Download Live cell imaging reveals actin-cytoskeleton

ß 2014. Published by The Company of Biologists Ltd | Journal of Cell Science (2014) 127, 1357 doi:10.1242/jcs.151852
CORRECTION
Live cell imaging reveals actin-cytoskeleton-induced
self-association of the actin-bundling protein WLIM1
Céline Hoffmann, Daniéle Moes, Monika Dieterle, Katrin Neumann, Flora Moreau, Angela Tavares Furtado,
Dominique Dumas, André Steinmetz and Clément Thomas
There was an error published in J. Cell Sci. 127, 583–598.
The last sentence of the Introduction section contained a typographical error. This sentence should read as follows: Together, our data
support a multistep process in which F-actin-induced NtWLIM1 self-association operates as a driving force for the zippering of
NtWLIM1-decorated AFs.
We apologise to the authors and readers for any confusion that this error might have caused.
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ß 2014. Published by The Company of Biologists Ltd | Journal of Cell Science (2014) 127, 583–598 doi:10.1242/jcs.134536
RESEARCH ARTICLE
Live cell imaging reveals actin-cytoskeleton-induced
self-association of the actin-bundling protein WLIM1
ABSTRACT
Crosslinking of actin filaments into bundles is essential for the
assembly and stabilization of specific cytoskeletal structures.
However, relatively little is known about the molecular
mechanisms underlying actin bundle formation. The two LIMdomain-containing proteins define a novel and evolutionarily
conserved family of actin-bundling proteins whose actin-binding
and -crosslinking activities primarily rely on their LIM domains.
Using TIRF microscopy, we describe real-time formation of actin
bundles induced by tobacco NtWLIM1 in vitro. We show that
NtWLIM1 binds to single filaments and subsequently promotes their
interaction and zippering into tight bundles of mixed polarity.
NtWLIM1-induced bundles grew by both elongation of internal
filaments and addition of preformed fragments at their extremities.
Importantly, these data are highly consistent with the modes of
bundle formation and growth observed in transgenic Arabidopsis
plants expressing a GFP-fused Arabidopsis AtWLIM1 protein.
Using two complementary live cell imaging approaches, a close
relationship between NtWLIM1 subcellular localization and selfassociation was established. Indeed, both BiFC and FLIM-FRET
data revealed that, although unstable NtWLIM1 complexes can
sporadically form in the cytosol, stable complexes concentrate
along the actin cytoskeleton. Remarkably, disruption of the actin
cytoskeleton significantly impaired self-association of NtWLIM1. In
addition, biochemical analyses support the idea that F-actin
facilitates the switch of purified recombinant NtWLIM1 from a
monomeric to a di- or oligomeric state. On the basis of our data, we
propose a model in which actin binding promotes the formation and
stabilization of NtWLIM1 complexes, which in turn might drive the
crosslinking of actin filaments.
KEY WORDS: Actin dynamics, Arabidopsis, BiFC, FLIM-FRET, LIM
proteins, TIRF microscopy, Tobacco BY-2 cells
INTRODUCTION
As a vital structure of eukaryotic cells, the actin cytoskeleton is a
key player in numerous processes, including cell division and
elongation, vesicle and organelle trafficking, adhesion and
1
Centre de Recherche Public-Santé, 84 Val Fleuri, L-1526 Luxembourg,
Luxembourg. 2Plateforme d’Imagerie Cellulaire et Tissulaire PTIBC-IBISA, CNRSUMR 7563 et FR3209. Université de Lorraine, F-54505 Vandoeuvre-lès-Nancy,
France.
{
These authors contributed equally to this work.
*Author for correspondence ([email protected])
Received 3 May 2013; Accepted 30 October 2013
motility, and establishing polarity. In line with this wide range
of functions, the actin cytoskeleton exhibits a high degree of
structural plasticity allowing a highly dynamic rearrangement of
its basic elements, namely the actin filaments. The creation and
turnover of actin filaments, as well as their assembly into higherorder structures, are tightly regulated at spatial and temporal
levels by a plethora of actin-binding proteins, which control
nucleation, polymerization, capping, severing and crosslinking
(dos Remedios et al., 2003; Higaki et al., 2007; Staiger and
Blanchoin, 2006; Thomas et al., 2009; Winder and Ayscough,
2005). Recent live cell studies combining reliable fluorescent
actin markers with novel high-resolution imaging techniques
such as spinning-disc confocal microscopy and variable-angle
epifluorescence microscopy (VAEM) have provided key insights
into actin cytoskeleton dynamics at the cell cortex of plant cells
(Augustine et al., 2011; Era et al., 2009; Henty et al., 2011;
Henty-Ridilla et al., 2013; Khurana et al., 2010; Konopka and
Bednarek, 2008; Li et al., 2012; Smertenko et al., 2010; Staiger
et al., 2009; Tóth et al., 2012). In striking contrast to in vitro actin
treadmilling (Bugyi and Carlier, 2010; Selve and Wegner, 1986), in
vivo remodeling of plant cortical actin arrays was found to follow a socalled ‘stochastic dynamics’ process that is dominated by fast
elongation and prolific severing of single filaments (Blanchoin et al.,
2010; Okreglak and Drubin, 2010; Staiger et al., 2009). Consistently,
single actin filaments have a surprisingly short lifetime, ,20 seconds
in Arabidopsis hypocotyl epidermal cells, and reach a maximum
length of 12–15 mm before disappearing. In this context, the
crosslinking of actin filaments into thick bundles emerges as a
strategy ‘used’ by cells to shape more stable and organized
cytoskeletal structures. Indeed, probably as a result of their higher
resistance to severing factors, actin bundles exhibit significantly
longer lifetimes (Smertenko et al., 2010; Staiger et al., 2009). In
addition, actin bundles appear stiffer than single actin filaments and
tend to align with the long cell axis (Era et al., 2009; Henty et al.,
2011; Staiger et al., 2009; Vidali et al., 2009).
Actin bundles are found in all plant cells and are involved
in cytoplasmic streaming and myosin-driven movement of
organelles and vesicles (Smith and Oppenheimer, 2005;
Thomas, 2012; Thomas et al., 2009; Walter and Holweg, 2008).
The role of actin bundles as long-distance tracks has been
particularly well described in pollen tubes, where longitudinally
arranged actin bundles enable the directional transport of Golgiderived vesicles from the shank to the apex (Ketelaar et al., 2002;
Lenartowska and Michalska, 2008; Miller et al., 1999; Ren and
Xiang, 2007; Tominaga et al., 2000). The recent characterization
of pollen-enriched villin, fimbrin and LIM proteins further
corroborates the importance of actin-bundling processes for
proper pollen tube elongation (Papuga et al., 2010; Staiger et al.,
2010; Wang et al., 2008; Wu et al., 2010; Zhang et al., 2010).
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Journal of Cell Science
Céline Hoffmann1,{, Danièle Moes1,{, Monika Dieterle1, Katrin Neumann1, Flora Moreau1,
Angela Tavares Furtado1, Dominique Dumas2, André Steinmetz1 and Clément Thomas1,*
Additional, more specific, roles have been assigned to actin
bundling during guard cell and chloroplast movements (Higaki
et al., 2010b), cell growth and morphogenesis (Baluska et al.,
2001; Higaki et al., 2010a; Nick, 2010; Nick et al., 2009; Smith
and Oppenheimer, 2005) and the set-up of plant defense
responses against pathogens (Clément et al., 2009; Day et al.,
2011; Hardham et al., 2007; Henty-Ridilla et al., 2013; Opalski
et al., 2005; Schmidt and Panstruga, 2007; Takemoto et al.,
2006).
Actin-bundling proteins exhibit a modular organization and
combine one or more actin-binding domains (ABDs), with
regulatory domains conferring sensitivity to specific stimuli
such as phospholipids or variations in pH or calcium
concentration (Bartles, 2000; Furukawa and Fechheimer, 1997;
Huang et al., 2006; Li et al., 2012; Papuga et al., 2010; Puius
et al., 1998; Wang et al., 2008). Despite the frequent pairwise
arrangement of their ABDs, some actin-bundling proteins
undergo dimerization to crosslink actin filaments (Mimura and
Asano, 1986; Thomas, 2012; Thomas et al., 2009). For instance,
dimerization as a prerequisite for actin-bundling activity has been
reported for members of the evolutionarily conserved villin and
formin families and for the recently described, plant-specific
SCAB1 protein (Chhabra and Higgs, 2006; George et al., 2007;
Harris et al., 2004; Li and Higgs, 2005; Michelot et al., 2006;
Michelot et al., 2005; Mimura and Asano, 1986; Xu et al., 2004;
Yokota et al., 1998; Zhang et al., 2012). By contrast, fimbrins use
the close proximity of their two ABDs to induce bundle formation
and function as monomers in yeast, animals and plants (Klein
et al., 2004; Nakano et al., 2001; Volkmann et al., 2001). We
previously reported that both LIM domains of plant LIM proteins,
which each consist of a double zinc finger motif, display intrinsic
actin-binding activity in vitro (Thomas et al., 2007). It is therefore
possible that plant LIMs crosslink actin filaments in a monomeric
form, although dimerization or oligomerization cannot be ruled
out. Indeed, a recent study focusing on the nuclear functions of
plant LIM proteins suggested that tobacco WLIM2 (NtWLIM2),
which also acts as an actin-bundling protein in the cytoplasm,
forms dimers in the nucleus of tobacco cells (Moes et al., 2013).
In support of dimerization, the mammalian counterparts of plant
LIM proteins, namely cysteine-rich proteins (CRPs), have been
reported to dimerize both in vitro and in live cells (Arber and
Caroni, 1996; Boateng et al., 2007; Feuerstein et al., 1994).
