9. Materials and methods are available as supplementary
materials on Science Online.
10. R. Nomura et al., Nature 473, 199–202 (2011).
11. D. Andrault et al., Nature 487, 354–357 (2012).
12. J. Zhang, C. Herzberg, J. Geophys. Res. 99, 17729 (1994).
13. R. G. Trønnes, D. J. Frost, Earth Planet. Sci. Lett. 197,
14. J. E. Dixon, L. Leist, C. Langmuir, J.-G. Schilling, Nature
420, 385–389 (2002).
15. B. Marty, Earth Planet. Sci. Lett. 313-314, 56–66 (2012).
16. A. K. McNamara, E. J. Garnero, S. Rost, Earth Planet.
Sci. Lett. 299, 1–9 (2010).
17. K. Sakamaki et al., Phys. Earth Planet. Inter. 174,
18. H. Terasaki et al., Phys. Earth Planet. Inter. 194-195,
19. K. Ohta et al., Earth Planet. Sci. Lett. 349-350, 109–115
20. G. M. Manthilake, N. de Koker, D. J. Frost, C. A. McCammon,
Proc. Natl. Acad. Sci. U.S.A. 108, 17901–17904
21. N. de Koker, G. Steinle-Neumann, V. Vlček, Proc. Natl.
Acad. Sci. U.S.A. 109, 4070–4073 (2012).
22. M. Pozzo, C. Davies, D. Gubbins, D. Alfè, Nature 485,
23. T. Lay, J. Hernlund, E. J. Garnero, M. S. Thorne, Science
314, 1272–1276 (2006).
24. J. W. Hernlund, S. Labrosse, Geophys. Res. Lett. 34,
25. H. Huang et al., Nature 479, 513–516 (2011).
26. H. Terasaki et al., Earth Planet. Sci. Lett. 304, 559–564
27. A. Shahar et al., Earth Planet. Sci. Lett. 288, 228–234
28. G. Morard et al., Phys. Chem. Miner. 38, 767–776 (2011).
29. R. A. Fischer et al., Earth Planet. Sci. Lett. 373, 54–64
30. J. Li, Y. Fei, in The Mantle and Core, R. W. Carlson,
Ed., vol. 2 of Treatise on Geochemistry, H. Holland,
K. K. Turekian, Eds. (Elsevier-Pergamon, Oxford, 2003),
31. T. Okuchi, Science 278, 1781–1784 (1997).
Acknowledgments: We thank M. Ishikawa, Y. Kudo, and
T. Tomomasa for their assistance in the experiments at
BL10XU and BL47XU, SPring-8 (proposals no. 2012B0087 and
2012B1706). Discussions with E. Takahashi and Y. Nakajima
were helpful. T. Kawamoto and F. Tomiyasu supported the
Fourier transform infrared and thermal conversion elemental
analyzer measurements, respectively. R.N. was supported by
a Japan Society for the Promotion of Science Fellowship for
Young Scientists. Data are available in the supplementary
Materials and Methods
Figs. S1 to S7
Tables S1 and S2
6 November 2013; accepted 30 December 2013
A Mechanosensory Pathway
to the Drosophila Circadian Clock
Alekos Simoni,1*† Werner Wolfgang,1 Matthew P. Topping,2 Ryan G. Kavlie,3
Ralf Stanewsky,1,4‡ Joerg T. Albert2,3,4‡
Circadian clocks attune the physiology of virtually all living organisms to the diurnal cycles of
their environments. In metazoan animals, multiple sensory input pathways have been linked to
clock synchronization with the environmental cycle (entrainment). Extrinsic entrainment cues
include light and temperature. We show that (12-hour:12-hour) cycles of vibration and silence (VS)
are sufficient to synchronize the daily locomotor activity of wild-type Drosophila melanogaster.
Behavioral synchronization to VS cycles required a functional clock and functional chordotonal
organs and was accompanied by phase-shifts of the daily oscillations of PERIOD protein
concentrations in brain clock neurons. The feedback from mechanosensory—and particularly,
proprioceptive—organs may help an animal to keep its circadian clock in sync with its
own, stimulus-induced activities.
The neurocellular network that adjusts an organism’s physiological needs to the di- urnal fluctuations of its environment is
summarily referred to as the “circadian clock”
(1). The tasks associated with the operation of
circadian clocks are computationally challenging. In metazoan animals, clock synchronization
requires integration of inputs from different sensory modalities, of which light and temperature
changes provide major cues.
