Download The plant formin AtFH4 interacts with both actin and microtubules

Short Report
1209
The plant formin AtFH4 interacts with both actin and
microtubules, and contains a newly identified
microtubule-binding domain
Michael J. Deeks1,*, Matyáš Fendrych2,3,*, Andrei Smertenko1, Kenneth S. Bell4, Karl Oparka4,
Fatima Cvrcková2, Viktor Zársky2,3 and Patrick J. Hussey1,‡
1
School of Biological and Biomedical Sciences, University of Durham, South Road, Durham DH1 3LE, UK
Department of Plant Physiology, Faculty of Sciences, Charles University, Prague 12844, Czech Republic
3
Institute of Experimental Botany, Academy of Sciences of the Czech Republic, Prague 16502, Czech Republic
4
Institute of Molecular Plant Sciences, University of Edinburgh, Mayfield Road, Edinburgh EH9 3JR, UK
2
*These authors contributed equally to this work
‡
Author for correspondence ([email protected])
Journal of Cell Science
Accepted 25 January 2010
Journal of Cell Science 123, 1209-1215
© 2010. Published by The Company of Biologists Ltd
doi:10.1242/jcs.065557
Summary
The dynamic behaviour of the actin cytoskeleton in plants relies on the coordinated action of several classes of actin-binding proteins
(ABPs). These ABPs include the plant-specific subfamilies of actin-nucleating formin proteins. The model plant species Arabidopsis
thaliana has over 20 formin proteins, all of which contain plant-specific regions in place of the GTPase-binding domain, formin homology
(FH)3 domain, and DAD and DID motifs found in many fungal and animal formins. We have identified for the first time a plantspecific region of the membrane-integrated formin AtFH4 that mediates an association with the microtubule cytoskeleton. In vitro
analysis shows that this region (named the GOE domain) binds directly to microtubules. Overexpressed AtFH4 accumulates at the
endoplasmic reticulum membrane and co-aligns the endoplasmic reticulum with microtubules. The FH1 and FH2 domains of formins
are conserved in plants, and we show that these domains of AtFH4 nucleate F-actin. Together, these data suggest that the combination
of plant-specific and conserved domains enables AtFH4 to function as an interface between membranes and both major cytoskeletal
networks.
Key words: Actin, Actin regulating proteins, Membrane, Microtubule
Introduction
The activities of the cytoskeleton impact multiple aspects of plant
biology (for a review, see Hussey et al., 2006). In addition to
supporting mitosis and subsequent cell division, actin filaments and
microtubules also guide cell-wall synthesis, endomembrane
trafficking and organelle motility. The organisation of the filaments
of the eukaryote cytoskeleton requires many accessory proteins,
including the formin family of actin-nucleating proteins. The model
plant species Arabidopsis thaliana contains more than 20 formin
isoforms. These can be divided according to sequence similarity
and domain structure into two distinct groups (groups I and II). The
divergence of these two groups is likely to be ancient, as
representatives of both can be found in mosses (Grunt et al., 2008).
Like their animal and fungal homologues, the formin homology
1 and 2 (FH1 and FH2) domains of group I plant formins modify
actin dynamics. The investigation of several isoforms in vitro has
identified F-actin nucleation, bundling, capping and severing
activities (Ingouff et al., 2005; Michelot et al., 2005; Deeks et al.,
2005; Yi et al., 2005). Extensive biophysical studies exploiting total
internal reflection fluorescence (TIRF) microscopy monitored the
behaviour of the FH1-FH2 domains from group I isoform AtFH1
(Michelot et al., 2005; Michelot et al., 2006). The FH1-FH2 unit
establishes filament polymerisation, but does not maintain a longterm association with the barbed end; this non-processive behaviour
is accompanied by an ability to enhance the bundling of F-actin
through a process of ‘zippering’ (Michelot et al., 2006). The FH1FH2 unit can simultaneously nucleate and bundle filaments in vitro,
creating actin cables that thicken by incorporating filaments that
have originated at the cable through de novo nucleation.