However, the oligomerization status of LIM proteins in the
cytoplasm of plant cells and, more particularly, along the actin
cytoskeleton has not been directly assessed so far. Despite the
importance of actin bundling in plant cells and the increasing
number of plant actin-bundling proteins identified, the questions
as how actin filaments are brought into contact by actin-bundling
proteins and how they are arranged inside the bundle have been
rarely addressed. A recent article by Khurana and co-workers
(Khurana et al., 2010) reported that Arabidopsis VILLIN1 and
VILLIN3 promote actin bundle formation by a so-called ‘catch
and zipper’ mechanism. However, whether this applies to other
types of plant actin-bundling proteins remains unclear. To the
best of our knowledge, the relative orientation of actin filaments
within bundles induced by a plant actin-bundling protein was
only reported in the case of the non-processive Arabidopsis
formin AFH1 (Michelot et al., 2006).
In the present work, we used state-of-the-art imaging
approaches to dissect the molecular mechanisms underlying
tobacco WLIM1 (NtWLIM1)-induced actin bundle formation in
both in vitro reconstituted assays and live cells. Together, our
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Journal of Cell Science (2014) 127, 583–598 doi:10.1242/jcs.134536
data support a multistep process in which self-association of
F-actin-induced NtWLIM1 operates as a driving force for the
zippering of NtWLIM1-decorated actin filaments.
RESULTS
NtWLIM1 binds to single actin filaments and promotes actin bundling
We previously reported that NtWLIM1 promotes the formation of
actin bundles in vitro and in live cells (Thomas et al., 2006;
Thomas et al., 2007). Here, we aimed to characterize NtWLIM1
bundling activity in a more quantitative manner as well as the
underlying molecular mechanisms. Fluorescently labeled actin
filaments (1 mM) were co-polymerized with 0–4 mM of NtWLIM1
and subsequently imaged by TIRF microscopy. The corresponding
pictures were pseudo-colored as a function of pixel fluorescence
intensities (see examples shown in Fig. 1A–C), whose distributions
were illustrated by histograms (supplementary material Fig. S1A–
C). In control experiments (no NtWLIM1), single actin filaments
formed a randomly organized array (Fig. 1A). In the presence of
NtWLIM1, filaments assembled into long and rather straight
bundles (Fig. 1B,C).
Noticeably, actin bundles grew larger with increasing NtWLIM1
concentrations. Fluorescence-intensity-based measurements revealed
that a maximal average number of 3–5 filaments per bundle was
Fig. 1. Quantification of NtWLIM1-induced actin bundling. A mix of 0.5 mM
unlabeled actin and 0.5 mM Alexa-Fluor-488-labeled G-actin was polymerized
in the presence of different NtWLIM1 concentrations ranging from 0 to 4 mM.
(A–C) Typical examples of TIRF microscopy images showing actin filaments
polymerized alone (A), with 1 mM NtWLIM1 (B) or 3 mM NtWLIM1 (C). Images
were pseudo-colored according to pixel intensity. Scale bars: 5 mm. (D) By
considering a direct correlation between the fluorescence intensity of actin
bundles and the number of filaments they include, we determined the number
of filaments per bundle. For each condition, the fluorescence intensity of at
least 150 bundles was measured and normalized to the signal of a single actin
filament from the same experiment. The central rectangle spans the 25th
percentile (first quartile) to the 75th percentile (third quartile) of all values; the
segment inside the rectangle shows the median (50th percentile or second
quartile). Bars denote the highest and lowest data point within 1.5 interquartile
range (IQR) starting from the upper and lower quartile, respectively. Open
circles indicate outliers. Asterisks indicate significant differences (P#0.0005,
based on Mann–Whitney U-test).
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reached at an actin-to-NtWLIM1 ratio of 1:3 (Fig. 1D). Consistently,
the same actin-to-NtWLIM1 ratio yielded the highest skewness value
(degree of asymmetry of pixel intensity distribution; Higaki et al.,
2010b; supplementary material Fig. S1D).
The distribution of NtWLIM1 along emerging actin bundles was
assessed using a NtWLIM1 protein labeled with HiLyte FluorTM
488 (subsequently referred to as LIM1-488). Remarkably, LIM1488 promoted the formation of actin bundles in a similar manner to
unlabeled NtWLIM1 (supplementary material Fig. S2A–E and
data not shown). As shown in supplementary material Fig. S2A–C,
LIM1-488 was distributed along the whole length of actin bundles.
To test the ability of NtWLIM1 to bind to individual filaments, we
lowered the LIM-488-to-actin ratio (1:1) and thereby kept a large
fraction of filaments in a non-bundled form (Fig. 1B). For a higher
resolution, samples were immobilized onto NEM-myosin-coated
coverslips and imaged by TIRF microscopy. LIM1-488 decorated a
mixture of bundles and fine fibers, which we assumed to be
individual, non-crosslinked actin filaments (supplementary
material Fig. S2D; asterisks and arrowheads, respectively).
Interestingly, as revealed by Rhodamine-Phalloidin co-labeling
(supplementary material Fig. S2E), the fluorescent signal intensity
due to LIM1-488 directly correlated with bundle thickness. A
close-up of the LIM1-488-decorated and Rhodamine-Phalloidin
co-labeled fine fibers (supplementary material Fig. S2F,G,
respectively) confirmed that they exhibited a shape and intensity
that was similar to those of control filaments polymerized without
LIM-488 (supplementary material Fig. S2H). We therefore
conclude that NtWLIM1 can efficiently bind to individual actin
filaments, and that this event probably precedes actin filament
crosslinking.
Real-time imaging of NtWLIM1-induced actin bundle formation in vitro
The formation of actin bundles induced by NtWLIM1 was
monitored by time-lapse TIRF microscopy. In the absence of
NtWLIM1, growing actin filaments occasionally co-aligned in a
transient manner, but never formed bundles of two or more
filaments over a long distance (.2 mm) and/or time (.5 second;
supplementary material Movie 1). The addition of NtWLIM1
promoted lateral contacts between filaments and their subsequent
crosslinking in parallel bundles. In two representative examples,
the extremity of an individual filament contacted a close-by fine
bundle in its terminal or more central region, respectively, and
subsequently fused with this bundle by a zippering-like process
(Fig. 2A,B; supplementary material Movies 2,3). NtWLIM1induced bundles were observed to elongate by two distinct
mechanisms. On the one hand, they grew at their terminal
regions by the addition of pre-formed short filaments (Fig. 2C;
supplementary material Movie 4). Interestingly, these short
filaments sometimes undertook several docking attempts before
finally joining the targeted bundle (Fig. 2D; supplementary
material Movie 5). These ‘touch and flip’ events probably reflect
local steric and/or electrostatic constraints, which favor a
preferential relative orientation between the added filament
fragments and growing bundles. On the other hand, bundle
elongation was achieved through polymerization of the inside
filaments (Fig. 2E, arrowheads; supplementary material Movie 6).
As shown by the kymograph in Fig. 2F, filaments elongate with
similar growth rates both inside and at the extremities of bundles.
In addition, we calculated similar average growth rates for
NtWLIM1-bundled actin filaments and individual actin filaments
polymerized alone (0.660.1 mm/minute), indicating that WLIM1
does not significantly alter the actin polymerization rate.
Journal of Cell Science (2014) 127, 583–598 doi:10.1242/jcs.134536
WLIM1-decorated actin filaments zipper into bundles in live cells
Advanced live cell imaging approaches have recently enabled
high-resolution monitoring of actin cytoskeleton remodeling in
plant cells (Henty et al., 2011; Li et al., 2012; Smertenko
et al., 2010; Staiger et al., 2009). To validate and extend our in
vitro biochemical data on LIM-protein-mediated actin bundle
assembly, we applied variable-angle epifluorescence microscopy
(VAEM) on Nicotiana benthamiana seedlings transiently
expressing NtWLIM1 fused to GFP (GFP-NtWLIM1). In
particular, we wanted to answer the following questions: does
NtWLIM1 bind to individual actin filaments in live cells? If so,
do such filaments undergo bundling through mechanisms similar
to those observed in in vitro reconstituted assays? In hypocotyl
epidermal cells, GFP-NtWLIM1 decorated single actin filaments,
which elongated by fast polymerization and disintegrated by
severing and depolymerization events (data not shown).