Chordotonal organs (ChOs) have been linked
to temperature entrainment of the circadian clock
in adult flies (2). ChOs are internal mechano-
receptors mediating proprioception and the detec-
tion of air- and substrate-borne vibrations (3, 4).
If signaling from ChOs provides sensory input
for the entrainment of the fly’s circadian clock to
temperature cycles (2), we reasoned that exposure
to a rhythmic mechanical stimulus (Fig. 1, A and
B, stimulus details) that excites the fly’s ChOs (fig.
S1, response details) might phenocopy temperature entrainment (5) and be sufficient to synchronize the clock and clock-controlled locomotor
behavior. To test this, we first entrained adult
wild-type flies to 12-hour:12-hour light-dark (LD)
cycles (6). Flies were then transferred to constant
darkness (DD) and constant temperature (7). One
group remained in silence to serve as controls (Fig.
1C, top), whereas a second group was exposed to
12-hour:12-hour vibration:silence (VS) cycles (Fig.
1C, bottom). In the first, 4-day-long VS regime
(VS1), vibration onset was delayed by 6 hours from
light onset in the preceding LD cycles (L+6h). In
a second, 5-day-long vibration regime (VS2), vibration onset was then delayed by another 6 hours
(thus now L+12h). At the end of VS2, the flies
were released into the final free running (FR)
conditions—that is, darkness and silence—in which
they were kept for another 5 days (Fig. 1C).
During VS1, wild-type flies showed an initial
activity peak after vibration onset, which decreased
throughout the remaining vibration part (Fig. 1C
and fig. S2). During VS2, flies again showed increased activity immediately after vibration onset,
which declined rapidly (Fig. 1, C and D, and fig.
S2). In contrast to VS1, flies now also exhibited
increased activity several hours before vibration
onset, which is reminiscent of anticipatory behavior in LD cycles (Fig. 1D and fig. S2). To
quantify the behavioral activity occurring before
vibration onset (the anticipatory activity component), we determined the ratio of the activity in
the 4-hour time window before the V-phase and
the total activity in the S-phase [compare with (8)].
The resulting entrainment index (EI) revealed that
anticipatory activity was significantly increased
(Fig. 2B). To further probe whether the activity
patterns during the VS cycles resulted from a
clock-controlled synchronization of behavioral
activity, as indicated by the EI calculation, we
conducted a phase analysis of the activity peaks
in the final FR conditions between flies exposed
to VS cycles and controls (fig. S3) (7, 9). The FR
activity peaks were in phase with those of the
last VS cycle, demonstrating that the circadian
clock driving these rhythms had indeed been
stably synchronized (Figs. 1, C and D, and 2C).
In the control group, activity peaks free-ran
from the synchronized phase set during the initial LD cycle and hence occurred significantly
earlier (mean difference, 4.9 hours; P < 0.001,
Watson-Williams-Stevens test) (Figs. 1, C and
D, and 2C; fig. S3; and table S1) than those of
the experimental group.
Not all flies synchronized their activity to the
vibration cycles (fig. S4). We therefore assessed
each fly’s synchronization by inspecting individual actograms without any knowledge about
the experimental treatments of the particular fly
under investigation. This “observer-blind” analysis revealed that across eight independent experiments, ~53% of all flies (n = 312) synchronized
to vibration cycles (table S1). The reasons for this
incomplete synchronization are unclear. The vibration stimulus used across our experiments
1School of Biological and Chemical Science, Queen Mary, University of London, London E1 4NS, UK. 2Centre for Mathematics and Physics in the Life Sciences and Experimental Biology
(CoMPLEX), University College London, Gower Street, London
WC1E 6BT, UK. 3The Ear Institute, University College London,
332 Gray’s Inn Road, London WC1X 8EE, UK. 4Department of
Cell and Developmental Biology, University College London,
London WC1E 6DE, UK.
*These authors contributed equally to this work.
†Present address: Department of Life Science, Division of Cell and
Molecular Biology, Imperial College London, South Kensington,
London SW7 2AZ, UK.
‡Corresponding author. E-mail: email@example.com (R.S.);
firstname.lastname@example.org (J. T.A.)