Ten of the eleven group I isoforms of Arabidopsis have an Nterminal secretory signal sequence followed by a transmembrane
domain. This domain architecture of formins has not been
recognised beyond the Plantae kingdom [with the possible exception
of a few invertebrate metazoans and protists (Grunt et al., 2008)].
The overexpression of full-length AtFH1 within pollen tubes causes
excessive bundling and membrane-associated accumulation of Factin in tip-growing cells (Cheung and Wu, 2004). Localisation of
this activity at the plasma membrane was dependent on the
signalling and transmembrane motifs of the AtFH1 N terminus,
confirming that the unique domain structure of group I formins is
capable of coupling aspects of the biophysical behaviour observed
in vitro to membrane processes in vivo. Another group I formin,
AtFH6, localises to the plasma membrane in expanding giant cells
of nematode-induced galls (Favery et al., 2004), again consistent
with an in vivo role in membrane-associated processes.
Many of the biological processes involving the cytoskeleton
depend on the coordination of both actin filaments and microtubules
at the plant cell cortex (Fu et al., 2005; Crowell et al., 2009). Preprophase band formation, epidermal cell interdigitation and
cytoskeletal polarisation in response to pathogen attack all require
local antagonism or cooperation between F-actin and microtubules.
Signalling networks orchestrating this coordination are beginning
to be characterised (Fu et al., 2005). We have identified a new plantspecific domain within the group I formin AtFH4 that is capable
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of associating with the microtubule cytoskeleton in vivo. In vitro,
this domain binds directly to microtubules. Full-length AtFH4-GFP
accumulates at the endoplasmic reticulum (ER) and causes
microtubule co-alignment. Plant group I formin AtFH4 therefore
has the potential to associate microtubule and actin arrays with lipid
membranes.
Results and Discussion
Journal of Cell Science
AtFH4 associates with microtubules in vivo
To identify cytoplasmic interaction partners of formin AtFH4, the
fusion construct AtFH4D1 was assembled, containing GFP coupled
to the N terminus of the cytoplasmic section of AtFH4 (Fig. 1A).
AtFH4 and its closest homologues, AtFH7 and AtFH8, constitute
the group Ie subfamily of Arabidopsis formins (Cvrcková et al.,
2004). Members of this clade share close sequence homology within
the FH2 domain, and significant sequence similarity in a region of
unknown function located between the transmembrane domain and
the FH1 domain (Fig. 1A). We have named this region the group
Ie (GOE) domain. The GOE domain corresponds to the
automatically generated ProDom (Bru et al., 2005) domains
PD038281 and PD224441, which are mutually related. The GFPAtFH4D1 construct retains the FH1-FH2 region of AtFH4 and
encompasses the 138-residue GOE domain.
GFP-AtFH4D1 was transiently transformed into leaves of
Nicotiana benthamiana. Observation of transformed leaves three
days after infiltration revealed the organisation of GFP-AtFH4D1
into a filamentous system resembling the cortical microtubule
network (Fig. 1B). To label microtubules in vivo, we used the kinesin
motor domain (KMD) of plant kinesin-7 fused to RFP (KMD-RFP).
Leaves co-infiltrated with GFP-AtFH4D1 and KMD-RFP showed
colocalisation of the GFP-AtFH4D1 and KMD-RFP networks (Fig.
1B). The identity of the GFP-AtFH4D1 filament system was probed
using drugs that target the cytoskeleton. Tissue transformed with
GFP-AtFH4D1 and incubated with 10 mM of the actindepolymerising drug latrunculin B retained an intact network
resembling control treatments, despite developing a disorganised
ER and cytoplasm (Fig. 1C). Equivalent treatments with the plant
microtubule-disrupting agent amiprophosmethyl (APM) caused
severe reduction in the number and length of filaments, progressing
to the complete loss of GFP-AtFH4D1 structures. These data,
together with the observed colocalisation of KMD-RFP, suggest
that GFP-AtFH4D1 associates with microtubules. To confirm that
the localisation of AtFH4D1 was not an artefact of the N.