Moreover, we noticed pair-wise interaction between single
filaments or fine bundles, which subsequently zippered into
thicker fibers. Because the biolistic approach used to express
NtWLIM1-GFP in tobacco hypocotyl only resulted in a low
number of transformed cortical epidermal cells (the cell type
amenable to VAEM analysis) and variable transgene expression
levels, we switched to transgenic Arabidopsis seedlings stably
expressing the Arabidopsis WLIM1 protein fused to GFP (GFPAtWLIM1) to conduct more quantitative and statistically relevant
analyses. Noticeably, AtWLIM1 shares 86% sequence similarity
(79% sequence identity) with NtWLIM1 and exhibits equivalent
actin-binding and -bundling activities in vitro (Papuga et al.,
2010; Thomas et al., 2006). As in the tobacco system, GFPAtWLIM1 extensively decorated actin cytoskeleton components
in Arabidopsis hypocotyl cells, ranging from individual filaments
to thick bundles (Fig. 3A; supplementary material Movie 7). In
agreement with previous imaging studies using standard
fluorescent F-actin reporters (Smertenko et al., 2010; Staiger
et al., 2009) (Henty et al., 2011; Li et al., 2012; Staiger et al.,
2009; Tóth et al., 2012), actin cytoskeleton remodeling (Fig. 3A)
was dominated by fast polymerization and prolific severing
(Fig. 3B). Tracking of individual filaments revealed that many of
them were readily destroyed by severing and usually exhibited
lifetimes ,30 seconds. To check whether the binding of
AtWLIM1 to single actin filaments affects the dynamics of
these filaments in vivo, we quantified well-defined stochastic
dynamics parameters such as filament elongation rates, severing
frequency and convolutedness (Staiger et al., 2009) in the cortex
of hypocotyl epidermal cells from 5-day-old Arabidopsis
seedlings expressing GFP-AtWLIM1 or the widely used GFPfABD2 actin marker (supplementary material Table S1, Movie
8). Both filament elongation rates and severing frequency showed
statistically significant but modest differences in GFP-AtWLIM1
and GFP-fABD2 seedlings. In addition, filament convolutedness
was not statistically different in GFP-AtWLIM1 and GFP-fABD2
plants. Together, these findings indicate that overexpression of
WLIM1 is not detrimental to single actin filament dynamics. In
addition, every optical section analyzed over a period of
200 seconds revealed filaments that interacted with each other
and subsequently zippered into bundles (Fig. 3C; supplementary
material Movie 9). Since several zippering events could be
observed over the whole time of observation, we concluded that
these events are relatively frequent and represent a major mode of
actin bundle formation in vivo. Statistical quantification analyses
of these events in GFP-AtWLIM1 hypocotyl epidermal cells
yielded a bundling frequency of 2.06102466.761025 zippering
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Journal of Cell Science (2014) 127, 583–598 doi:10.1242/jcs.134536
events?mm22?second21 (supplementary material Table S1). A
parallel analysis conducted in the hypocotyls of seedlings
expressing GFP-fABD2 (Sheahan et al., 2004; Staiger et al.,
2009; Tóth et al., 2012) indicated a threefold lower frequency of
filament bundling processes (6.96102562.661025 zippering
events?mm22?second21). Together, these data are consistent
with an in vivo actin bundling activity of AtWLIM1. Compared
with individual filaments, GFP-AtWLIM1-labeled actin bundles
exhibited less-convoluted shapes and longer lifetimes (Fig. 3A;
supplementary material Movie 7). As in NtWLIM1-based in vitro
bundling assays, bundles grew by both elongation of internal
filaments and fusion/recycling of nearby-severed filament
fragments. In addition to straight bundles, some ring-shaped
bundles were observed to form through actin filament
circularization (supplementary material Movie 10). Similar
ring-shaped bundles have been previously referred to as
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acquosomes and have been proposed to serve as actin storage
organelles (Smertenko et al., 2010). However, because we never
observed acquosomes in vitro, their formation probably requires
additional or simply factors other than LIM proteins.
On the whole, live cell analyses are highly consistent with our in
vitro data and support a model in which LIM proteins bind to both
individual filaments and bundles and promote the zippering of
filaments or bundles that come into contact with each other.
Furthermore, the unaffected dynamics of single AtWLIM1-decorated
filaments support the idea that WLIM1 proteins promote actin
bundling directly through their actin-crosslinking activity.
WLIM1 proteins predominantly crosslink actin filaments in
antiparallel orientation
Depending on their respective functions, actin bundles can be
either of uniform polarity with the barbed ends of all filaments
Journal of Cell Science
Fig. 2. Real-time imaging of NtWLIM1-induced actin bundling in vitro. Alexa-Fluor-488-actin was co-polymerized with NtWLIM1 (ratio 1:3) in a glass flow
chamber and the formation and elongation of bundles were imaged by time-lapse TIRF microscopy. (A,B) Typical examples of NtWLIM1-induced actin
bundling by a ‘catch-and-zip’ mechanism. After docking onto an actin bundle, a single filament zips together with the targeted structure to form a thicker bundle.
Asterisks indicate the initial filament-docking site on the bundle. (C) Actin bundle elongation by consecutive addition of four small filaments. Each new filament is
labeled with a different color. The final relative positions of the newly integrated filaments are given in the last panel. (D) Time-lapse of single filament integration
into an actin bundle via ‘touch-and-flip’ mechanism: red and blue circles visualize filament redirection after a first integration attempt. (E) Actin filament
polymerization inside and at the extremity of a bundle (green arrowheads). (F) Kymograph of the two filaments traced in E. Scale bars: 3 mm.
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pointing in the same direction (parallel filaments) or of mixed
polarity (anti-parallel filaments). To measure the polarity of
NtWLIM1-induced actin bundles, we first measured actin bundle
polarity using a dual labeling fluorescence microscopy assay
(Harris et al., 2006). In brief, actin (1 mM) was co-polymerized
with NtWLIM1 (2 mM) and subsequently labeled by Alexa-Fluor488-phalloidin. These bundles were incubated with profilin-bound
actin monomers (mixed at a profilin-to-actin ratio of 4:1) in order
to inhibit actin polymerization at the pointed end of filaments and
resume filament elongation exclusively at the barbed end (Kovar
et al., 2006; Pantaloni and Carlier, 1993; Pollard and Cooper, 1984;
Pring et al., 1992). Finally, these newly polymerized bundle
sections were labeled with Rhodamine-Phalloidin. Data revealed
that 75% (n582) of NtWLIM1-induced bundles resumed bidirectional growth as indicated by red filaments emerging from
each extremity of the green initial bundle (Fig. 4A, left). The rest
of the bundles (25%) showed a unipolar conformation, with only
one of their extremities labeled in red (Fig. 4A, middle). By
contrast, control experiments with the human actin-bundling
protein fascin yielded almost exclusively unipolar bundles (90%,
n520) (Fig. 4A, right), a result consistent with the high selectivity
previously reported for fascin (Breitsprecher et al., 2011; Courson
and Rock, 2010; Ishikawa et al., 2003; Skau et al., 2011).
To further examine actin filament orientation in NtWLIM1triggered bundles, we traced fast barbed-end elongation of
individual and bundled filaments using real-time TIRF
microscopy (Fig. 4B–D). The majority of bundles (<90%)
contained several filaments that grew in opposite directions
(Fig. 4C). In addition, we regularly observed that anti-parallel
filaments, although elongating toward each other, crossed and
continued their growth along the bundle axis (Fig. 4D;
supplementary material Movie 11). In accordance with the
above dual-fluorescence labeling experiments, some NtWLIM1induced bundles (<10%) exhibited an exclusively parallel
filament orientation (data not shown). Together, these data
indicate that NtWLIM1 has no or only weak selectivity for actin
filament polarity and predominantly generates bundles of mixed
polarity.
To extend the above data, we tracked the orientation of
elongating filaments after they zippered into fine growing
bundles in Arabidopsis hypocotyl cells expressing GFPAtWLIM1 (supplementary material Movie 12). Data show that
55% of filaments that zipper into a thin bundle were elongating in
the same direction, whereas the rest of the analyzed population
(45%) grew in opposite directions. These findings are therefore
consistent with the above-characterized weak intrinsic selectivity
of NtWLIM1 for actin filament polarity. It should be noted that
we could not reliably characterize the relative orientation of actin
filaments in thick bundles. We assume that most of these bundles
have a mixed polarity, although this awaits confirmation.
NtWLIM1 self-associates along the actin cytoskeleton
The dimerization of NtWLIM1 in live cells as well as its potential
implication in actin bundling (Thomas et al., 2007) were directly
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Fig. 3. Real-time dynamics and
remodeling of GFP-AtWLIM1
decorated actin filaments at the
cell cortex. Hypocotyl epidermal
cells of transgenic Arabidopsis plants
expressing GFP-fused Arabidopsis
AtWLIM1 were submitted to timelapse VAEM. (A) Superposition of
three time-points (0 seconds in red,
10 seconds in green and 20 seconds
in blue) showing the extensive
remodeling of fine filaments (in color)
and the more stable pattern of thick
actin bundles (white). (B) Typical
sequential images showing fast actin
polymerization (red arrowheads) and
prolific filament severing (yellow
arrowheads). (C) Typical sequential
images of a short cell cortex region
illustrating the frequent zippering of
GFP-AtWLIM1-decorated actin
filaments. In this example, three
zippering events (highlighted with
colored dots) were observed in
30 seconds. Red arrowheads
indicate fast-growing filament ends.
Scale bars: 10 mm (A,B), 3 mm (C).
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assessed by two complementary approaches, namely bimolecular
fluorescence complementation (BiFC) (Hu et al., 2002; Hu and
Kerppola, 2003; Kodama and Hu, 2012) and fluorescence
resonance energy transfer based on fluorescence lifetime
imaging microscopy (FLIM-FRET) (Becker, 2012; IshikawaAnkerhold et al., 2012). In the first set of experiments, two BiFC
constructs, consisting of complementary N-terminal and Cterminal fragments of the enhanced yellow fluorescent protein
(eYFP) fused to NtWLIM1 (YN-NtWLIM1 and YC-NtWLIM1,
respectively) were co-expressed in tobacco BY-2 cells and
checked for their association. An intense signal of reconstituted
eYFP indicative of NtWLIM1–NtWLIM1 interaction was
observed in almost 90% of cells (Fig. 5Aa). Remarkably, as
revealed by treatment with the F-actin-disrupting drug
Latrunculin B (Lat B), NtWLIM1 BiFC complexes sharply
decorated the actin cytoskeleton (supplementary material Fig.
