The Plant Journal (2011) 66, 354–365
doi: 10.1111/j.1365-313X.2011.04497.x
Building bridges: formin1 of Arabidopsis forms a connection
between the cell wall and the actin cytoskeleton
Alexandre Martinière1, Philippe Gayral2, Chris Hawes1 and John Runions1,*
Department of Life Sciences, Oxford Brookes University, Gipsy Lane, Headington, Oxford OX3 0BP, UK, and
2
Institut des Sciences de l’Evolution, CNRS UMR 5554, Université Montpellier 2, Place E. Bataillon, 34095 Montpellier, France
1
Received 23 November 2010; revised 6 January 2011; accepted 7 January 2011; published online 28 February 2011.
*
For correspondence (fax +44 1865 483242; e-mail [email protected]).
SUMMARY
Actin microfilament (MF) organization and remodelling is critical to cell function. The formin family of actin
binding proteins are involved in nucleating MFs in Arabidopsis thaliana. They all contain formin homology
domains in the intracellular, C-terminal half of the protein that interacts with MFs. Formins in class I are usually
targeted to the plasma membrane and this is true of Formin1 (AtFH1) of A. thaliana. In this study, we have
investigated the extracellular domain of AtFH1 and we demonstrate that AtFH1 forms a bridge from the actin
cytoskeleton, across the plasma membrane and is anchored within the cell wall. AtFH1 has a large,
extracellular domain that is maintained by purifying selection and that contains four conserved regions, one
of which is responsible for immobilising the protein. Protein anchoring within the cell wall is reduced in
constructs that express truncations of the extracellular domain and in experiments in protoplasts without
primary cell walls. The 18 amino acid proline-rich extracellular domain that is responsible for AtFH1 anchoring
has homology with cell-wall extensins. We also have shown that anchoring of AtFH1 in the cell wall promotes
actin bundling within the cell and that overexpression of AtFH1 has an inhibitory effect on organelle actindependant dynamics. Thus, the AtFH1 bridge provides stable anchor points for the actin cytoskeleton and is
probably a crucial component of the signalling response and actin-remodelling mechanisms.
Keywords: formin, actin, cell wall, cytoskeleton, microtubule.
INTRODUCTION
Actin microfilaments (MF or F-actin) are a major component
of the cytoskeleton and are essential for many cellular processes including division, expansion, differentiation, and
are components of cellular response mechanisms. Within
plant cells, most of the dynamic processes, e.g. organelle
movement, vesicle trafficking and cytoplasmic streaming
occur via interaction with MFs. Plant MFs themselves are
extremely dynamic structures with a rate of polymerization
of up to 1.7 lm sec)1 (Staiger et al., 2009; Smertenko et al.,
2010). To make new filaments, actin nucleation proteins are
required.
One of the main gene families involved in this function is
formin. The formin gene family is very diverse, with more
than 20 genes in A. thaliana (Deeks et al., 2002; Cvrckova
et al., 2004b; Blanchoin and Staiger, 2010). They all share a
formin homology domain (FH2) in the C-terminal half of the
protein which is involved in nucleating MFs. In monocotyledonous and dicotyledonous plants, the formin gene family
is organized into two classes. Most class I formins have a
membrane targeting domain in the N-terminal region of
354
the protein. This domain is composed of a signal peptide
followed by a transmembrane domain and appears similar
to that of type I membrane proteins in general. Formins in
class I are in most cases targeted to the plasma membrane
(PM) as has been demonstrated for AtFH1, AtFH5 and AtFH6
by GFP fusions (Cheung and Wu, 2004; Favery et al., 2004;
Van Damme et al., 2004), and by immuno-localization for
AtFH4 (Deeks et al., 2005). Class II formins (AtFH12–AtFH21)
are characterized by a phosphatase tensin (PTEN)-like
domain at the N-terminus. This domain is hypothesized
to mediate membrane targeting and probably determines
subcellular localization of the protein (Cvrckova et al., 2004a;
Vidali et al., 2009). Vidali et al. (2009) have shown by RNA
silencing that class II formins are involved in polarized
growth in moss (Physcomitrella pattens). In vascular plants,
AtFH14 has recently been shown to be crucial for cell
division and is able to induce co-alignment between MFs
and microtubules (Li et al., 2010).