benthamiana-Agrobacterium tumeficians transformation system,
mature A. thaliana leaves were transformed with GFP-AtFH4D1
using a particle bombardment method, also revealing a microtubule
network (supplementary material Fig. S1). The identity of the
cytoskeletal network labelled by AtFH4D1 was further confirmed
by imaging dynamic AtFH4D1-decorated filaments (Fig. 1D;
supplementary material Movie 1). The elongation rate of GFPAtFH4D1 is approximately 0.05 mm per second, similar to rates of
microtubule elongation measured in Arabidopsis of approximately
0.1 mm per second (e.g. Kawamura and Wasteneys, 2008; Yao et
al., 2008).
A domain for microtubule association is located within the
GOE region
To identify the domain responsible for microtubule association, a
further three deletion constructs of AtFH4 fused to GFP were
transformed into leaves (Fig. 2A). AtFH4D2 contains the FH1 and
FH2 domains, AtFH4D3 encompasses the FH2 domain only, and
Fig. 1. Formin AtFH4 interacts with plant microtubules in vivo.
(A) AtFH4 contains a secretory signal peptide (SP), a transmembrane domain
(TM), an FH1 domain (FH1) and an FH2 domain (FH2). Additionally, AtFH4
shares a novel 138-residue domain with other members of the group Ie
subfamily of formins (labelled GOE). In construct GFP-AtFH4D1, the N
terminus of AtFH4, including the transmembrane domain, has been replaced
with GFP. (B) Imaging of leaf epidermal cells transformed with GFPAtFH4D1 identified a dynamic filamentous network resembling the
microtubule cytoskeleton (left panel). Weak accumulation of GFP-AtFH4D1
within the cytoplasm and ER was also visible. Plants were co-labelled with
KMD-RFP (centre panel), a plant microtubule-binding kinesin motor domain.
KMD-RFP consistently colocalised with GFP-AtFH4D1 (right panel). (C) Leaf
tissue expressing GFP-AtFH4D1 was treated with the actin-depolymerising
drug latrunculin B (centre panel) or the microtubule-disrupting drug APM
(right panel). Latrunculin B treatment disrupted the organisation of the
cytoplasm, but did not affect the filamentous distribution of GFP-AtFH4D1.
By contrast, APM treatment dramatically shortened or eliminated GFPAtFH4D1 filaments (right panel). (D) Time series of leaf cells transformed
with CFP-AtFH4D1 and GFP-Lifeact. Microtubules decorated with CFPAtFH4D1 remain dynamic. The microtubule end highlighted by the arrow is
growing at a rate of 2.7 mm per minute (time points are displayed in the bottom
left corner of each panel).
AtFH4D4 contains the GOE and FH1 domains. Besides AtFH4D1,
only AtFH4D4 associates with microtubules (as summarised in Fig.
2A). AtFH4D1 and AtFH4D4 contain both the GOE domain and
Microtubule- and actin-binding plant formin
the FH1 domain, but the isolated FH1 domain of AtFH4D2 was
not sufficient for microtubule association. A further deletion
construct containing only the GOE domain, AtFH4D5, was
transformed into plants to test the hypothesis that the GOE region
encompasses the microtubule-association domain. AtFH4D5
localised to microtubules, as shown by a co-transformation with
KMD-RFP (Fig. 2B); however, the ratio of cytoplasmic to
microtubule association was noticeably higher for AtFH4D5 than
AtFH4D4, suggesting that, although the GOE domain is sufficient
for microtubule association, the neighbouring FH1 region has a
minor influence on microtubule association in vivo.
Journal of Cell Science
The GOE domain binds to microtubules directly in vitro
The microtubule association of AtFH4 in vivo leads to the
hypothesis that AtFH4 can interact directly with microtubules.
Deletion fragments of AtFH4 were expressed and purified from
Escherichia coli. The microtubule-binding capability of AtFH4D4
was compared with that of AtFH4D5 (the GOE domain) and with
a new deletion, AtFH4D6, which contains only the FH1 domain.