S3A–D). This contrasted with the dual diffuse cytoplasmic and
cytoskeletal localization observed when NtWLIM1 was fused to
full-length eYFP (Fig. 5Ab). Together, these data support the
idea that NtWLIM1 complexes concentrate along the actin
cytoskeleton and might hint at the existence of NtWLIM1
monomeric and di/oligomeric pools. However, we cannot exclude
the possibility that the diffuse cytoplasmic localization observed
for eYFP-NtWLIM1 results from the interference of the fulllength eYFP tag with NtWLIM1 self-association and/or by the
elevated levels of overexpressed eYFP-NtWLIM1 and resulting
cytoplasmic accumulation of the fusion protein occurring on top
of its appropriate cytoskeletal localization. No significant
fluorescence could be detected in control cells transformed with
the two empty BiFC vectors, nor with YN- or YC-NtWLIM1, on
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the one hand, and the complementary empty vector on the other
hand (data not shown). To confirm that the formation of
NtWLIM1 BiFC complexes was not only due to the close
vicinity of both NtWLIM1 BiFC constructs on the actin
cytoskeleton, we performed additional control assays with
complementary BiFC constructs respectively fused to WLIM1
and another actin filament-binding protein, namely the actin
binding domain 2 of Arabidopsis FIMBRIN1 (fABD2) (Ketelaar
et al., 2004). In contrast to the strong signal typically observed for
NtWLIM1–NtWLIM1 complexes, no fluorescence could be
detected in 75% of cells co-expressing YN-NtWLIM1 and YCfABD2 (Fig. 5Ac). The rest of cells (25%) exhibited significant
but relatively weak fluorescence (Fig. 5Ad). This BiFC signal
could be entirely abolished by pre-treating cells with Lat B (data
not shown), indicating that it was due to unspecific BiFC
complexes that formed along the cytoskeleton. However, as
previously stated, both the frequency and intensity of unspecific
actin-associated BiFC complexes were much lower than those
observed for NtWLIM1–NtWLIM1 BiFC complexes, supporting
the specificity of the latter.
In addition to BiFC analyses, NtWLIM1 self-interaction was
evaluated using FLIM-based FRET measurements (Becker, 2012;
Ishikawa-Ankerhold et al., 2012). Basically, the extent of
quenching of a donor fluorophore lifetime by an acceptor
fluorophore allowed us to quantitatively evaluate the physical
interaction between the respectively fused candidate proteins. Cterminal fusions of NtWLIM1 to AmCyan and eYFP were used
as donor and acceptor constructs (AmCyan-NtWLIM1 and
eYFP-WLIM1, respectively). For AmCyan-NtWLIM1 alone,
we measured a fluorescence lifetime of 2.8360.08 nseconds
Journal of Cell Science
Fig. 4. Filament orientation in NtWLIM1induced actin bundles. (A) Determination of
actin filament orientation within NtWLIM1assembled bundles by dual-labeling
fluorescence microscopy. Actin monomers
were first co-polymerized with either
NtWLIM1 or human fascin (in green), then
elongation was performed in the presence of
actin monomers (1 mM) and profilin (4 mM)
and visualized using Rhodamine-Phalloidin.
Red arrowheads indicate elongating bundle
extremities. (B–D) Growth orientation of
single filaments in NtWLIM1-triggered actin
bundles monitored by TIRF microscopy.
Images and associated kymographs were
derived from 12 minute time lapses. In the
absence of NtWLIM1, individual actin
filaments show one-directional elongation
(B, arrowheads). In the presence of NtWLIM1,
bundles elongate into opposite directions
owing to the polymerization of anti-parallel
orientated single filaments (C, arrowheads).
(D) Growing and intersecting of parallel and
anti-parallel filaments within one NtWLIM1induced bundle. Scale bars: 4 mm (A),
3 mm (B–D).
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Journal of Cell Science (2014) 127, 583–598 doi:10.1242/jcs.134536
Fig. 5. Characterization of NtWLIM1 self-association by BiFC and FLIMFRET analyses. (A) BiFC constructs were co-bombarded with an mRFPexpressing plasmid allowing the comparison of expression levels between
transformed cells (inserts in red). (a) BY-2 cells co-expressing YN-NtWLIM1
and YC-NtWLIM1 showed a sharp cytoskeleton-located signal of
reconstituted eYFP. (b) By comparison, full-length eYFP fused to NtWLIM1
shows a dual cytoskeletal and diffuse cytoplasmic distribution. Coexpression of YN-NtWLIM1 with YC-fABD2 yielded no fluorescence in 75%
(c) and low fluorescence in 25% of cells (d). Number of cells, n$30.
(B) Fluorescence intensity (gray scale, a,c,e) and corresponding lifetime
images (pseudo-color, b,d,f) of BY-2 cells expressing AmCyan-NtWLIM1
alone (a,b) or AmCyan-NtWLIM1 in the presence of eYFP-NtWLIM1 (c,d) or
eYFP-fABD2 (e,f). The histogram curves ranging from 1.5 nseconds
(orange) to 3.0 nseconds (blue) indicate occurrence frequency for values
within the lifetime image (b9,d9,f9). (C) Mean fluorescence lifetime (nseconds)
was calculated for cells expressing AmCyan-NtWLIM1 alone (white column)
or AmCyan-NtWLIM1 in the presence of eYFP-NtWLIM1 (dark gray column)
or YFP-fABD2 (light gray column). Number of cells, n$10. Mean FRET
efficiency values (percentage) are highlighted in bold, error bars indicate s.d.
Scale bars: 10 mm.
complexes (Fig. 5Bc–d9, arrowheads). Along actin bundles, we
measured an average lifetime of 2.4360.08 nseconds (Fig. 5C)
corresponding to a FRET efficiency of 14%. As a control, we
performed additional FLIM-FRET analyses on cells co-expressing
AmCyan and eYFP fusions of NtWLIM1 and fABD2. Although
eYFP-fABD2 decorated the actin cytoskeleton similar to AmCyanNtNtWLIM1 (supplementary material Fig. S4A–C), no significant
change of AmCyan-NtWLIM1 fluorescence lifetime was noticed
(Fig. 5Be–f9). Indeed, we measured an AmCyan-NtWLIM1
lifetime of 2.7560.07 nseconds, corresponding to a FRET
efficiency of 2.8% (Fig. 5C), a value below the typical 5%
significance threshold (Ciubotaru et al., 2007). Together, these data
confirm the BiFC analyses and provide compelling evidence that
NtWLIM1 self-associates along the actin cytoskeleton.
(Fig. 5Ba–b9,C), a value similar to those previously published for
AmCyan-based donor constructs (Ismail et al., 2010). In the
presence of eYFP-NtWLIM1, the overall fluorescence lifetime
of AmCyan-NtWLIM1 significantly decreased, indicating the
presence of NtWLIM1 complexes (Fig. 5B). Importantly,
particularly low values were reached along the actin cytoskeleton,
supporting the asymmetrical subcellular distribution of NtWLIM1
Both BiFC and FLIM-FRET analyses revealed NtWLIM1 selfassociation and highlighted the preferential localization of
NtWLIM1 complexes to the actin cytoskeleton. Because the
two LIM domains of NtWLIM1 (supplementary material Fig.
S5A) were previously shown to function as autonomous actinbundling modules in vitro (Thomas et al., 2007), we hypothesized
that they hold the ability to self-associate. Therefore, the single
LIM domains of NtWLIM1, namely LIM1 and LIM2, were fused
to either GFP (donor) or mRFP (acceptor) and assessed for their
interaction ability in BY-2 cells using FLIM-FRET analyses.
Consistent with the lack of actin-binding activity previously
reported for LIM1 and LIM2 in live cells (Thomas et al., 2007),
FLIM-FRET constructs exhibited a diffuse cytoplasmic
distribution (supplementary material Fig. S5B). Donor
constructs (GFP-LIM1 and GFP-LIM2) yielded an unquenched
GFP fluorescence lifetime average of 2.4560.04 nseconds and
2.4260.02 nseconds, respectively (Fig. 6Aa–b9 and data not
shown, respectively). When GFP-LIM1 was co-expressed with
mRFP-LIM1 or mRFP-LIM2, GFP fluorescence lifetime average
decreased to 2.3660.06 nseconds and 2.3360.09 nseconds,
respectively (Fig. 6Ac–d9 and e–f9). Similarly, when GFP-LIM2
was co-expressed with mRFP-LIM2, the overall GFP
fluorescence lifetime decreased to 2.3460.06 nseconds
(Fig. 6Ag–h9). In all cases, FRET efficiencies were below the
typical 5% threshold, suggesting no significant interaction
between LIM domains (Fig. 6B). However, we noticed on
lifetime images that obvious GFP quenching occurred in
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LIM domains are involved in NtWLIM1 self-association
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Journal of Cell Science (2014) 127, 583–598 doi:10.1242/jcs.134536
small restricted cytoplasmic areas (Fig. 6Ad,f,h, arrowheads), a
phenomenon that was not observed with donor constructs alone
(Fig. 6Ab). We therefore suspected that single LIM domains could
sporadically assemble into weak complexes that accumulate in the
whole cytoplasm. Supporting such a scenario, quantitative analyses
conducted on restricted positive cytoplasmic areas chosen from
lifetime images yielded FRET efficiencies ranging from 7.7% to
9.3% for the different LIM domain combinations (Fig. 6B). By
contrast, only background FRET efficiencies were obtained from coexpression of GFP-LIM1 with the mRFP fusion of either fABD2 or
the C-terminal domain of NtWLIM1 (2.4% and 3.3%, respectively).