The formin homology domains (FH1 and FH2) of the
Arabidopsis class I formin proteins AtFH1, AtFH3, AtFH4,
ª 2011 The Authors
The Plant Journal ª 2011 Blackwell Publishing Ltd
AtFH1 links the cell wall and actin cytoskeleton 355
AtFH5 and AtFH8, are able to nucleate actin filaments in vitro
(Deeks et al., 2005; Michelot et al., 2005; Yi et al., 2005; Ye
et al., 2009). The MF nucleation mechanism of AtFH1 has
been described (Michelot et al., 2006). AtFH1 is a nonprocessive formin, it becomes localized to the side of the
MF after a nucleation event and so, by repeated nucleation,
AtFH1 causes formation of bundles of MFs during in vitro
assays (Michelot et al., 2006). In vivo, overexpression of
AtFH1, AtFH3 and AtFH4 induces formation of an excess of
MFs at the cell cortex. Overexpression of AtFH8 induces
aberrant root hair development (Deeks et al., 2005; Yi et al.,
2005), and overexpression of AtFH1 and AtFH3 has an effect
on pollen tube growth (Cheung and Wu, 2004; Ye et al.,
2009). Down-regulation of AtFH3 by RNAi results in a
decrease in the numbers of MFs and a reduction of
cytoplasmic streaming in pollen tubes (Ye et al., 2009). More
specific functions have been described for some formin
genes. AtFH5 is localized at the cell plate and its loss of
function results in impaired cytokinesis (Ingouff et al., 2005).
More recently, AtFH5 has shown to be involved in pollen
tube polarization (Cheung et al., 2010). Finally, RNA
expression data show that AtFH1, AtFH6 and AtFH10 are
up-regulated during nematode infection which predicts that
Class I formins are involved in response to pathogen attack
(Favery et al., 2004) and generally have a signalling function
in environmental response pathways.
In animal cells, numerous studies describe protein complexes involved in interaction between the extracellular
space of the cell and the actin cytoskeleton. One of the best
described is the integrin/actin interaction, which is involved
in mechanisms such as focal adhesion. There is no reason
to suspect that interaction between cytoskeletal systems
and the extracellular matrix – mediated by proteins with
extracellular domains – does not occur in plant cells (Baluska
et al., 2003). In fact, a rapid cytoskeleton, endoplasmic
reticulum and Golgi apparatus reorganization occurs after
extracellular stimulation (Hardham, 2007). Similar observations have been made during pathogen infection in which
fungus attack induces a strong and fast reorganization of the
actin cytoskeleton (Koh et al., 2005; Hardham, 2007).
According to protein topology prediction, class I formins
have an extracellular domain. In the case of AtFH1, it is
comprised of nearly 100 amino acids. For this reason, type I
formins are good candidates for linking between the outside
of cells and the actin cytoskeleton, and might be involved
in signal transduction through the PM (Deeks et al., 2002).
Alignment of formin 1 proteins from nine dicotyledonous
and monocotyledonous plants enabled us to find a conserved region which suggests conserved functions for these
domains (Figure S1). This study is focused on the potential
role of the membrane targeting signal and extracellular
domain in the function of formin1 of Arabidopsis (AtFH1).
By resolving the topology, we first show that AtFH1 has an
extracellular domain with subdomains conserved by purify-
ing selection. Then, we show several proofs of interaction
between this region and the extracellular matrix of the cell.
Finally, we demonstrate a correlation between anchoring of
AtFH1 within the extracellular matrix and its actin polymerization activity.
RESULTS
The extracellular domain of Arabidopsis formin 1 evolved
under strong purifying selection
The amino acid alignment of a set of plant formin1 proteins
clearly reveals a conserved region in the potential extra
cellular domain (Figure S1). Estimation of the nonsynonymous to synonymous substitution ratio (x = dN/dS ratio)
was used to quantify the selective pressure acting on the
AtFH1 gene and orthologous sequences in plants. Genes
or domains with no function evolve neutrally (x = 1), while
genes evolving under selection have x „ 1. Positive selection (x > 1) is observed when amino acid replacement is
favored by natural selection. Purifying selection (x < 1) is
observed when amino acid replacement is counter-selected.
Evolution under strong purifying selection pressure tends to
have low values of x, typically a few percent.
The x estimation of the putative extracellular domain
studied here (amino acids 1–107) was 0.070, that of the
intermediate protein region (amino acids 108–520) was
0.200, that of the 3¢ part of the gene which included the
FH1 and FH2 conserved domains (amino acids 521–1051)
was 0.078. This result indicates that the putative extracellular
domain evolved under strong purifying selection of the
same order of magnitude as the intracellular region which
contains the conserved FH1 and FH2 actin-interacting
domains and demonstrates the evolutionary importance of
the extracellular domain for AtFH1 function in plants.
