DOI: 10.1126/science.1222376
, 260 (2012);338Science
Roger C. Hardie and Kristian Franze
PhotoreceptorsDrosophilaPhotomechanical Responses in
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friction is present, total force on the EVL also has
a geometry-independent contribution, where retrograde
flow of actin and myosin-2 is resisted by
friction (termed “flow-friction motor”; Fig. 3A).
In the embryo, friction will arise when the flow
velocity in the ring is different from the velocity
of the adjacent material, such as the yolk cell
plasma membrane and the yolk cytoplasm. Consistent
with this, we observed differential flow velocities
between the actomyosin in the ring and
adjacent microtubules within the YSL (fig. S6).
This flow-friction motor pulls the EVL in a direction
opposite to the actomyosin flow, operates
at any stage, and can drive epiboly before passing
the equator (Fig. 3A and fig. S11).Notably, frictionresisted
flow provides additional tension in the
AV direction, consistent with the small degree of
tension anisotropy observed in the laser ablation
experiments(Fig.2B).Furthermore,experimentally
measured flow profiles within the EVL and the actomyosin
ring, as well as the relative tensions obtained
from laser ablation, are accurately predicted
by our theoretical description at all stages when
friction against the substrate is taken into account
(Fig. 3B). To conclude, we identified two distinct
modes of ring propulsion: a cable-constriction motor
due to circumferential contraction of the YSL
actomyosin network, and a flow-friction motor due
to contraction along the AVaxis of the network.
We next asked if the flow-friction motor is
sufficient to drive EVL epiboly. To this end, we
took advantage of the predicted geometry dependence
of the cable-constriction motor. Because
the cable-constriction motor cannot exert a net
force on the EVL when positioned right at the
equator, propulsion by this motor would be hindered
when the yolk cell is deformed from its original
spherical geometry into a cylindrical shape.
We thus deformed the yolk cell into a cylindrical
shape by aspirating pre–gastrula-stage embryos
(2.5 hpf ) into agarose tubes of a diameter smaller
than that of the embryo and analyzed resulting
changes in EVL movements. To verify that the
actomyosin ring is unperturbed in cylindrical embryos,
we analyzed the distribution and flow of
actin and myosin-2 within the YSL of cylindrical
embryos. We found that both the accumulation of
actin and myosin-2 in a ring-like structure adjacent
to the EVL/YSL border and their retrograde
flows from the vegetal pole toward the EVL/YSL
border were largely unaffected in cylindrical embryos
as compared to normal-shaped control embryos
(Fig. 4 and movie S10). This suggests that
the actomyosin ring remains intact in cylindrical
embryos. We observed that EVL movements were
largely unaffected in cylindrical embryos and proceeded
with velocities similar to those of spherical
control embryos (2.0 T 0.2 mm/min compared to
1.9 T 0.1 mm/min at 60 to 70% epiboly; compare
Figs. 4D and 2D). This shows that the cableconstriction
motor is not essential for EVL epiboly
movements and indicates that the flow-friction
motor is sufficient to drive this process.
Our findings have major implications for the
function of actomyosin rings in morphogenesis.
Whereas the prevalent model of actomyosin ring
function assumes circumferential contraction as
the main force-generating process, we present
evidence that friction-resisted actomyosin flows
can represent an equally important process mediating
ring function. This raises the possibility of
a more general role of cortical flows in morphogenetic
pattern formation processes (18).
References and Notes
1. S. E. Lepage, A. E. E. Bruce, Int. J. Dev. Biol. 54, 1213 (2010).
2. D. A. Kane et al., Development 123, 47 (1996).
3. M. Siddiqui, H. Sheikh, C. Tran, A. E. E. Bruce,
Dev. Dyn. 239, 715 (2010).
4. M. Köppen, B. G. Fernández, L. Carvalho, A. Jacinto,
C.-P. Heisenberg, Development 133, 2671 (2006).
5. F. A. Barr, U. Gruneberg, Cell 131, 847 (2007).
6. J. M. Sawyer et al., Dev. Biol. 341, 5 (2010).
7. A. Jacinto, S. Woolner, P. Martin, Dev. Cell 3, 9 (2002).
8. M. S. Hutson et al., Science 300, 145 (2003).
9. P. Martin, J. Lewis, Nature 360, 179 (1992).
10. J. C. Cheng, A. L. Miller, S. E. Webb, Dev. Dyn. 231,
313 (2004).