Although the above data supported transient interactions
between LIM domains, this required further confirmation. Of
interest, BiFC complexes have been reported to over-stabilize
protein interactions, thereby displacing the equilibrium toward the
dimeric form of the proteins fused to the complementary eYFP
fragments. This property, which is sometimes considered to be a
disadvantage, provides the opportunity to reveal transient and weak
interactions (Ding et al., 2006; Hu et al., 2002; Kerppola, 2008;
Kerppola, 2009; Kodama and Hu, 2010; Robida and Kerppola,
2009; Shyu and Hu, 2008). We therefore conducted additional
590
BiFC experiments to assess self-association of single LIM
domains. Various combinations of LIM1 and LIM2 BiFC
constructs were tested. They all yielded an intense and diffuse
eYFP signal in the cytoplasm of BY-2 cells revealing, on the one
hand, the formation of LIM1 and LIM2 homomers in 100% and
80% of transformed cells, respectively (supplementary material
Fig. S6Aa,b; Fig. S6B) and the formation of LIM1–LIM2
heteromers in about 100% of transformed cells (supplementary
material Fig. S6Ac; Fig. S6B). Remarkably, no eYFP fluorescence
could be detected in cells co-transformed with either LIM1 or
LIM2 BiFC construct and a complementary fABD2 BiFC
construct (supplementary material Fig. S6Ad; Fig. S6B). In
conclusion, these data are highly consistent with our FLIMFRET analyses and confirm that LIM domains can form homo- and
heteromeric complexes, although they are much less stable than
full-length NtWLIM1 complexes.
Formation and/or stabilization of the NtWLIM1 complex requires an
intact actin cytoskeleton
The difference in the ability to self-associate between single LIM
domains and full-length NtWLIM1 is intriguing. Because,
Journal of Cell Science
Fig. 6. Quantitative FLIM-FRET analyses of
individual LIM domain interactions.
(A) Fluorescence intensity (a,c,e,g) and lifetime
images (b,d,f,h) of BY-2 cells expressing GFP-LIM1
alone (a,b), GFP-LIM1 in the presence of either mRFPLIM1 (c,d) or mRFP-LIM2 (e,f) and GFP-LIM2 in the
presence of mRFP-LIM2 (g,h). The histogram curves
ranging from 2.0 nseconds (orange) to 2.8 nseconds
(blue) indicate the occurrence frequency for values
within the fluorescence lifetime image (b9,d9,f9,h9).
Detail views in fluorescence (c,e,g) and lifetime
pictures (d,f,h) represent a twofold magnification of the
selected region of interest (squares) and highlight
restricted areas with a prominent decrease in
fluorescence lifetime (arrowheads). (B) Mean
fluorescence lifetime (nseconds) was determined on a
whole-cell level (black columns) or limited to
subcellular patches (gray columns) for cells
expressing either GFP-LIM1 alone or in the presence
of either mRFP-LIM1 or mRFP-LIM2, as well as for
cells expressing LIM2-GFP alone or in the presence of
mRFP-LIM2. Number of analyzed cells, n58. Mean
FRET efficiency values are indicated as percentages
and highlighted in bold. Error bars denote s.d.;
significant differences are indicated by asterisks
(P,1028, obtained by Student’s t-test).
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contrary to NtWLIM1, single LIM domains do not bind to the
actin cytoskeleton in live cells and because NtWLIM1 complexes
concentrate along actin bundles, we wondered whether the weak
interaction between single LIM domains could be related to their
inability to bind the actin cytoskeleton. The induction of protein
self-association by actin binding has already been reported for the
actin-bundling protein vinculin and, recently, the association of
villin with F-actin has been shown to promote villin dimerization
(George et al., 2013; Johnson and Craig, 2000). Based on these
observations, we wondered whether the actin cytoskeleton could
directly promote the formation and/or stabilization of NtWLIM1
dimers or oligomers.
To answer this question, we first investigated the assembly state
of recombinant NtWLIM1 in the absence of F-actin using sizeexclusion chromatography and dynamic light-scattering experiments.
Collectively, these approaches provide strong evidence that NtWLIM1
exhibits a single, lower, oligomeric state (data not shown). However,
the lack of knowledge about the spatial conformation of NtWLIM1
precludes a definite conclusion on the precise oligomeric state of
NtWLIM1. We thus performed sedimentation velocity analytical
ultracentrifugation experiments. The data established that NtWLIM1
has an elongated confirmation and behaves as a monomer
(supplementary material Fig. S7; Table S2).
None of the above approaches is appropriate to examine the
oligomeric state of NtWLIM1in the presence of F-actin. As an
alternative, we determined the NtWLIM1-to-actin molar ratio at
saturation in low-speed cosedimentation assays conducted at
fixed actin levels and increasing NtWLIM1 concentrations
(Fig. 7A,B). From four independent experiments, we calculated
an average of 1.9360.19 NtWLIM1 bound per actin subunit. This
result was very consistent with live cell data and further indicates
that NtWLIM1 switches from monomeric to a di- or oligomeric
state when associated with F-actin. In an attempt to directly
characterize NtWLIM1 complexes, the protein was incubated
with F-actin and the samples were subsequently mixed with a bmercaptoethanol-deprived buffer and analyzed by SDS-PAGE.
The NtWLIM1 species formed under these non-reducing
conditions were analyzed by western blotting. In control
samples without F-actin, NtWLIM1 was mostly detected at the
expected monomeric size (arrow at ,25 kDa; Fig. 7C, lane 1).
However, an additional very faint band of approximately twice
the size (double arrow at ,50 kDa) was scarcely detectable
supporting a predisposition of NtWLIM1 for self-association.
Consistent with this idea and the promoting effect of F-actin
previously indicated by cosedimentation data, the higher band
was markedly enhanced in the presence of F-actin (Fig. 7C, lane
2). In a control blot, actin was detected at its expected size, i.e.
,42 kDa (supplementary material Fig. S8). The ability of
NtWLIM1 to self-associate was further tested by incubating the
protein with the chemical crosslinker DFDNB. As expected,
DFDNB promoted the formation of 50 kDa NtWLIM1 complexes
(Fig. 7C, lanes 3 and 4). Additional higher bands of low intensity
suggest that NtWLIM1 can reach different oligomeric states
although unspecific chemical crosslinking of NtWLIM1 cannot
be excluded. Together, these data strongly suggest that NtWLIM1
self-association is promoted and/or enhanced by F-actin.
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Journal of Cell Science
Fig. 7. Impact of the actin cytoskeleton on NtWLIM1 protein assembly state. (A,B) Determination of NtWLIM1-to-actin ratio at saturation by low-speed
cosedimentation assays. Actin filaments (4 mM) were polymerized in the presence of increasing NtWLIM1 concentrations (0–24 mM). Samples were centrifuged
at 12,500 g, supernatant and pellet fractions were analyzed by SDS-PAGE and the respective amounts of NtWLIM1 were quantified. The ratio of actin-bound
NtWLIM1 versus total actin concentration was plotted against total NtWLIM1 concentration and data points were fitted to a hyperbolic function (A). Four
independent experiments yielded a mean value of 1.9360.19 NtWLIM1 molecules bound per actin subunit. For the representative experiment, a mean value of
1.94 was calculated from the cosedimentation gel shown in B. Filled circles indicate the set of experimental data points and blue lines give the 95% confidence
intervals of the fitted black curve. (C) Western blot analysis of NtWLIM1 dimer formation in vitro. 1 mM NtWLIM1 mixed (lane 2) or not (lane 1) with 8 mM F-actin
was incubated for 60 minutes under non-reducing conditions. 1 mM NtWLIM1 with DMSO (lane 3) or with a 20-fold molar excess of DFDNB (lane 4) was
incubated for 40 minutes under non-reducing conditions. After separation by SDS-PAGE omitting b-mercaptoethanol, samples were analyzed by western blot
using an anti-poly-histidine antibody.
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Journal of Cell Science (2014) 127, 583–598 doi:10.1242/jcs.134536
To confirm that efficient self-association of NtWLIM1 relies
on an intact actin cytoskeleton in live cells, additional FLIMFRET analyses were conducted on BY-2 cells treated with
the actin-disrupting drug Lat B. Data revealed that the donor
lifetime average (AmCyan-NtWLIM1) decreased in the presence
of eYFP-NtWLIM1 acceptor from 2.7760.03 nseconds to
2.6460.06 nseconds, corresponding to a FRET efficiency
below the significance threshold (4.75%; Fig. 8A,a,b9 versus
c,d9 and Fig. 8B). However, similar to what we observed with
single LIM domains, some restricted cytoplasmic areas exhibited
a greater reduction in AmCyan fluorescence lifetime (Fig. 8A;
arrowheads in d). Quantitative analysis of AmCyan fluorescence
lifetime limited to those areas yielded a lifetime average of
2.4960.06 nseconds, corresponding to a significant FRET
efficiency of 9.9% (Fig. 8B). By contrast, no such FRETpositive areas were observed in control experiments conducted
with eYFP-fABD2 (Fig. 8B,e,f9 and Fig. 8C). Thus, when
deprived of an intact actin cytoskeleton, NtWLIM1 exhibits a
reduced ability to form complexes.
Fig. 8. Impact of the actin cytoskeleton on NtWLIM1 protein
interactions in live cells. (A) Fluorescence intensity (a,c,e) and
lifetime images (b,d,f) of BY-2 cells treated with Lat B (2.5 mM) and
expressing either AmCyan-NtWLIM1 alone (a,b) or AmCyan-NtWLIM1 in
the presence of eYFP-NtWLIM1 (c,d) or eYFP-fABD2 (e,f). Histogram
curves ranging from 2.1 nseconds (orange) to 3.0 nseconds (blue)
indicate the occurrence frequency for values within the fluorescence
lifetime image (b,d9,f9). Arrowheads highlight areas of locally restricted
AmCyan-NtWLIM1 lifetime reduction (d). Scale bars: 10 mm.
(B) Mean fluorescence lifetime (nseconds) has been calculated for cells
expressing AmCyan-NtWLIM1 alone (white column) or in the presence of
either eYFP-NtWLIM1 (black column) or eYFP-fABD2 (light gray column).
Mean values for locally restricted decreases in AmCyan-NtWLIM1
fluorescence lifetime were calculated separately (dark gray column).