Arabidopsis formin1 is targeted to the plasma membrane
and is excluded by cortical microtubules
We decided to confirm the sub-cellular localization of AtFH1
by fluorescent protein fusion. AtFH1 cDNA was cloned as a
C-terminal fusion with fluorescent proteins (GFP, YFP and
RFP) under control of the 35S promoter and expressed
transiently in tobacco (Nicotiana tabacum) leaf tissue
(Figure 1a). This experimental system was selected as it
was already known that AtFH1 has a dramatic effect on MF
organization and it has proven difficult to generate stably
transformed Arabidopsis plants (Banno and Chua, 2000;
Cheung and Wu, 2004). AtFH1–GFP labelled very clearly the
outline of leaf epidermal cells, produced a sheet of fluorescence at the cell surface (Figure 1b) and co-localized with a
known plasma membrane protein, PIP2;1–CFP (Figure S2)
thus demonstrating that AtFH1 is targeted to the PM. We
also tested whether AtFH1 is targeted to the PM via
the secretory pathway. Inhibition of ER/Golgi transport with
brefeldin A or with Sar1-GTP locked (daSilva et al., 2004)
ª 2011 The Authors
The Plant Journal ª 2011 Blackwell Publishing Ltd, The Plant Journal, (2011), 66, 354–365
356 Alexandre Martinière et al.
(a)
(b)
(c)
Figure 1. AtFH1 is localized to the plasma membrane and marks it in a non-homogeneous way.
(a) Schematic drawing of the different constructs used in the study. FORMIN1 of Arabidopsis thaliana (AtFH1) contains a transmembrane domain (TM) and formin
homology domains (FH1 and FH2) within the intracellular part of the protein (not to scale), and a series of conserved domains (Domains A–D) within the extracellular
region. SP, signal peptide; FP, green, yellow, or red fluorescent protein.
(b) Cortical section of AtFH1–GFP expressed in tobacco leaf epidermal cells. Arrow points to one of the unlabeled structures that resemble microtubules in the
fluorescent membrane.
(c) The stripe pattern of AtFH1–YFP labelling (magenta) and the stripes visible when microtubules are labelled with CFP–TuA6 (green) are exactly coincidental when
overlain. Scale bars = 10 lm.
induced an accumulation of AtFH1 in the ER (Figure S3).
AtFH1 is therefore co-translated inserted into the ER membrane and then targeted to the PM. Curiously, AtFH1–GFP
does not have an homogeneous distribution at the PM
as unlabeled stripes were visible (Figure 1b, arrow). The
pattern of non-fluorescent stripes in the PM resembled
microtubules (MT) in shape and quantity. To verify this
hypothesis, AtFH1–YFP was co-expressed with the MT
labelling construct CFP–TuA6. Microtubules were present
exactly underlying the non-fluorescent pattern of AtFH1–YFP
(Figure 1c). Moreover, when the AtFH1–GFP tobacco leaves
were incubated with 20 lM oryzalin, a microtubule-depolymerizing drug, the non-fluorescent stripes disappeared
(Figure S4). Actin MFs are not involved in formation of this
pattern as MF depolymerization with latrunculin B did not
abolish the stripes (Figure S4).
AtFH1 has a large extracellular domain which interacts
with the cell wall
The predicted intracellular part of the protein was removed
by cloning the first 140 amino acids of the N-terminus of the
protein as a fusion with GFP (AtFH1-dFH–GFP) (Figure 1a).
AtFH1-dFH–GFP-expressing cells were plasmolyzed and
GFP fluorescence was observed in the withdrawn PM and
in Hechtian strands but not in the apoplastic space
(Figure 2a, *). AtFH1-dFH–GFP labelled the PM and proved
that the first 140 amino acids are sufficient for PM
targeting and protein anchoring in the PM. We then
removed the predicted transmembrane domain by fusing
only the N-terminal 107 amino acids of AtFH1 with RFP
(AtFH1-107–RFP). As predicted, the truncated protein was
not able to remain fixed within the PM and was mainly
localized instead at the cell wall (Figure 2b). We conclude
that AtFH1 has an extracellular domain at its N-terminus
of around 100 amino acids. Interestingly, in plasmolyzed
tissue, AtFH1-107–RFP cells show labelling of the cell wall
(as opposed to the apoplastic space created between cell
wall and PM by plasmolysis) which was never observed
when cells expressed the secreted form of RFP (compare
Figure 2b,c, arrows). This finding suggests a potential
interaction between the extracellular domain of AtFH1 and
the cell wall.
