snuggly. The tape may need to
be cut horizontally so that it does not interfere with the culture vial. Modify multiple caps as a supply
for many traps. Alternatively, this trap can be used without a bottle cap by taping the culture vial to
the bottle itself. However, the cap makes it much easier to remove and replace the culture vial
Figure 2. Procedures for making the trap.
The containment portion of the trap is
made with one transparent 2-L soda bottle.
Alternatively, for a smaller trap, a 20-oz
(591-mL) bottle can be used. The first step is
to make a small hole near the middle of the
bottle (Figure 2a). From this hole, cut a
curved slit into the bottle (Figure 2b). The
best tool to use for this is a box cutter. Bait
will be introduced through this slit. After the
trap is hung, this slit can be taped over to
ensure larger animals, such as wasps, do not
enter the trap. However, this usually is not
an issue. Second, using a narrow, sharp tool,
puncture small holes (~ 4 – 5 mm in
diameter) in the trap at random locations above the curved slit (Figure 2c). Make these holes large
enough for fly entry but small enough to limit the entry of larger insects. Make 15 to 30 of these
holes for a 2-L trap. Next, add the bait via the curved slit (Figure 2d). Finally, place the culture vial
onto the modified cap at the top of the trap (Figure 2e). The trap can be hung from vegetation or
hooks using a bent wire coat hanger (Figure 2f).
To collect flies from the trap, have several culture vials and plugs available. Flick the trap by
gently tapping it a few times. Flies will migrate upwards into the culture vial. Quickly remove the
culture vial and immediately plug it using a foam plug (Carolina Biological Supply item # 173122)
while holding the vial upside down. Replace the culture vial with an empty one. Repeat the
Dros. Inf. Serv. 96 (2013) Technique Notes 201
collection process until the trap is empty. A filtered aspirator (for example, Bioquip item #1135A)
can be used through the slit to selectively remove specimens.
Using a sleeve expedites the collection process. Hollow cardboard mailing tubes (Uline
model # S-10723) with plastic end caps (Uline model # S-7020) can be used as a sleeve (Figure 3a).
Using a rotary tool, drill, or sharp blade, make a hole into the plastic cap of the tube so that the
culture vial can tightly fit through (Figure 3b). Next, cut the tube to roughly the height of the bottle
without the shell vial attached (Figure 3c). When collecting, slide the tube over the trap (Figure 3d).
Specimens migrate upwards toward light into the culture vial (Figure 3e). Natural light or a lamp
may be used.
Figure 3. Collecting specimens using a sleeve.
A trap of this design is efficient because it
is capable of trapping thousands of specimens 2
to 3 days after deployment, and it can be emptied
quickly and easily. This trap has captured flies
from many different species, including D.
melanogaster, D. simulans, D. busckii, D.
robusta, D. affinis, D. tripunctata, D. immigrans,
D. suzukii (Freda and Braverman, 2013), and
References: Freda, P.F., and J.M. Braverman 2013, Entomol. News 123(1): 71-75;
Medeiros, H.F., and L.B. Klaczko 1999, Dros. Inf. Serv. 82: 100-102.
The impact of pheromones on sexual behavior in D. melanogaster:
Recommendations for laboratory protocols.
Beck, Aaron P.1,3, Erica F. Hamlin2,3, Erika L. Hume1,3, and Robert M. Hallock1,4.
Skidmore College; 1Neuroscience program, Skidmore College, 815 N Broadway,
Saratoga Springs, NY 12866 2Psychology Department, Skidmore College, 815 N Broadway,
Saratoga Springs, NY 12866; 3These authors contributed equally to this work; 4Corresponding
Pheromones are conspecific chemical signals used throughout the animal kingdom that elicit
behavioral responses in other organisms and are essential for intraspecies communication. D.