Number of cells, n$10. Mean values of FRET efficiency (indicated as
percentage) are highlighted in bold. Error bars denote s.d.; significant
differences are indicated by asterisks (P,10210, obtained by Student’s
t-test).
592
Actin bundles define a ubiquitous and highly abundant type of
cytoskeletal elements. In mammals, they play particularly
important roles in the formation and function of various
protrusive and contractile cellular structures such as filopodia,
microvilli, stereocilia, nerve growth cones and stress fibers. In
plants, the functional portfolio of actin bundles has steadily
expanded over the past years (Higaki et al., 2010a; Higaki et al.,
2010b; Thomas, 2012; Thomas et al., 2009). In addition to their
general role as tracks for myosin-dependent transport, actin
bundles were recently implicated in a series of more-specific
processes, including stomata and chloroplast movements,
polarization and recycling of auxin transporters or alteration of
turgor pressure by modifications in the cell wall, vacuole and
transvacuolar strand (Higaki et al., 2010a; Higaki et al., 2011;
Higaki et al., 2010b; Nick, 2010; Nick et al., 2009; Staiger et al.,
1994; Szymanski and Cosgrove, 2009).
Contrasting with our increasing knowledge about the functions
of actin bundles, relatively little is known regarding their mode of
formation. We recently characterized a novel family of small
actin-bundling proteins that are widely and abundantly expressed
in plants: the LIM proteins (Papuga et al., 2010; Thomas et al.,
2006; Thomas et al., 2007). Noticeably, counterparts of plant
LIM proteins are found in mammals, suggesting that this subset
of LIM domain proteins triggers basic, evolutionarily conserved,
functions (Weiskirchen and Günther, 2003). Like plant LIM
proteins, the so-called cysteine-rich proteins (CRPs) promote
actin bundle assembly in both reconstituted in vitro assays and
live cells (Jang and Greenwood, 2009; Kihara et al., 2011; Ma
et al., 2011; Tran et al., 2005). In addition, recent data support the
idea that CRP-bundling activity is involved in the stabilization of
stress fibers in smooth muscle cells and the formation of dendritic
filopodia (Kihara et al., 2011; Ma et al., 2011). However, the
molecular mechanisms underlying the actin-bundling activity of
plant LIM proteins and mammalian CRPs remain unknown. Here,
we provide evidence that NtWLIM1 promotes local interaction
between adjacent actin filaments and their subsequent zippering
into tight bundles. A similar two-step bundling process was
recently described for the Arabidopsis villin proteins VLN1 and
VLN3 (Khurana et al., 2010), fission yeast fimbrin Fim1 (Skau
et al., 2011), mouse TRIOBP (Kitajiri et al., 2010) and human
fascin (Breitsprecher et al., 2011), suggesting that it defines a
Journal of Cell Science
DISCUSSION
basic mechanism ‘employed’ by structurally unrelated actinbundling proteins. In addition, in vitro reconstituted assays and
live cell analyses conducted with NtWLIM1 and AtWLIM1,
respectively, show that both proteins can effectively bind to
individual unbundled actin filaments, suggesting that WLIM1
binding chronologically precedes formation of actin bundles.
In contrast to fascin, which almost exclusively assembles
unipolar bundles (90%), NtWLIM1 exhibits weak or no intrinsic
selectivity regarding polarity of actin filaments and induces a
majority (75–90%) of mixed polarity bundles in vitro. Data from
VAEM analyses confirm the weak selectivity of WLIM1 proteins
for actin filament polarity in live cells. So far, few studies have
addressed the orientation of actin filaments within bundles and
they have been limited to fixed tissues. Apart from pollen tubes
and root hair cells, where unipolar bundles have been
characterized by decoration with myosin S1 (Lenartowska and
Michalska, 2008; Tominaga et al., 2000; Yokota and Shimmen,
1999; Yokota et al., 2003), the polarity of bundles remains largely
unknown. With regard to these and our present data, we assume
that the formation of unipolar bundles in vivo (e.g. in pollen
tubes) requires factors other than LIM proteins: nucleators such as
formins (Michelot et al., 2006) and/or other, more selective,
actin-bundling proteins. Consistent with this hypothesis, plant
LIM proteins are highly expressed in various types of cell
(Arnaud et al., 2012; Eliasson et al., 2000; Papuga et al., 2010;
Wang et al., 2008) and are accordingly expected to contribute to
the formation of structurally different (unipolar and mixed
polarity) bundles.
The activity of several actin-crosslinking proteins was
previously suggested to rely on protein di- or oligomerization
(Bachmann et al., 1999; Johnson and Craig, 2000; Sanders et al.,
1996) and, more recently, dimerization of human villin was
reported along actin-bundle-rich structures in living cells and was
suggested to regulate actin crosslinking and filopodial assembly
by villin (George et al., 2013; George et al., 2007). Using a series
of biochemical approaches, we provide evidence that NtWLIM1
exhibits an almost exclusive monomeric conformation in the
absence of actin, but reaches a dimeric, and possibly higher
oligomeric, state(s) following incubation with F-actin. In support
of a close relationship between NtWLIM1 state and F-actin, the
sharp cytoskeletal localization of NtWLIM1 BiFC complexes
indicates that NtWLIM1 self-associates along actin filaments or
bundles. In addition, a strong FRET efficiency indicative of
substantial NtWLIM1 self-association was exclusively detected
along actin filaments, validating the accumulation of NtWLIM1
complexes at this location. The disruption of actin filaments by
Lat B was sufficient to decrease the overall FRET efficiency
below the typical significance threshold, pointing to the
implication of actin filaments in NtWLIM1 complex formation
or stabilization in vivo. Interestingly, we nevertheless observed
low, but significant, FRET efficiency in restricted cytoplasmic
areas, suggesting that NtWLIM1 sporadically self-associates in
the cytoplasm but does not form stable complexes without actin
filaments. Consistent with this hypothesis, weak NtWLIM1
complexes in F-actin-free samples could be resolved by western
blot following chemical crosslinking.
Together, our results not only draw a simple parallel between
WLIM1 self-association and cytoskeletal localization, but also,
and above all, disclose a causal link between interaction of
WLIM1 with actin filaments and formation or stabilization of
WLIM1 di- or oligomers. Similar scenarios of actin-bindinginduced protein oligomerization have been reported for vinculin
Journal of Cell Science (2014) 127, 583–598 doi:10.1242/jcs.134536
and villin – two non-related actin-bundling proteins (George
et al., 2013; Johnson and Craig, 2000). These findings, in concert
with our results, hint at a mechanism common to structurally
different actin-bundling proteins that allows the control of actinbundling by actin itself.
We previously reported that, although individual LIM domains
of NtWLIM1 can bind to and bundle actin filaments in vitro, they
retain an affinity that is too low to interact with the actin
cytoskeleton in live cells (Thomas et al., 2007). On the basis of
these data and the here-established role of actin filaments in
NtWLIM1 self-association, it can be predicted that if individual
LIM domains hold the ability to self-associate, the resulting
complexes should exhibit low stability. In line with this
assumption, FRET analyses conducted with various
combinations of individual LIM domains revealed discrete
cytoplasmic areas exhibiting positive but weak FRET
efficiencies (ranging from 7% to 9%) which were strikingly
similar to the FRET efficiency calculated for full-length
NtWLIM1 in Lat-B-treated cells (9.9%). Noticeably, selfinteraction of LIM domains was confirmed in BiFC assays,
which upon eYFP complementation generate irreversible
complexes and thereby allow the characterization of weak
protein interactions. In conclusion, NtWLIM1 self-association
relies, at least in part, on LIM-domain-based interactions, which
might, in turn, be facilitated by actin filaments. We thus end with
a multi-step mechanism in which LIM domains sequentially
function as actin-binding, protein dimerization and actin-bundling
modules. Based on our data, we propose a model for actin
crosslinking by WLIM1 dimers or oligomers (Fig. 9). In the
cytoplasm, WLIM1 mainly exhibits a monomeric conformation.
Although dimers and possibly higher-order oligomers
sporadically assemble in the cytosol, they are not stable and the
equilibrium is thus displaced toward the monomeric state. Upon
binding to actin filaments through its LIM domains, WLIM1
becomes more competent for dimerization or oligomerization.
Such an increase in competency of self-association might result
from a conformational change of LIM domains induced by their
interaction with actin filaments. Through its LIM-domaintriggered self-interaction, WLIM1 brings neighboring actin
filaments or bundles into contact and promotes their zippering
without marked selectivity for the polarity of actin filaments.
Considering the extremely fast cytoskeletal turnover observed
in plant cells as well as the ubiquitous and high expression of
plant LIM proteins, such a mechanism might correspond to a
default cellular program by which actin filaments increase their
own stability and lifetime. Our data might have wider
implications and, for instance, could lead to a better
understanding of why the human LIM protein counterpart
CRP3 (also known as muscle LIM protein) dimerizes or
oligomerizes in the cytoplasm but remains exclusively
monomeric in the nucleus, and how shuttling of CRP3 between
both subcellular compartments is regulated (Boateng et al., 2007;
Boateng et al., 2009). These are particularly important questions
because abnormalities in the subcellular distribution of CRP3
have been associated with the pathogenesis of heart failure.
Future work should confirm and characterize the conformational
changes triggered by LIM domain binding to actin filaments and
leading to protein dimerization or oligomerization. More than 60
genes containing one or more LIM domains have been identified
in the human genome and the list of their specific partners
is continuously expanding (Kadrmas and Beckerle, 2004).