11. B. A. Holloway et al., PLoS Genet. 5, e1000413
(2009).
12. M. Mayer, M. Depken, J. S. Bois, F. Jülicher, S. W. Grill,
Nature 467, 617 (2010).
13. K. Kruse, J. F. Joanny, F. Jülicher, J. Prost, K. Sekimoto,
Eur. Phys. J. E 16, 5 (2005).
14. G. Salbreux, J. Prost, J. F. Joanny, Phys. Rev. Lett. 103,
058102 (2009).
15. D. Bray, J. G. White, Science 239, 883 (1988).
16. L. P. Cramer, Front. Biosci. 2, d260 (1997).
17. J. Howard, Mechanics of Motor Proteins and Cytoskeleton
(Sinauer Associates, Sunderland, MA, 2001).
18. N. W. Goehring et al., Science 334, 1137 (2011).
Acknowledgments: We are grateful to M. Sixt, T. Bollenbach,
and E. Martin-Blanco for advice and the service facilities of
the IST Austria and MPI-CBG for continuous help. M.B., G.S.,
S.W.G., and C.-P.H. synergistically and equally developed the
presented ideas and the experimental and theoretical
approaches. M.B. and P.C. performed the experiments; G.S.
developed the theory; and R.H., F.O., and J.R. contributed to
the experimental work. This work was supported by a grant
from the Fonds zur Förderung der wissenschaftlichen
Forschung (FWF) and the Deutsche Forschungsgemeinschaft
(DFG) (I930-B20) to C.-P.H., S.W.G., and G.S.
Supplementary Materials
www.sciencemag.org/cgi/content/full/338/6104/257/DC1
Supplementary Text
Figs. S1 to S16
Materials and Methods
References (19–36)
Movies S1 to S10
1 May 2012; accepted 10 September 2012
10.1126/science.1224143
Photomechanical Responses in
Drosophila Photoreceptors
Roger C. Hardie* and Kristian Franze
Phototransduction in Drosophila microvillar photoreceptor cells is mediated by a G protein–activated
phospholipase C (PLC). PLC hydrolyzes the minor membrane lipid phosphatidylinositol 4,5-bisphosphate
(PIP2), leading by an unknown mechanism to activation of the prototypical transient receptor potential
(TRP) and TRP-like (TRPL) channels. We found that light exposure evoked rapid PLC-mediated contractions
of the photoreceptor cells and modulated the activity of mechanosensitive channels introduced into
photoreceptor cells. Furthermore, photoreceptor light responses were facilitated by membrane stretch
and were inhibited by amphipaths, which alter lipid bilayer properties. These results indicate that, by
cleaving PIP2, PLC generates rapid physical changes in the lipid bilayer that lead to contractions of the
microvilli, and suggest that the resultant mechanical forces contribute to gating the light-sensitive channels.
I
n most invertebrate photoreceptor cells, the
visual pigment (rhodopsin) and other components
of the phototransduction cascade are
localized within tightly packed microvilli (tubular
membranous protrusions), together forming a
light-guiding rod-like stack (rhabdomere; Fig. 1).
After photoisomerization, rhodopsin activates a
heterotrimeric guanine nucleotide–binding protein
(Gq protein), releasing its guanosine triphosphate–
bound a subunit, which in turn activates phospholipase
C (PLC; Fig. 1C). How PLC activity
leads to gating of the light-sensitive transient receptor
potential channels (TRP and TRPL) in the
microvilli is unresolved (1–3). PLC hydrolyzes the
minor membrane phospholipid phosphatidylinositol
4,5-bisphosphate (PIP2), yielding soluble inositol
1,4,5-trisphosphate (InsP3), diacylglycerol (DAG,
which remains in the inner leaflet of the microvillar
lipid bilayer), and a proton. The light-sensitive
channels in Drosophila photoreceptors can be
activated by a combination of PIP2 depletion and
protons (4), but it remains unclear how PIP2 depletion
might contribute to channel gating. It has
been speculated that changes in membrane properties
play a role (4, 5), and members of the TRP
ion channel family have been repeatedly, although
controversially, implicated as mechanosensitive
channels (6, 7). This led us to ask whether cleavage
of the bulky, charged inositol head group of
PIP2 (Fig. 1C) from the inner leaflet might alter
the physical properties of the lipid bilayer in the
microvilli, resulting in mechanical forces that contribute
to channel gating.