The protein fragments were mixed with taxol-stabilised
microtubules and centrifuged to pellet the microtubules and
associated proteins. Fig. 2C shows that both AtFH4D4 and AtFH4D5
sedimented with microtubules, whereas AtFH4D6 remained in the
supernatant. Fragment AtFH4D4 noticeably decreased the
proportion of tubulin remaining in the supernatant (Fig. 2C;
supplementary material Fig. S2). To test its bundling potential,
AtFH4D4 was mixed with taxol-stabilised and Oregon-Greenlabelled microtubules. Increasing concentrations of AtFH4D4 were
correlated with the co-alignment of microtubules (supplementary
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material Fig. S3), suggesting that the presence of AtFH4D4 promotes
microtubule bundling. Together, these data show that the GOE
domain exhibits microtubule-binding activity in vivo and in vitro,
and that the neighbouring FH1 region also influences AtFH4microtubule interactions.
In animals, three formins have been found to associate directly
with microtubules: mDia1-2, Capu and INF1. The FH2 domains
of mDia2 and Capu are responsible for mediating microtubule
interactions in these proteins (Bartolini et al., 2008; Rosales-Nieves
et al., 2006), with mDia1 also containing an N-terminal region in
an analogous position to the AtFH4 GOE domain that is essential
for mitotic spindle association (Kato et al., 2001). INF1 is a formin
with an unusual domain structure, with two novel microtubulebinding motifs in an extended C terminus (Young et al., 2008).
Neither the mDia1 N terminus nor the INF1 C terminus have
primary sequence similarity to the GOE domain. When considering
the evolutionary distance between the formin families of animals
and angiosperm plants (Grunt et al., 2008; Chalkia et al., 2008), as
well as the absence of GOE-related non-plant sequences detectable
in GenBank by BLAST or PSI-BLAST, it seems likely that the
microtubule affinity of the AtFH4 GOE domain originates from
independent convergence towards similar functions rather than the
ancient conservation of a shared function.
Formins of animals associate with the plus ends of microtubules
using intermediates such as CLIP170, APC and EB1 (Lewkowicz
et al., 2008; Wen et al., 2004). Our data do not exclude such
additional interactions in plants; however, we have not isolated any
known plus-end-binding proteins from AtFH4 interactor screens.
In animals, the plus-end-associated interactions show functional
Fig. 2. The GOE domain of AtFH4 contains a newly identified microtubule-interaction domain. (A) AtFH4 was divided into subfragments to identify
microtubule (MT)-binding sites (for residues, see Materials and Methods). The subfragments were fused with GFP (using vector pMDC43) and transformed into
plant cells. GFP-AtFH4D1, GFP-AtFH4D4 and GFP-AtFH4D5 associated with the microtubule cytoskeleton. These constructs all include the GOE domain.
Constructs without the GOE domain did not label microtubules. (B) Images showing leaf epidermal cells transformed with GFP-AtFH4D5 (left panel) and KMDRFP (centre panel). Colocalisation of the two protein fusions (right panel) indicates that the GOE domain of AtFH4 is sufficient to confer the microtubule
association of AtFH4. (C) Protein fragments AtFH4D4, AtFH4D5 and AtFH4D6 were incubated with taxol-stabilised microtubules. AtFH4D5 (containing the GOE
domain) remained associated with the microtubule pellet (P), whereas fragment AtFH4D6 (containing only the FH1 domain) remained in the supernatant (S)
fraction. In both experiments, positive control AtFH4D4 (containing both the GOE and FH1 domains) associated with microtubules.