ª 2011 The Authors
The Plant Journal ª 2011 Blackwell Publishing Ltd, The Plant Journal, (2011), 66, 354–365
AtFH1 links the cell wall and actin cytoskeleton 357
(a)
(b)
(c)
Figure 2. AtFH1 interaction with the cell wall in plasmolysis experiments.
(a–c) Cells expressing different fluorescent protein labelled constructs were incubated with 0.5 M mannitol to induce plasmolysis.
(a) The formin deletion construct with intracellular domain removed (AtFH1-dFH–GFP) labels the plasma membrane and Hectian strands within the apoplastic space
created by plasmolysis (*). The cell wall (arrow) is unlabelled.
(b) In cells expressing only the N-terminal 107 amino acids of AtFH1 (AtFH1-107–RFP) which does not include the transmembrane domain, the protein remains for
the most part within the cell wall (arrow) and marks the apoplastic space (*) and plasma membrane weakly.
(c) Secreted RFP (Sec–RFP) filled the apoplastic space (*) while fluorescence was not apparent within the cell wall (arrow). Scale bars = 10 lm.
AtFH1 is immobilized within the PM by a cell wall/plasma
membrane connection
To test the lateral mobility of AtFH1 as a measure of its
binding to other cellular components, we performed Fluorescence Recovery After Photobleaching (FRAP) experiments over a short time scale (1 or 2 mim) to exclude any
side effects that might result from endo-, or exocytotic
removal or insertion of protein from the membrane (KleineVehn et al., 2008; Men et al., 2008). AtFH1–GFP produced a
very unusual FRAP recovery curve. Approximately 20% of
initial fluorescence was recovered in the bleached region
within 10–20 sec but then nearly no fluorescence increase
was recorded for the next 100 sec (Figure 3a,c; dashed blue
line). The plateau value of fluorescence intensity recovery
was designated I100s and taken to represent the mobile
fraction of AtFH1–GFP. AtFH1–GFP, therefore, has a surprisingly low mobile fraction (I100s = 20.2 5.1%, Figure 3c
and Table 1). Indeed, we expected from observations of
other plasma-membrane anchored proteins (such as GFPLTI6b in Figure 3b) that AtFH1 would laterally diffuse in the
membrane and have a much higher I100s. One hypothesis to
explain the low mobility of AtFH1–GFP is that it interacts
with other proteins or structures which constrain its mobility. Formins are known to interact with the cell cytoskeleton
(Michelot et al., 2005; Deeks et al., 2010; Li et al., 2010) so
interaction with MF or microtubules might reduce the lateral
mobility of AtFH1. To test this hypothesis, we treated AtFH1–
GFP-expressing cells with latrunculin B and oryzalin to
depolymerise both actin and microtubule networks. These
treatments had no effect on protein mobility (Table 1). Furthermore, the truncated construct AtFH1-dFH–GFP, which
we predict to be unable to interact with cytoplasmic elements, remained largely immobile in FRAP experiments
(Figure 3c; solid blue line and Table 1). The relative immobility of AtFH1 within the PM is not due to an intracellular
interaction so we turned to investigation of its extracellular
domain. We analyzed lateral mobility of AtFH1-dFH–GFP in
cells which were plasmolyzed with various concentrations
of mannitol to separate the plasma membrane from the cell
wall and found that it more than doubled in plasmolyzed
cells compared with untreated cells (Figure 3c; green and
red lines and Table 1). This increase of I100s was not due to
a general effect of osmotic treatment on protein mobility
within the PM as no increase of I100s was observed for PIP2a–
CFP under similar conditions (Table 1). Spatial proximity
between cell wall and PM is required for AtFH1 to remain
largely immobile within the PM.
Cell wall regeneration inhibits the lateral mobility
of AtFH1-dFH
Interaction with the cell wall seems to reduce the lateral
mobility of AtFH1-dFH–GFP. To support these observations,
we recorded protein mobility during cell wall regeneration
of protoplasts. AtFH1-dFH–YFP-expressing protoplasts were
used in FRAP experiments to record lateral mobility of
AtFH1-dFH–YFP at times 0, 24, and 48 h during development
of the new primary cell wall (Figure 3d). Protein lateral
mobility decreased gradually from t0 to t48 (Figure 3e–g)
(Tukey test of I100s, t0 versus t24, P = 0.046; t0 versus t48,
P = 0.001; Table 2). During cell wall regeneration, a concomitant stabilization of AtFH1-dFH–YFP occurs in the PM.