Remarkably, Drosophila photoreceptors responded
to light flashes with small (<1 mm) but
rapid contractions that were directly visible in
Department of Physiology, Development and Neuroscience,
University of Cambridge, Cambridge CB2 3EG, UK.
*To whom correspondence should be addressed. E-mail:
rch14@cam.ac.uk
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Fig. 1. AFM measurements of photomechanical
responses. (A) An AFM cantilever contacts distal
tips of ommatidia in an excised Drosophila retina.
(B) An ommatidium, containing photoreceptors
(orange) and pigment cells (red). Elements of the
phototransduction cascade are contained within
microvillar rhabdomeres (two shown in longitudinal
section, seven in cross section), which are
rodlike stacks ~80 mm in length containing
~30,000 microvilli. Right: Electron micrograph
cross section of a rhabdomere (scale bar, 1 mm),
showing tubular microvilli, each ~50 nm in
diameter, with lumen in diffusional continuity with
the cell body. (C) Phototransduction cascade.
Rhodopsin (R) is photoisomerized to metarhodopsin
(M*), which catalyzes release of the Gq
protein a subunit to activate PLC. PLC hydrolyzes
PIP2 (red), leaving DAG (green) in the membrane.
Ca2+
influx via TRP channels inhibits PLC. (D) Lower
traces: AFM measurements of contractions (cantilever
z-position) in a wild-type retina in response to
5-ms flashes, with intensity increased from ~200
to 8000 effectively absorbed photons per photoreceptor.
Blue traces: Whole-cell current-clamped
voltage responses to the same stimuli recorded
from a dissociated photoreceptor cell. (E) Contractions
evoked by 5-ms flashes covering the full
intensity range (~200 to 106
photons) in a wildtype
retina. (F) Same on faster time base. (G)
Response versus intensity (R/I) functions of contractions
(nm) from wild-type retinae (mean T SEM,
N = 13), and peak voltage (mV) recorded from
dissociated photoreceptors (blue; means T SEM,
N = 6). (H) Responses to flashes (~5 × 104
photons)
in trpl mutant before and after (red) channel block
by 50 mM La3+
and 10 mM ruthenium red (RR),
which prevents inhibition of PLC by Ca2+
influx.
Blue trace: Lack of response in norpAP24
(PLC
mutant) despite using flashes of higher intensity
(~2 × 105
photons; N = 3). (I) R/I function of
contractions from trpl retinae before and after (red)
channel block by La3+
and RR (means T SEM, N = 5). For intensity calibration, see fig. S2.
Fig. 2. Light-induced modulation of gramicidin channels.
(A) Whole-cell recordings from trpl;trp mutant photoreceptor
cell lacking all native light-sensitive channels.
Perfusion with gramicidin induced a constitutive inward
current. (B) Flashes of increasing intensity (5 × 103
to 4 ×
105
effective photons), each 1 s (denoted by bar at upper
left), up-regulated the current. (C) Averaged responses
(TSEM) to 100-ms flashes containing 1.3 × 104
(middle
trace, N = 4) and 3.7 × 104
(lower trace, N = 10) effective
photons (data pooled from trpl;trp and trpl mutants recorded
in the presence of La3+
and RR). The same flashes delivered
before gramicidin application (controls) induced residual,
noise-free transient currents of uncertain origin. (D) R/I
function (after subtracting control responses measured
before gramicidin perfusion) expressed as a fraction of the
steady-state gramicidin current I/ISS (means T SEM, N = 4).
Dotted curve: R/I function of contractions measured by AFM
in trpl mutant in the presence of La3+
and RR, replotted from
Fig. 1I.
www.sciencemag.org SCIENCE VOL 338 12 OCTOBER 2012 261
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dissociated cells via bright-field microscopy (see
movie S1). To obtain improved temporal and
spatial resolution, we recorded these photomechanical
responses with an atomic force microscope
(AFM) (8), positioning the AFM cantilever
on the distal tips of photoreceptors in a whole
excised retina glued to a coverslip (Fig. 1A). Contact
force (~100 pN) was maintained constant, so
that changes in sample height resulted in imme-
diate,matchingchangesinthecantilever’sz-position.