Journal of Cell Science
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specificity. For example, mDia1 interactions with CLIP170 are only
essential for phagocytosis mediated by CR3 and not for phagocytosis
mediated by FcR (Lewkowicz et al., 2008). For plants, it is possible
that formin isoforms other than AtFH4 use alternative methods to
interact with microtubules, such as plus-end-binding proteins.
together, these data would suggest a dual role for AtFH4, whereby
microtubules act as a scaffold to which the N-terminal GOE domain
of AtFH4 is attached, allowing the C-terminal FH2 domain to freely
regulate the nucleation of actin filaments from a relatively fixed
position
The actin cable network is distinct from AtFH4-decorated
microtubules in vivo
Full-length AtFH4 associates with microtubules in vivo
The potential for the AtFH4D1 fragment to associate with actin
filaments was assessed by coating Ni-NTA sepharose beads with
6⫻His-tagged AtFH4D1. Coated and uncoated beads were exposed
to a polymerisation solution containing 2 mM of rhodamine-labelled
actin. Rhodamine fluorescence accumulated in a growing corona
around the coated beads before reaching a steady state
approximately 10 minutes after initial exposure (Fig. 3A;
supplementary material Movie 2). A growing corona of fluorescence
was not observed to develop around either beads coated with
AtFH4D4, which lacks the FH2 domain (supplementary material
Fig. S4A), or uncoated beads (supplementary material Fig. S4B).
The inclusion of 10 mM latrunculin B in the polymerisation medium
prevented the formation of an AtFH4D1 corona (Fig. 3A;
supplementary material Movie 3), confirming that the fluorescence
was generated by rhodamine-actin polymerisation. Leaves were cotransformed with CFP-AtFH4D1 and GFP fused to the N terminus
of Lifeact, a 17-residue peptide with affinity for G-actin and Factin (Riedl et al., 2008). GFP-Lifeact revealed a network of Factin cables distinct from the network of CFP-AtFH4D1 (Figs
2D,3B), showing that, under these in vivo conditions, the actininteraction domains of AtFH4D1 do not serve to decorate F-actin
arrays. Similar behaviour has been reported for Arabidopsis formin
AtFH1. This formin in vitro nucleates actin filaments and has sidebinding activity (Michelot et al., 2005; Michelot et al., 2006), but
an equivalent AtFH1 fragment expressed transiently in pollen tubes
does not decorate actin filaments (Cheung and Wu, 2004). Taken
Fig. 3. AtFH4 affects actin polymerisation in vitro. (A) AtFH4 interacts
with actin filaments in vitro. Rhodamine-labelled actin was polymerised in the
presence of AtFH4D1-coated sepharose beads. Images of bead surfaces were
captured during the course of actin polymerisation. The far right panel shows a
control experiment in the presence of 10 mM latrunculin B. Scale bar: 10 mm.
(B) CFP-AtFH4D1 (left panel) was co-transformed with the F-actin marker
GFP-Lifeact (centre panel). Actin cables labelled by GFP-Lifeact were not
labelled by CFP-AtFH4D1, suggesting that the two constructs labelled nonoverlapping networks (right panel).
AtFH4 contains a secretory signal sequence and a single
transmembrane domain within the N terminus. These domains are
absent from the AtFH4D1 deletion, but are adjacent to the GOE
microtubule-binding domain in the full-length protein (Fig. 1A).
The influence of these domains on microtubule interaction was
assessed by expressing full-length AtFH4-GFP in leaf epidermal
cells. AtFH4-GFP was found to align with microtubules, but was
simultaneously associated with an unidentified compartment (Fig.
4A; supplementary material Fig. S5). The coexpression of HDELCFP (a fusion protein retained by the ER) showed that AtFH4-GFP
was localised to the ER (Fig. 4B). The organisation of the HDELCFP-labelled ER is aberrant in the presence of overexpressed
AtFH4-GFP (Fig. 4B). Epidermal cells expressing GFP-AtFH4D1,
which lacks the transmembrane domain, do not show ERmicrotubule co-alignment (Fig. 4B, panel ii; supplementary material
Fig. S6A). This contrasts with cells expressing AtFH4-GFP, in
which ER networks adopt the configuration of the microtubule
cytoskeleton (Fig. 4B panel vii; supplementary material Fig. S6B).
ER tubules can be observed bridging co-aligned sections of the ER
(supplementary material Fig. S7). The strong ER-microtubule coalignment induced by AtFH4-GFP suggests that full-length AtFH4
can associate membranes and microtubules.