The agent of this stabilization is absent or reduced in newly
made protoplasts and gradually appears during the early
phase of cell-wall deposition.
A conserved subdomain of the proline-serine-rich
extracellular domain is involved in anchoring of AtFH1
to the cell wall
The extracellular domain of AtFH1 has four highly-conserved sub-domains: Domain A residue 23–32, Domain B
residue 37–56, Domain C residue 66–80, and Domain D
residue 83–96 (Figure 1a). Complete removal of all four
domains resulted in a fluorescent construct that remained
ª 2011 The Authors
The Plant Journal ª 2011 Blackwell Publishing Ltd, The Plant Journal, (2011), 66, 354–365
358 Alexandre Martinière et al.
(a)
(c)
(b)
(d)
(e)
(f)
(g)
Figure 3. Lateral mobility of Formin is constrained by the cell wall.
(a, b) Fluorescence Recovery After Photobleaching experiments to compare the lateral mobility of AtFH1–GFP with another plasma membrane marker, GFP-LTI6b.
(a) AtFH1 is very immobile during 80 sec recovery within the plasma membrane due to the cell wall/plasma membrane connection. Fluorescence is slow to recover
when compared with that of (b) GFP-LTI6b. Scale bar = 5 lm.
(c) FRAP curves of AtFH1 and AtFH1-dFH–GFP before and after plasmolysis. Mannitol treatment causes the PM to separate from the cell wall and results in a
significant increase in mobility of AtFH1-dFH–GFP.
(d) Cellulose stained with Calcofluor white M2R (blue). At time 0 no staining is apparent. By 48 h after making protoplasts the cell wall is well developed and stains
strongly. Scale bar = 10 lm.
(e) FRAP curves show decreasing mobility of AtFH1-dFH–YFP during the 48 h of cell wall regeneration in protoplasts as the connection between cell wall and the
extracellular domain of AtFH1 is re-established.
(f, g) Examples of FRAP of AtFH1-dFH protoplasts without (e) and with (f) cell wall. Fluorescence recovery at 110 sec is much less in protoplasts with primary cell wall
(f) Pixels values are rainbow coloured so that high intensity fluorescence is indicated with red. The rectangle shows the bleaching area.
Table 1 Mobility of AtFH1 within the plasma membrane
Construct
Treatment
Mobile fraction
(%)
AtFH1–GFP
AtFH1–GFP
AtFH1-dFH–GFP
AtFH1-dFH–GFP
AtFH1-dFH–GFP
PIP2;1–CFP
PIP2;1–CFP
–
Latrunculin b/oryzalin
–
0.35 M mannitol
0.5 M mannitol
–
0.5 M mannitol
20.2
19.2
21.4
42.0
48.0
15.8
14.2
a
5.1
6.6
3.6
11.7a
13.7a
5.7
5.7
Mannitol treatments compared with no treatment for AtFH1-dFH–
GFP, P < 0.05 Tukey HSD test, n = 10 cells. PIP2a-CFP did not differ in
lateral mobility between control and plasmolyzed cells (n = 9 cells).
Mobile fraction is expressed as mean standard deviation.
in the ER (Figure S5b). Proper targeting to the PM was
achieved when Domain A was included in the deletion
constructs. To determine which of Domains B, C or D are
involved in anchoring of AtFH1 to the cell wall, three
deletion constructs were made from the full length protein.
AtFH1-TR1 in which Domains B, C and D are removed,
AtFH1-TR2 in which Domain B and C are removed, and
AtFH1-TR3 in which only Domain B is removed (Figure 1a).
All three constructs labelled the PM strongly without any
obvious differences compared with the full length protein
(Figure S5a,c–e). FRAP was used to record the lateral
mobility of these different deletion constructs. None of
them had a fluorescence-intensity recovery value that differed from that of the control protein AtFH1–YFP (P = 0.785;
Table 3). Because formins are known to interact with
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The Plant Journal ª 2011 Blackwell Publishing Ltd, The Plant Journal, (2011), 66, 354–365
AtFH1 links the cell wall and actin cytoskeleton 359
Table 2 Lateral mobility of AtFH1-dFH–YFP decreases during cell
wall regeneration
Hours post cell wall digestion
Mobile fraction (%)
0
24
48
29.9 15.9
19.3 5.7*
12.3 6.7*
*P < 0.05 Tukey HSD test, n = 12 cells. Mobile fraction is expressed as
mean standard deviation.