Thus, it would be highly worthwhile to examine whether a
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Journal of Cell Science (2014) 127, 583–598 doi:10.1242/jcs.134536
Fig. 9. Model for actin bundling by WLIM1 selfassociation. For space reasons and ease of
understanding, the present model only considers
WLIM1 dimerization, although higher
oligomerization states for WLIM1 should be
considered as an alternative. (1) Dimers of WLIM1
sporadically form in the cytosol but they are unstable
and the equilibrium is displaced toward the
monomeric form. (2) WLIM1 binds to actin filaments
through its two LIM domains and thereby acquires a
higher competency for dimerization (stars). (3,4)
Through its dimerization, WLIM1 promotes
interactions between actin filaments (3) and their
subsequent zippering into tight bundles (4). WLIM1
assembles both unipolar and mixed polarity
bundles, a property that probably results from the
capacity of WLIM1 N- and C-terminal LIM domains
to form homo- and heterodimers. WLIM1-induced
bundles elongate by both polymerization of
crosslinked actin filaments (5) and addition of short
filament or bundle fragments at their extremities (6).
conformational change of LIM domains upon partner interaction
is a general process by which LIM-domain-containing proteins
trigger functional switches between their partners and/or regulate
their own activity.
MATERIALS AND METHODS
Denver, CO) was reconstituted at 116 mM. Rabbit muscle actin-AlexaFluor-488 (200 mM) was purchased from Molecular Probes (Life
technologies, Merelbeke, Belgium). Before each experiment, a 5 mM
solution of G-actin in G Buffer (2 mM Tris-HCl, pH 8.0, 0.2 mM CaCl2,
0.4 mM DTT, 0.2 mM ATP) was prepared. After overnight incubation
on ice, G-actin was centrifuged for 30 minutes at 100,000 g to remove
actin oligomers.
Plasmid construction
Co-labeling of recombinant NtWLIM1 protein along
actin structures
A previously described pQE60 construct allowing the production of
recombinant 66His-tagged NtWLIM1 protein (Thomas et al., 2006) was
expressed in E. coli and NtWLIM1-66His was purified under native
conditions according to the manufacturer’s instructions (Qiagen, Hilden,
Germany) and dialysed against 50 mM Tris-HCl, pH 8, 200 mM NaCl,
5 mM b-mercaptoethanol. For dye-coupling, the protein was purified and
buffer-exchanged with 10 mM MOPS, pH 7.4, 150 mM NaCl using a
Superdex 75 10/300 column (GE healthcare). Fractions containing
NtWLIM1-66His were combined and concentrated using centrifugal
filters (Ultracell 10K; Amicon, Merck, Darmstadt, Germany).
NtWLIM1-66His was labeled using the AnaTagTM Hilyte FluorTM 488
Protein labeling Kit (Anaspec, Fremont) according to the manufacturer’s
instructions using a protein-to-dye ratio of 1:2. Micro Bio-SpinH
Chromatography Columns (Bio-Gel P-6; Bio-Rad, Nazareth Eke,
Belgium) were used to remove unincorporated dye and to exchange the
buffer to 10 mM MOPS, 150 mM NaCl, pH 7.4. Labeling efficiency and
protein concentration were determined spectrophotometrically (Implen
NanoPhotometerTM). Lyophilized rabbit muscle actin (Cytoskeleton,
594
Low-speed actin cosedimentation assays
Low-speed cosedimentation assays were performed with NtWLIM166His purified under native conditions and dialysed against 10 mM TrisHCl, pH 7, 150 mM NaCl, 1 mM DTT, 50 mM ZnCl2. Following protein
pre-clarification at 200,000 g, 4 mM of rabbit muscle actin
(Cytoskeleton) were polymerized in the presence of increasing
NtWLIM1 concentrations (0–24 mM) for 1 hour at room temperature in
5 mM Tris-HCl, pH 7, 0.2 mM CaCl2, 50 mM KCl, 2 mM MgCl2,
0.4 mM ATP and 0.4 mM DTT. Samples were then centrifuged for
30 minutes at 12,500 g. The resulting pellet and supernatant fractions
were analyzed by SDS-PAGE and the respective amounts of NtWLIM1
were quantified using ImageJ (http://rsbweb.nih.gov/ij/). To calculate the
molar NtWLIM1-to-actin ratio at saturation, the ratio of bound
NtWLIM1 versus total actin concentration was plotted against the
respective total NtWLIM1 concentration and the data points of one
experiment were fitted to a hyperbolic function (SigmaPlotH10 software).
The value (6 s.d.) indicated in the Results corresponds to the mean of
four independent experiments.
Dual-labeling fluorescence microscopy assays
F-actin was obtained following a 30 minute polymerization of 1 mM Gactin in 16 KMEI buffer (10 mM Imidazole, pH 7, 50 mM KCl, 1 mM
EGTA, 1 mM MgCl2). To localize NtWLIM1 on actin bundles, F-actin
was mixed with Hilyte-Fluor-488-labeled NtWLIM1 protein (LIM1-488)
at a 1:2-ratio. After 15 minutes, the sample was labeled with RhodaminePhalloidin and diluted 20-fold in fluorescence buffer (16 KMEI supplied
with 100 mM DTT, 0.5% methylcellulose, 20 mg/ml catalase, 100 mg/ml
glucose oxidase, 15 mM glucose) before imaging. Orientation of actin
filaments in WLIM1-induced actin bundles was determined by a duallabeling fluorescence assay (Harris et al., 2006). In brief, actin monomers
(1 mM) were polymerized for 10 minutes in the presence of NtWLIM1
(2 mM) or fascin (250 nM). After addition of Alexa-Fluor-488-phalloidin
(Life technologies, Belgium) during 5 minutes, actin filaments were
washed in polymerization buffer and centrifuged for 5 minutes at
55,000 g. A second elongation step of 15 minutes was performed with a
mix of actin monomers (1 mM), profilin (4 mM), NtWLIM1 protein
(2 mM) and Rhodamine-Phalloidin. Bundles were diluted in two volumes
Journal of Cell Science
Coding sequences of NtWLIM1, its two LIM domains (LIM1 and LIM2)
and fABD2 were amplified by PCR (for primers and templates see
supplementary material Table S3). For BiFC analyses, coding sequences
were ligated into vectors pSPYNE(R)173 and pSPYCE(MR) (Waadt et al.,
2008), which allow the expression of proteins fused to the N-terminal part
(YN, amino acids 1–173) or C-terminal part (YC, amino acids 156–239) of
eYFP, respectively. Constructs used for FLIM-FRET analyses were mainly
derived from previously described in-house plasmids pNTL2103 and
pNTL3103 (Thomas et al., 2006; Thomas et al., 2007), where existing GFP
fusion constructs of NtWLIM1, LIM1, LIM2 and the C-terminal domain
were changed into the reporter gene fusions of interest (AmCyan-, eYFPor mRFP-fusions, respectively). The FLIM control plasmid expressing
eYFP-fABD2 was a derivative of the pSPYNE(R)173 vector (Waadt et al.,
2008), in which the initial N-terminal eYFP fragment was replaced with
full-length eYFP and the fABD2 coding sequence (amino acids 325–687)
was subcloned in-frame with eYFP.
Journal of Cell Science (2014) 127, 583–598 doi:10.1242/jcs.134536
of fluorescence buffer, placed on a poly-L-Lysine-coated slide and
imaged by confocal microscopy. Bundles formed during the first copolymerization step are imaged in green, filament elongations arising
from the second step are shown in red. The experiment was performed
three times and 82 filaments were observed.
emitted light was collected with a 505–530 nm band pass filter. RFP and
Rhodamine-Phalloidin were excited with the 543 nm helium neon laser
line and emitted light was collected by a 580–650 nm band-pass filter.
Stacks with 0.4 mm optical sections were captured and processed for
deconvolution using Huygens essential software (SVI, Netherlands).
Analytical ultracentrifugation (AUC), dynamic light scattering
(DLS) and crosslinking experiments
TIRF microscopy and VAEM live cell imaging
After purification of NtWLIM1-66His, elution buffer was exchanged
using size exclusion chromatography to buffer A, B or C using a
Superdex 75 10/300 column (GE healthcare). Sedimentation velocity
AUC experiments were performed in an XL-I analytical ultracentrifuge
(Beckman-Coulter) at 4 ˚C and 50,000 rpm using double-sector charcoalfilled Epon centerpieces (1.2 cm) with sapphire windows. Measurements
were carried out in PBS buffer (buffer A) or 50 mM Tris-HCl, pH 7.5,
150 mM NaCl, 10 mM ZnCl2 and 5 mM 2-mercaptoethanol (buffer B) or
50 mM Tris-HCl, pH 7.5, 50 mM NaCl, 10 mM ZnCl2 and 5 mM 2mercaptoethanol (buffer C) with 400 ml of each sample. Absorbance
scans were taken at 280 nm every 5 minutes for 16 hours. The partial
specific volume of NtWLIM1-66His was calculated from the amino acid
composition to be 0.7177 ml?g21 using the program SEDNTERP
(SEDNTERP, Sedimentation Interpretation Program. Philo, J., Hayes,
D. and Laue, T. Alliance Protein Laboratories, Thousand Cloaks, CA).
The sedimentation data were analyzed with program SEDFIT using the
continuous c(s) distribution analysis (Schuck, 2000). To analyze
NtWLIM1 self-association under non-reducing conditions, the protein
was first eluted from a Ni-NTA matrix, then buffer exchanged by dialysis
to 10 mM MOPS, 150 mM NaCl, pH 7.7. Actin was exchanged for
modified G-Buffer (5 mM MOPS, pH 7.5, 2 mM ATP, 0.2 mM CaCl2).