Contractions were elicited indefinitely by repeated
brief flashes of modest intensity, with kinetics
similar to those of electrical responses recorded
from dissociated photoreceptors (Fig. 1D and fig.
S1). The latencies of contractions induced by the
brightest stimuli (4.9 T 0.9 ms, mean T SEM, N =
11; Fig. 1F) were significantly shorter than the
latencies of voltage responses to the same stimuli
(6.6 T 0.6 ms, N = 6; P = 0.002, unpaired twotailed
t test). Only a few hundred effectively absorbed
photons per photoreceptor were required
to elicit detectable contractions, which saturated
with flashes containing ~106
photons, corresponding
to ~30 effectively absorbed photons per microvillus
(Fig. 1, E and G). This intensity dependence
overlapped with that of the electrical response
(Fig. 1G and supplementary text). Like the electrical
responses, the contractions were eliminated
in mutants lacking PLC (norpAP24
), which shows
that they too required PLC activity (Fig. 1H).
Because PLC activity is normally terminated
by Ca2+
influx through the light-sensitive channels,
net PIP2 hydrolysis is enhanced when the
light-sensitive current is blocked (9, 10). We therefore
measured contractions in trpl mutants expressing
only TRP light-sensitive channels before
and after blocking TRP channel activity with La3+
and ruthenium red (RR). Indeed, after blocking
the light-sensitive channels the contractions were
enhanced, more sensitive to light, and saturated at
lower intensities, corresponding to only ~1 to 5
effectively absorbed photons per microvillus
(Fig. 1, H and I). In the absence of Ca2+
influx,
such intensities deplete virtually all microvillar
PIP2, resulting in temporary loss of sensitivity to
light (10). After such saturating flashes, the
photomechanical response was also temporarily
refractory, recovering sensitivity with a time course
(t1/2 ~ 40 s) similar to that of PIP2 resynthesis
(10). By contrast, without channel blockers, sensitivity
recovered within ~10 s (fig. S3).
Blockade of all light-sensitive current in these
experiments also shows that the contractions cannot
result from any downstream effects of Ca2+
influx or osmotic changes caused by ion fluxes associated
with the light response. Therefore, these
results indicate that the contractions result from
hydrolysis of PIP2. Although we do not exclude
other downstream effects of PLC, the speed of
the contractions supports a simple and direct mechanism.
Cleavage of the bulky head groups from
PIP2 molecules, which represent 1 to 2% of
lipids in the plasma membrane, leaves DAG in
the membrane, which occupies a substantially
smaller area than PIP2. This should increase membrane
tension, leading to shrinkage of the microvillar
diameter, as reported for the action of PLC
on the diameter of artificial liposomes (11). Integrated
over the stack of ~30,000 microvilli,
such a mechanism seems capable of accounting
for the observed macroscopic contractions, which
represent at most ~0.5% of the rhabdomere length
(see supplementary text). Within each microvillus,
we suggest that the alteration to the mechanical
properties of the lipid bilayer may
contribute to channel gating.
To test whether phototransduction generates
sufficient mechanical forces to gate mechanosensitive
channels (MSCs), we made whole-cell
patch-clamp recordings from dissociated photoreceptors
lacking all light-sensitive channels
(trpl;trp double mutants, or trpl mutants exposed
to La3+
and RR). We then perfused the photoreceptors
with gramicidin, a monovalent (Ca2+
impermeable)
cation channel and one of the
best-characterized MSCs, which is known to be
regulated by changes in bilayer physical properties
(12, 13). Incorporation of gramicidin channels
into the membrane generated a constitutive
inward current that stabilized after a few minutes
(Fig. 2A). Despite having replaced the native
light-sensitive channels with MSCs, the photoreceptors
still responded to light, with a rapid
increase in the gramicidin-mediated current (Fig.
2, B and C). Like the photomechanical responses
recorded after blocking Ca2+
influx through the
light-sensitive channels (Fig. 1H), these gramicidinmediated
responses inactivated slowly (Fig. 2C),
were temporarily refractory to further stimulation
and had a similar intensity dependence (Fig. 2D).