Table 3 Lateral mobility of AtFH1 is affected in extracellular domain
deletion constructs when the cytoskeletons are removed
Mobile fraction
(I100%) (cytoskeleton
present)
AtFH1–YFP
AtFH1-TR1–YFP
AtFH1-TR2–YFP
AtFH1-TR3–YFP
15.2
12.7
13.3
15.3
7.9
4.2
5.9
5.7
Mobile fraction
(%) (cytoskeleton
absent)
15.7
19.6
21.1
17.7
4.6
4.0*
5.8*
2.4
*Tukey HSD test, untreated versus latrunculin B/oryzalin treated,
n = 9 cells. ANOVA Between constructs, P = 0.785; ANOVA Between
treatments (cytoskeleton removal), P = 0.001. Mobile fraction is
expressed as mean standard deviation.
cytoskeletal elements (Michelot et al., 2006; Deeks et al.,
2010), we treated cells with 25 lM latrunculin B and 20 lM
oryzalin to remove MF and microtubules, respectively. In
these conditions, the full length construct and AtFH1-TR3–
YFP did not behave differently between control and treated
cells, but fluorescence-intensity recovery of AtFH1-TR1 and
AtFH1-TR2 increased significantly compared with that in
untreated cells (Tukey HSD test, untreated versus treated,
for AtFH1–YFP, P = 0.889; for AtFH1-TR1–YFP, P = 0.004; for
AtFH1-TR2–YFP, P = 0.008, for AtFH1-TR3–YFP, P = 0.284;
Table 3). The only difference between AtFH1-TR2 and
AtFH1-TR3 is the addition of the 18 amino acids of Domain
C (PFFPLYPSSPPPPSPASF). This sequence is sufficient for
keeping AtFH1 immobilized and contains the motif SPPPP
which is homologous with the extensin glycosylated
domain (Banno and Chua, 2000). Consequently, this domain
appears to be involved in a cell wall/AtFH1 interaction.
AtFH1 overexpression induces an increase in the number
of actin filaments when AtFH1 interacts with the cell wall
Overexpression of AtFH1 in tobacco pollen tubes induces an
increase in the number of actin filaments at the cell cortex
(Cheung and Wu, 2004). To determine if overexpression of
AtFH1-xFP induces any similar effect in tobacco leaf epidermal cells, it was co-expressed with one of the actin
labelling constructs Lifeact–RFP or Lifeact–GFP (Era et al.,
2009; Riedl et al., 2008; Roca et al., 2010). Long, thick bundles of MFs, as would be observed without AtFH1 overexpression, disappeared and a dense meshwork of thin,
mainly parallel MFs was observed (Figure 4a–c). This finding
was also observed with other actin labelling constructs
(Figure S6a,b). To confirm that this effect was not due to the
fluorescent-protein tag fused to AtFH1, we expressed a nontagged AtFH1 with Lifeact–GFP and the same altered MF
pattern was observed (Figure 4d). When the FH domain was
removed from AtFH1, the actin network appeared as in
AtFH1 non-overexpressing cells (Figure 4e). These two
experiments indicate that AtFH1-xFP is functional in our
system. But, in a somewhat unexpected manner, when the
extracellular domain truncation constructs AtFH1-TR1–YFP
and AtFH1-TR2–YFP were co-expressed with Lifeact–RFP,
the MF pattern did not resemble the actin structure
described for AtFH1 overexpressing cells (Figure 4f,g).
Rather, for these deletions (TR1 and TR2), the effect on actin
network morphology was relatively weak with only 31% of
cells for AtFH1-TR1–YFP and 34% of cells for AtFH1-TR2–YFP
having an increase in the number of MFs (n = 100 transformed cells from four independent experiments). In contrast, the Lifeact–RFP pattern in AtFH1-TR3–YFP-expressing
cells was similar to that in cells expressing the full length
protein AtFH1–YFP (Figure 4h). These results show that
presence of Domain C – and its cell-wall anchoring function –
is required to produce the increase in MF number that results
in an actin sheet in the cell cortex. Unfortunately, it is difficult
to quantify precisely the effect of the AtFH1 truncation
constructs on the MF network, especially as we use actin
labelling tags that might also affect actin morphology and
dynamics. For this reason, we decided to follow Golgi body
motility as a reporter of the AtFH1-overexpression effect on
the actin cytoskeleton.