In animal cells, interactions with the microtubule cytoskeleton
drive rearrangements of the ER. Kinesin motor proteins and plusend-binding proteins both contribute to this process (Bola and Allan,
2009). Complementing these interactions are proteins such as
CLIMP-63 and VAP-B, which provide static links between
microtubules and the ER. Both CLIMP-63 and VAP-B are type II
membrane proteins integrated into the ER membrane; the cytosolic
domain of CLIMP-63 binds microtubules directly, but the
mechanism for VAP-B remains unclear (Amarilio et al., 2005).
Overexpression of both these classes of proteins causes ER
rearrangement and ER-microtubule co-alignment (Amarilio et al.,
2005; Vedrenne et al., 2005). To date, formins have not been
identified as contributing to these ER-microtubule interactions, but
mammalian formin INF2 was recently found to associate with the
ER periphery of Swiss 3T3 cells (Chhabra et al., 2009).
Constitutively active INF2 mutants caused actin rearrangements that
were detrimental to the ER organisation, but the contribution of
wild-type INF2 to ER function remains unknown. In plants, it has
been thought that actin plays the major role in ER organisation and
motility (Boevink et al., 1998), although during cell division the
ER uses microtubules of the spindle and phragmoplast as an
organisational template (Hepler and Jackson, 1968; Gupton et al.,
2006). In characean algae, the alignment of the ER at the cell cortex
during interphase is reliant on the cortical microtubule array during
periods of cell elongation (Foissner et al., 2009).
Although AtFH4-GFP accumulates at the ER, AtFH4 does not
contain a characterised ER membrane retention motif. Type I
membrane proteins in plants are thought to be delivered by default
to the plasma membrane in the absence of other targeting signals
presented to the cytoplasm or encoded within transmembrane
domains (Brandizzi et al., 2002). Membrane-integrated plant
formins AtFH1 and AtFH6 have been shown to be trafficked to the
Microtubule- and actin-binding plant formin
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Journal of Cell Science
Fig. 4. AtFH4-GFP co-aligns microtubules and ER.
(A) Full-length AtFH4 fused to GFP (green)
colocalises simultaneously with the microtubule
cytoskeleton (labelled with KMD-RFP; red) and a
globular compartment. (B) Coexpression of AtFH4GFP, KMD-RFP and the ER marker HDEL-CFP within
cells identified co-alignment between AtFH4-GFP, the
microtubule cytoskeleton and the ER. Panels i, ii, iii
and iv show cells expressing the control construct
GFP-AtFH4D1, which contains only the cytosolic
domains of AtFH4. Panels v, vi, vii and viii show cells
expressing full-length AtFH4-GFP. Panels i and v
compare AtFH4-GFP localisation; panels ii and vi
compare ER organisation; panels iii and vii compare
microtubule organisation, whereas panels iv and viii
show merged images of all panels. (C) A model
depicting a putative role for AtFH4 at the plant
membrane-cytoskeletal interface. Formins are depicted
in contact with the barbed end of filaments, but the
non-processive nature of AtFH1 and the ability of
AtFH1 to bind to the flanks of filaments (Michelot et
al., 2006) might suggest an alternative F-actin
arrangement for AtFH4. Actin filament anchoring
could theoretically occur in parallel to microtubule
association, or microtubule and F-actin binding could
be mutually exclusive.
plasma membrane (Cheung and Wu, 2004; Favery et al., 2004).
Although AtFH4 accumulates within the ER of transiently
transformed N. benthamiana epidermal cells, it is plausible that
AtFH4 can also be trafficked to the plasma membrane, because,
for example, the passage of some transiently expressed integral
membrane proteins is dependent on the coexpression of specific
cofactors (Ribeiro et al., 2009). The destination for AtFH4 along
the default secretory pathway could also be regulated by
developmental or environmental factors. Immunofluorescence
imaging of sections of Arabidopsis mesophyll labelled with antiAtFH4 showed that endogenous AtFH4 in this tissue is localised
to the cell cortex in proximity to the plasma membrane (Deeks et
al., 2005); however, the transverse mesophyll tissue sections did
not permit the observation of filamentous or reticulate structures.