To test whether the light-sensitive channels
were mechanically sensitive,we manipulatedmembrane
tension osmotically. Channels were not
directly activated by perfusing cells with hyperor
hypo-osmotic solutions; however, we reasoned
Fig. 3. Modulation of light-sensitive channels by osmotic pressure. (A to D) Whole-cell voltage-clamped
responses to 1-ms flashes (~50 effective photons) in control bath (300 mOsm) were reversibly increased
by perfusion with 200 mOsm solution and suppressed by 400 mOsm in wild-type (A), trpl (B), and trp (C)
mutants as well as in wild-type photoreceptors recorded in Ca2+
-free solutions (D). (E) Response amplitudes
(I/I300) after hyperosmotic (400 mOsm) and hypo-osmotic (200 mOsm) challenges normalized to
control responses in 300 mOsm bath. Data are means T SEM; N = 4 to 8 cells. All conditions plotted were
significantly increased (200 mOsm) or decreased (400 mOsm) relative to control responses from the same
cells [P < 0.005; analysis of variance (ANOVA) followed by posttest for trend]. (F) Spontaneous TRP
channel activity (from trpl mutant) after several minutes of recording, using pipettes lacking nucleotide
additives. Perfusion with 400 or 200 mOsm (bar) reversibly suppressed and facilitated this “rundown
current” (RDC). (G) Channel noise resolved on a faster time base, plus trace recorded in dark before onset
of RDC. (H) Left: Amplitude of steady-state RDC normalized to value at 300 mOsm. Right: Effective singlechannel
conductance (g) estimated by variance/mean ratio (means T SEM, N = 7). Although macroscopic
RDC was substantially modulated (P < 0.001; ANOVA, posttest for trend), single-channel conductance was
not significantly affected by osmotic manipulation (P > 0.2).
12 OCTOBER 2012 VOL 338 SCIENCE www.sciencemag.org262
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that it would be impossible to mimic the exact
physical effects of PIP2 hydrolysis, which would
include a specific combination of changes in
membrane tension, curvature, thickness, lateral
pressure profile, charge, and pH. We therefore
tested whether osmotic manipulation could enhance
or suppress light-sensitive channel activity.
In wild-type photoreceptors, light-induced currents
were rapidly and reversibly facilitated by ~50%
after perfusion with hypo-osmotic solutions (200
mOsm), which, like PIP2 depletion, would be expected
to alleviate crowding between phospholipids,
increase tension, and reduce membrane
thickness. Conversely, responses in hyperosmotic
(400 mOsm) solutions were about half those
in control solutions (Fig. 3). Analysis of singlephoton
responses (quantum bumps) indicated
that modulation resulted from changes in both
quantum efficiency (fraction of rhodopsin photoisomerizations
generating a quantum bump) and
bump amplitude (fig. S4 and supplementary text).
Recordings from trp and trpl mutants showed
that both TRP and TRPL channels were modulated,
although facilitation of currents mediated
by TRPL channels (in trp mutants) was more
pronounced (Fig. 3). Modulation of the light response
by osmotic manipulation was at least as
pronounced in Ca2+
-free bath (Fig. 3D), indicating
that facilitation by membrane stretch did not
result from leakage of Ca2+
into the cell from the
extracellular space.
To test whether modulation might have been
mediated by effects on upstream components of
the cascade such as PLC (14), we measured the
activity of spontaneously active TRP channels in
recordings made with pipettes lacking adenosine
triphosphate. Under these conditions, PIP2 becomes
depleted (thereby removing PLC’s substrate),
sensitivity to light is lost, and the TRP
channels enter a constitutively active (“rundown”)
state uncoupled from the phototransduction cascade
(15, 16). Nonetheless, the channels were
still similarly modulated by osmotic manipulation,
whereas single-channel conductance, estimated
by noise analysis, was unaffected (Fig. 3,
F to H). These results indicate that osmotic pressure
directly modulated the open probability of
both TRP and TRPL channels.