NtWLIM1 was copolymerized with 8 mM actin (final buffer
composition: 7 mM MOPS, pH 7.5, 100 mM KCl, 5 mM ATP, 2 mM
MgCl2, 0.2 mM CaCl2). For crosslinking experiments 1 mM stock
solution of 1,5-difluoro-2,4-dinitrobenzene (DFDNB; Thermo Scientific)
was freshly prepared in DMSO. The final concentration of DMSO in the
sample was 2%. Proteins were separated by non-reducing SDS-PAGE
and subsequently analyzed by western blot using a poly-histidine or actin
antibody (Sigma, H1029 and abcam, ab7813, respectively).
Cell culture and transformation
BY-2 (Nicotiana tabacum L. cv. Bright Yellow-2) cell suspensions were
maintained in the dark on a rotary shaker (27 ˚C, 130 rpm). Every week,
3 ml of 7-day-old BY-2 cell culture were transferred into 80 ml fresh
BY-2 medium.
For biolistic transformation, 3 ml of 3-day-old BY-2 cells were filtered
onto a Whatman filter and placed on 0.8% BY-2 agar plates. Particle
preparation and biolistic assays were performed as previously described
(Thomas et al., 2007) with the following modifications: 3 mg of gold
particles (,10 mm, Sigma-Aldrich, Diegem, Belgium) prepared in 50%
glycerol were coated with 5 mg of each plasmid in the presence of 1 M
CaCl2 and 16 mM spermidine in a final volume of 60 ml. After ethanol
precipitation, coated particles were resuspended in 25 ml of absolute
ethanol. Biolistic transformation was achieved with 6 ml of DNA particle
solution using the particle delivery system Biolistic PDS-1000/He from
Bio-Rad. Each transformation was performed under vacuum at 1100 psi.
Transformed cells were incubated in the dark at 27 ˚C for 24 hours and
then observed by confocal microscopy. To disrupt actin cytoskeleton
structures, BY-2 cells were incubated 5 hours prior to bombardment on
BY-2 agar plates supplied with 2.5 mM Lat B. After transformation, cells
were kept on the same Lat B agar plates for 24 hours.
Confocal laser-scanning microscopy
Cells were observed with a Zeiss LSM 510META laser-scanning
confocal microscope. For BiFC experiments, cells were mounted in
BY-2 medium between slide and coverslip 24 hours after bombardment.
The same settings were used for all samples with a gain set at 56%.
Observations were performed using a 636 oil immersion (NA 1.4) Plan
Apochromat objective. Complemented eYFP and Alexa-Fluor-488phalloidin were excited with the 488 nm line of an Argon laser and
For TIRF microscopy, glass flow chambers of ,50 ml were prepared as
previously described (Breitsprecher et al., 2009). Chambers were perfused
with 2.5 nM NEM-myosin in myosin buffer (10 mM imidazole, pH 7.0,
0.5 M KCl, 10 mM MgCl2). After 1 minute, the chamber was washed with
myosin buffer, then 1% BSA was flowed into the chamber and incubated for
4 minutes. The chamber was finally washed with fluorescence buffer. A
mixture of Alexa-Fluor-488-labeled and unlabeled G-actin (1 mM) with or
without actin binding protein (diluted in protein dialysis buffer) was prepared
in 40 ml of fluorescence buffer; allowed to flow into the chamber and imaged
immediately. Actin filaments were imaged by TIRF microscopy on a Zeiss
inverted microscope equipped with an alpha-Plan Apochromat 1006 /1.46
TIRF objective. Excitation ray 488 nm was provided by an argon laser and
emission light was collected with a BP filter 525/50. Time-lapse images were
acquired at 5 second intervals over 25 minutes with a Zeiss Axiocam HRm
camera. Typical exposure time was 700 mseconds with a laser filter
of 5%. Pictures were analysed via ImageJ and, if necessary,
x-y drift during TIRF timelapses was corrected by the TurboReg
plugin (http://bigwww.epfl.ch/thevenaz/turboreg/). Kymographs were built
along actin filaments or bundles with the MultiKymograph plug-in (http://
www.embl.de/eamnet/html/body_kymograph.html). Skewness was measured
after 40 minutes of G-actin polymerization with WLIM1 protein concentrations increasing from 0 to 4 mM. Measurements were performed on three
independent experiments with 40 pictures using the plugin Kbi_Filter2d
(ThinLine) (Higaki et al., 2010b). For better visualization of skewness,
representative pictures of one ratio are presented with a custom LUT Fire. To
quantify actin bundle thickness, we supposed that the Alexa Fluor 488
fluorescence signal was proportional to the amount of actin present within
analyzed bundles (Breitsprecher et al., 2011; Smertenko et al., 2010). The
absolute signal of each bundle was measured and normalized to the
fluorescence intensity of a single actin filament to obtain the number of
filaments per bundle.
The cortical actin cytoskeleton in epidermal cells from Arabidopsis
hypocotyls expressing Arabidopsis GFP-AtWLIM1 (Papuga et al., 2010)
was examined using time-lapse VAEM (Konopka and Bednarek, 2008).
Transgenic seeds were stratified for 2 days at 4 ˚C, then exposed for
24 hours to white light and finally grown in the dark for 6 days (both at
21 ˚C). Seedlings were mounted in water between slide and coverslip and
analyzed using the above-described TIRF microscope platform and
imaging equipment. Images were captured at 1–2 second intervals and
filament bundling was examined.
Quantitative analyses of actin filament dynamics
Parameters of actin filament dynamics were measured in the cortex of
epidermal cells from the middle third of 5-day-old, dark-grown
Arabidopsis GFP-fABD2 (Sheahan et al., 2004) and GFP-NtWLIM1
hypocotyls (Papuga et al., 2010). Seedlings were imaged over
200 seconds with 1.3 second intervals and image sequences were
imported into ImageJ. At least 50 filaments from 10 cells in at least
five seedlings were tracked to calculate convolutedness and severing
frequency according to Staiger et al., 2009. Briefly, severing frequency of
filaments was evaluated by measuring the maximal length of one
filament and then counting breaks along this filament over time. Only
long filaments (.10 mm) were considered and tracked as long as possible
to document severing events. Severing frequency was calculated as the
number of breaks per length of the original filament per time (breaks/mm/
second). Filament flexibility was described by convolutedness, which
indicates how much a given line deviates from a straight line.
Convolutedness was defined as the ratio of free-hand-traced filament
length to the longest side of the bounding rectangle (series of $10 frames
were observed). Frequently-appearing acquosomes were not taken into
consideration. Elongation rates were determined by tracking filament
595
Journal of Cell Science
RESEARCH ARTICLE
assembly over at least four time frames. The frequency of actin filament
bundling was calculated as the average of catch and zip events counted
within ROI-squares of 196 mm2 over a time span of 200 seconds. To
quantify actin-bundling events, a total number of 36 ROI-squares from at
least 12 cells and six seedlings were analyzed.
FLIM-FRET data acquisition
Lifetime imaging was done on a confocal microscope Leica SP2 with a
time-correlated single photon counting module TCSPC-730 (Becker &
Hickl). BY-2 cells were observed in BY-2 medium with a 406 waterimmersion objective. Samples containing AmCyan fusion proteins were
excited by a white laser at 470 nm (75% laser power), samples with
eGFP constructs with the same laser at 488 nm (20% laser power). To
avoid a pulse pile-up effect, laser power was adjusted to an average
photon-counting rate of 105–106 photons?seconds21. Fluorescence was
detected with a 480–500 nm band pass filter. Typically, the samples
were continuously scanned for about 120 seconds to achieve sufficient
photon statistics for the fitting of the fluorescence decays. Data were
analyzed using a commercial software package (SPCImage V2.8;
Becker and Hickl). Curves were fit to a mono-exponential decay with a
x2 close to 1. FRET efficiency was calculated as % FRET512(tDA)/
(tD), where tDA is fluorescence lifetime of donor in the presence of
acceptor; tD, the fluorescence lifetime of donor alone. Fluorescence
lifetime was collected along bundles that showed colocalization of
AmCyan and eYFP and, for each cable, three points were randomly
selected; 30 actin bundles from 10 cells were analyzed. For Lat-Btreated samples and for single-LIM-domain analyses, regions of interest
(ROIs) showing homogenous fluorescence intensity were selected (three
ROIs/cell) from eight cells.
Acknowledgements
We thank Christopher Staiger and Jessica Henty (Purdue University, USA) for
TIRF microscopy and VAEM imaging training, and for the NEM-myosin supply.
We are grateful to Jörg Kudla (University of Münster, Germany) for providing us
with BiFC plasmids and David Kovar (University of Chicago, USA) for the gift of
human fascin. We thank Annette Kuehn and Stéphanie Kler (Laboratory of
Immunogenetics and Allergology, CRP-Santé, Luxembourg) for support in size
exclusion chromatography and Petr Nazarov (Microarray Center, CRP-Santé,
Luxembourg) for advice on statistical analyses. We thank Catherine Birck for her
expertise and assistance in AUC and DLS measurements (Structural Biology and
Genomics Platform, IGBMC, Illkirch, France).
Competing interests
The authors declare no competing financial interests.
Author contributions
Authors engaged in conducting experiments: C.H., D.M., M.D., K.N., F.M., A.T.F.;
experiment design, evaluation and interpretation: C.H., D.M., D.D., A.S. and C.T.;
writing the manuscript: C.H., D.M., C.T.
Funding
This work was supported by the Ministry of Culture, Higher Education and
Research [grant number REC-LBMV-20100902]. Financial support from the
National Research Fund Luxembourg (FNR) is gratefully acknowledged [grant
number C10/BM/784171-HUMCRP].
Supplementary material
Supplementary material available online at
http://jcs.biologists.org/lookup/suppl/doi:10.1242/jcs.134536/-/DC1
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