The arrangement of the cytoskeleton immediately adjacent to the
plasma membrane is essential for the development of the cell wall.
Membrane-integrated cellulose synthase complexes use
microtubules to guide the deposition of cellulose microfibrils
(Paredez et al., 2006). Moreover, coordination with the actin
cytoskeleton is thought to be necessary for the correct delivery of
cellulose synthase particles to sites of cell-wall assembly (Wightman
and Turner, 2008; Crowell et al., 2009). A recent study using highresolution scanning electron microscopy has confirmed the existence
of ordered microtubule arrays in contact with the plasma membrane,
and resolved physical linkages between microtubules and the
membrane (Barton et al., 2008). The identity of these linkages
remains unknown. Few protein candidates have been proposed, apart
from plant phospholipase D (PLD), a peripheral membrane protein
(Gardiner et al., 2001). Our data show that AtFH4 can co-align
microtubules with membranes, demonstrating the potential for an
additional microtubule-plasma membrane coupling mechanism
(Fig. 4C), whereby the AtFH4 integral membrane protein acts as a
scaffold for cytoskeletal organisation.
AtFH4 is one of a small group of formins known to interact
directly with both the actin and microtubule cytoskeletons, although
the AtFH4 microtubule-binding region is unique to plants. Many
developmental processes and environmental responses of plants
require interplay between actin filaments and microtubules (e.g. Fu
et al., 2005; Crowell et al., 2009; Wightman and Turner, 2008).
This requirement for coordination is perhaps reflected in PLD,
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because it was shown that, in addition to binding microtubules, PLD
can interact directly with F-actin (Kusner et al., 2003) and the
phosphatidic acid produced by PLD activity can also bind actincapping protein and thus stimulate F-actin polymerization (Huang
et al., 2006). Our data and those of others show that the FH1-FH2
region of group I formins can nucleate and anchor actin filaments.
Integration of actin and microtubule interactions into one protein
provides an effective tool for enabling local cross-talk between the
two cytoskeletal systems.
Materials and Methods
Journal of Cell Science
Tagged protein expression and live cell imaging
The following fragments of AtFH4 were amplified and recombined into Gateway
entry vector pDONR207: AtFH4D1 (codons 102 to 763), AtFH4D2 (codons 221763), AtFH4D3 (codons 302-763), AtFH4D4 (codons 102-338), AtFH4D5 (codons
102-237) and AtFH4D6 (codons 221-302). For in vivo analysis, LR reaction mix was
used to transfer inserts to N terminus fusion GFP vector pMDC43 (Curtis and
Grossniklaus, 2003) and to CFP fusion vector pH7WGC2 (Karimi et al., 2002).
Construct KMD-RFP encodes 400 residues of the N terminus of N. benthamiana
A90 (see supplementary material Fig. S8), a member of the kinesin-7 (or CENP-E)
family. The KMD fragment was isolated by screening random N. benthamiana cDNA
fusions to GFP (Escobar et al., 2003). The KMD fragment was cloned into a modified
version of pMDC83 (where GFP was replaced by RFP) to form the KMD-RFP fusion.