MSCs such as gramicidin are sensitive to
amphiphilic compounds, which insert into the
lipid bilayer. Because they are attracted to anionic
phospholipids, cationic amphipaths insert preferentially
into the inner leaflet, where they increase
crowding, promote negative (concave) curvature,
and decrease membrane stiffness (17, 18). We
found that four structurally unrelated cationic
amphipaths were all effective, reversible inhibitors
of the light-induced current. Neither light-induced
PLC activity (measured using a genetically targeted
PIP2-sensitive biosensor to monitor PIP2
hydrolysis) nor single-channel conductance were
substantially affected (fig. S5). The 50% inhibitory
concentrations (IC50 values) were much
higher than those of their traditional drug targets
and ranged over approximately three orders of magnitude.
However, after correcting for pKa and
partitioning, the effective concentration of the compounds
in the membrane was similar (~5 mM) in
each case (Fig. 4). Thus, their mode of action is
likely related to their physicochemical properties
rather than conventional drug-receptor interactions.
Because cationic amphipaths are also lipophilic
weak bases, and because we propose
that protons are also critical for activating the
light-sensitive channels (4), an alternative but not
mutually exclusive possible mechanism of action
is as lipophilic pH buffers of the membrane
environment. We also note that polyunsaturated
fatty acids (PUFAs) such as arachidonic and linolenic
acid—which are effective activators of both
TRP and TRPL (5, 19)—are not only anionic
amphipaths (predicted to have opposite effects
to cationic amphipaths) but also, as weak acids,
natural protonophores; such a dual action could
account for their agonist effect.
The mechanism of activation of the lightsensitive
channels in invertebrate microvillar
photoreceptors has long remained an enigma
(2, 3, 20). Neither InsP3 nor DAG—the two obvious
products of PIP2 hydrolysis—are reliable
agonists for the light-sensitive channels. Although
PUFAs are effective agonists and might be generated
from DAG, a DAG lipase with the appropriate
specificity has not been found in the photoreceptors
(21). By contrast, two neglected consequences
of PLC activity—the depletion of its substrate
(PIP2) together with protons released by PIP2
hydrolysis—were recently shown to potently activate
the light-sensitive channels in a combinatorial
manner (4). Our results support the hypothesis that
the effect of PIP2 depletion is mediated mechanically
by changes to the physical properties of the
lipid bilayer, thereby introducing the concept of
mechanical force as an intermediate or “second
messenger” in metabotropic signal transduction.
References and Notes
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Acknowledgments: We thank D. G. Stavenga, S. B. Laughlin,
and M. Postma for comments on the manuscript; O. Andersen
for advice on the use of gramicidin; and J. Grosche (Effigos AG)
for artwork for Fig. 1A. Supported by UK Biotechnology and
Biological Sciences Research Council grant BB/G0068651
(R.C.H.) and a UK Medical Research Council Career Development
Award (K.F.). Author contributions: project initiation, R.C.H.;
whole-cell electrophysiology experiments, R.C.H.; AFM
measurements, K.F., R.C.H.; paper written by R.C.H. with
contribution from K.F.
Supplementary Materials
www.sciencemag.org/cgi/content/full/338/6104/260/DC1
Materials and Methods
Supplementary Text
Figs. S1 to S5
Movie S1
References (22–31)
26 March 2012; accepted 7 August 2012
10.1126/science.1222376
Fig. 4. Cationic amphipaths
inhibit the light response.
(A) Response to 1-ms flashes
in trp mutants in the pres-
enceofprocaine(PROC),imipramine
(IMP), trifluoperazine
(TFP), and chlorpromazine
(CPZ). Control responses before
(turquoise) and after
washout (blue) are superimposed.
(B) Dose response
functions (means T SEM:
PROC, N = 4; IMP, N = 3;
TFP,N=3;CPZ, N = 6) fitted
by inverse Hill curves (slope
constrained at n = 2), based
on raw values (IC50: PROC,
3.1 mM; IMP, 27 mM; TFP,
4.9 mM; CPZ, 4.5 mM). (C)
Same as in (B) after correction
for pKa and octanol partition
coefficients, reflecting
predicted concentration in the lipid membrane (IC50: PROC, 2.7 mM; IMP, 7 mM; TFP, 3.4 mM; CPZ, 2.3 mM).
www.sciencemag.org SCIENCE VOL 338 12 OCTOBER 2012 263
REPORTS
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