Overexpression of AtFH1 inhibits Golgi body motility
Golgi bodies are highly motile organelles in plant cells, and
this movement is dependent on presence of the actin cytoskeleton and myosin proteins (Brandizzi et al., 2002; Sparkes
et al., 2008). We expect that changes in MF organization
might perturb Golgi motility. Sialyl transferase-mRFP (STmRFP), a Golgi body fluorescent tag, was co-expressed with
our set of AtFH1 constructs. Golgi body motility was
recorded by time lapse imaging. The Meandering Index (MI –
Golgi displacement:track length) was calculated. The value
of MI varies from close to 0 when the Golgi body is immobile
or moves erratically, to close to 1, when a Golgi body follows
a straight trajectory (Runions et al., 2006; Sparkes et al.,
2008). When ST-mRFP is expressed on its own,
MI = 0.41 0.26 (standard deviation) (Figure 5a,h). This
value is consistent with previous studies and, as others
have shown, has a high standard deviation because of the
co-occurrence of moving and non-moving Golgi bodies in
the same cell (Runions et al., 2006; Sparkes et al., 2008).
After removal of the actin cytoskeleton by latrunculin B
treatment, Golgi body motility was significantly reduced
(MI = 0.07 0.07) (Figure 5b,h). This reduction in MI is
ª 2011 The Authors
The Plant Journal ª 2011 Blackwell Publishing Ltd, The Plant Journal, (2011), 66, 354–365
360 Alexandre Martinière et al.
(a)
(b)
(c)
(d)
(e)
(f)
(g)
(h)
Figure 4. Overexpression of AtFH1 induces a re-organization of the actin cytoskeleton.
(a–h) Actin structure labelled with Lifact–GFP (green) or Lifact–RFP (magenta) in presence or absence of different AtFH1 constructs.
(a) Actin cytoskeleton in the absence of AtFH1 overexpression.
(b) A cell co-expressing AtFH1–GFP and Lifeact–RFP (arrow) has a dense mat of actin MFs very different to the cells which surround it that only express Lifact–RFP
(stars).
(c) The same effect was recorded with co-expression of Lifeact–GFP and AtFH1–RFP.
(d) Overexpression of un-tagged AtFH1 resulted in formation of a dense array of MFs similar to that induced by the tagged version of AtFH1.
(e) Overexpression of the AtFH1 construct with no actin-binding intracellular domain (AtFH1-dFH–YFP) had no effect on organization of the actin cytoskeleton.
(f–h) Actin cytoskeleton morphology in cells overexpressing various deletions of the AtFH1 extracellular domain.
(f) Overexpression of the construct in which extracellular subdomains B, C, and D are deleted (AtFH1-TR1–YFP), or (g) the construct in which extracellular
subdomains B and C are deleted (AtFH1-TR2–YFP), had very little apparent effect on actin cytoskeleton morphology.
(h) When subdomain C was added back into the extracellular domain deletion (AtFH1-TR3–YFP) the actin cytoskeleton appeared as if the full length AtFH1 was being
overexpressed [compare with (c–h)]. Scale bars = 10 lm.
because Golgi bodies can no longer move along straight
trajectories when the actin cytoskeleton is depolymerized.
When AtFH1–YFP is co-expressed with ST-mRFP in cells with
intact actin cytoskeletons, Golgi movement is strongly
reduced (MI = 0.20 0.16; Figure 5c,h), but not stopped
as with latrunculin B treatment (Tukey HSD test, ST-mRFP
alone versus ST-mRFP + AtFH1–YFP, P = 0.001). AtFH1
expression, because of its effect on actin network reorganization, inhibits Golgi body motility. Expression of AtFH1dFH–YFP, which does not contain the intracellular FH1 and
FH2 domains that bind actin, had no effect on Golgi body
movement (MI = 0.42 0.26; Figure 5d,h). AtFH1 constructs with various truncations in the conserved extracellular domains all reduced Golgi body movement with
AtFH1-TR3–YFP having the most severe effect and confirming the requirement of cell wall anchoring by extracellular
Domain 3 for producing the observed overexpression
phenotype (Figure 5e–h).