The Lifeact actin-binding domain (Riedl et al., 2008) was amplified and cloned into
Gateway vector pDONR207 (Invitrogen) before being recombined into destination
vector pMDC43 to create a fusion to GFP at the N terminus of Lifeact. HDEL-CFP
was as described by Nelson et al. (Nelson et al., 2007). Constructs for plant transfection
were transformed in A. tumefaciens strain GV3101. N. benthamiana were infiltrated
as described previously (Smertenko et al., 2008) and plants were imaged using a
Zeiss LSM510 scanning confocal microscope 3 days after infiltration. Biolistic
transformations were carried out with the Helios gene gun (Bio-Rad Laboratories)
with 0.6 mm gold beads coated with DNA according to manufacturer’s instructions,
using 0.05 mg/ml PVP and 0.6 mg DNA per shot. Approximately 5-week-old A. thaliana
leaves were placed on an agar plate, covered with nylon mesh and bombarded at 250
psi. Fluorescence was observed 12-18 hours after bombardment. Drug stocks were
the following: 10 mM latrunculin B (Molecular Probes) in DMSO, 100 mM APM
(Sigma) in ethanol. Sectors of transformed leaf (approximately 25 mm2) were
incubated in latrunculin B, APM or control treatments containing equivalent volumes
of ethanol or DMSO for 1.5 hours prior to imaging.
Protein expression and microtubule-binding assay
For protein expression, AtFH4 fragments were recombined into vector pGAT4
(Ketelaar et al., 2004), resulting in the fusion of a 6⫻His tag to the N terminus of
the respective fragments. Constructs were expressed in E. coli strain BL21 DE3 pLysS
Rosetta 2 and purified using Ni-NTA resin (Qiagen) according to manufacturer’s
guidelines. For the microtubule-binding assay, purified recombinant proteins were
dialysed against PEM buffer (50 mM PIPES pH 6.8, 1 mM EGTA, 1 mM MgCl2,
20% glycerol, 50 mM NaCl, 2 mM DTT). Microtubules were polymerised from
purified porcine brain tubulin at 30°C for 30 minutes in the presence of 1 mM GTP
and 100 mM taxol. 4 mM of polymerised tubulin was co-incubated for 10 minutes in
PEM buffer with approximately 1 mM of recombinant protein before centrifugation
at 230,000g for 10 minutes. Pellet and supernatant fractions were mixed immediately
after spinning with SDS-PAGE loading buffer. For microtubule-binding assays,
Oregon-Green-labelled tubulin (Molecular Probes) was mixed with unlabelled tubulin
at a ratio of 1:5. Tubulin was polymerised as described above and microtubules
incorporating Oregon Green tubulin were sedimented, then resuspended in
polymerisation buffer (10 mM imidazole pH 7.0, 50 mM KCl, 1 mM MgCl2, 2 mM
EGTA, 1 mM DTT, 0.2 mM ATP, 10 mM taxol). 1 mM of polymerised tubulin was
co-incubated briefly with recombinant protein and mounted on a slide for imaging
using a Zeiss LSM510 scanning confocal microscope.
Actin polymerisation in the presence of formin-coated beads
Formin constructs were expressed in E. coli as described above. Proteins were purified
using Ni-NTA beads (Qiagen) according to manufacturer’s instructions, but with the
omission of the elution procedure. The Ni-NTA resin was equilibrated with
polymerisation buffer (10 mM imidazole pH 7.0, 50 mM KCl, 1 mM MgCl2, 2 mM
EGTA, 1 mM DTT, 0.2 mM ATP) and diluted to a concentration of approximately
10 beads per microlitre. 1 ml of bead suspension was mixed with 2 mM of rhodaminelabelled actin monomer (Cytoskeleton) and polymerisation buffer to a total volume
of 10 ml. After gentle mixing, the reaction was transferred to a microscope slide. The
slide was coated with a parafilm spacer with a cut square (5 mm sides) into which
the bead suspension was deposited. A cover slip 10 mm⫻10 mm was used to seal
the chamber. Images were taken every 100 seconds using a Zeiss LSM510 scanning
confocal microscope (excitation 543 nm, 40⫻ objective).
This work was supported by the BBSRC and by Ministry of
Education of the Czech Republic (MSM0021620858 and LC06004)
and Grant Agency of the Czech Republic (P305/10/0433) grants to M.F.,
F.C. and V.Z. HDEL-CFP was constructed by A. Nebenführ (University
of Tennessee, Knoxville, TN) and distributed by Arabidopsis Biological
Resource Center (ABRC).
Supplementary material available online at
http://jcs.biologists.org/cgi/content/full/123/8/1209/DC1
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