DISCUSSION
Actin microfilaments are extremely dynamic structures
(Staiger et al., 2009). Many physiological processes, e.g.
pathogen response or re-arrangement during cell division
illustrate the plasticity of this network. The constant
re-modelling activity of actin microfilaments (nucleation,
bundling and severing) is mainly driven by actin interacting
proteins (Staiger et al., 2009). Regulation of actin cytoskeleton morphology is crucial and a mechanism for coordinating these re-arrangements in response to signals from
the external medium is necessary. Many different actininteracting proteins have been postulated as the physical
link between cell wall and the actin cytoskeleton (Baluska
et al., 2003) and we can now demonstrate that the plasma
membrane protein Formin1 connects the cell wall to the
actin cytoskeleton.
AtFH1 forms a link between the actin cytoskeleton
and the cell wall
We believed that to organize and coordinate a complex
structure, having spatially stable anchor points is crucial.
For instance, the endoplasmic reticulum has a constantly
changing shape. Recently, stable ER anchoring points were
described (Sparkes et al., 2009). Our observations suggest
that Arabidopsis Formin1 (AtFH1) has a similar function in
ª 2011 The Authors
The Plant Journal ª 2011 Blackwell Publishing Ltd, The Plant Journal, (2011), 66, 354–365
AtFH1 links the cell wall and actin cytoskeleton 361
(a)
(b)
(c)
(d)
(e)
(f)
(g)
(h)
Figure 5. Overexpression of AtFH1 inhibits the mobility of Golgi bodies.
(a–g) Time series data of ST-mRFP-expressing cells was used for tracking Golgi-body mobility. Golgi body paths are indicated with colored lines.
(a) Control cells.
(b) Latrunculin b treated cells. Golgi-body mobility is inhibited when the actin cytoskeleton is depolymerized as can be seen by the reduced path length.
(c–g) ST-mRFP co-expressed with different AtFH1 constructs, (c) the full length protein AtH1–YFP, (d) AtFH1-dFH–YFP, (e) AtFH1-TR1–YFP, (f) AtFH1-TR2–YFP and (g)
AtFH1-TR3–YFP. Golgi bodies are immobilized when either the FH domain or the extracellular domain of AtFH1 is present.
(h) Golgi-body Meandering Index (MI), calculated as the ratio between Golgi displacement and total track length. MI is lower when mobility is inhibited. Removal of
the actin cytoskeleton by treatment with latrunculin B almost completely eliminates Golgi movement. Overexpression of both the full length AtFH1 and the AtFH1
extracellular deletion construct with Domain C present (AtFH1-TR3) significantly reduced Golgi mobility. When the intracellular, actin binding domains of AtFH1
were removed (AtFH1-dFH), or when Domain C of the extracellular domain was absent (AtFH1-TR1 or AtFH1-TR2), Golgi mobility was significantly increased relative
to that in AtFH1 expressing cells. Mean SEM; * mobility of Golgi in presence of AtFH1 compared with other constructs, P < 0.05 t-test; n = 10 cells. Scale
bar = 10 lm.
that it forms a bridge across the plasma membrane between
actin filaments and the extracellular matrix (Figure 6). AtFH1
and most members of the class I plant formins have been
predicted to be type I integral membrane proteins (Cvrckova,
2000). In this study, we have confirmed that AtFH1 is a
plasma membrane protein and have shown that it has a
conserved N-terminal domain, which is localized within the
extracellular space.
Four experimental techniques have been used to demonstrate the presence of a link between the extracellular part
of AtFH1 and the extra-cellular matrix (ECM). Firstly, when
plasma membrane/cell wall proximity was disrupted by
plasmolysis, AtFH1 remained in the PM and its lateral
mobility within the plasma membrane increased. This
finding suggests that a physical link with a protein-anchoring function between AtFH1 and the ECM was disrupted by
plasmolysis. Secondly, the PM lateral mobility of AtFH1-dFH
was high in protoplasts and steadily decreased during 48 h
of cell-wall regeneration. Thirdly, removal of the majority of
the extracellular N-terminus of full length AtFH1 induced an
increase in the protein’s lateral mobility when the cytoskeleton was removed. Finally, removal of the transmembrane
domain (AtFH1-107–RFP) resulted in labelling of the cell wall
specifically in plasmolyzed cells. As a consequence of these
observations, we can say that AtFH1 forms a relatively
immobile link from the cell wall through the PM and acts as
an anchor point for the actin cytoskeleton. This model can be
related to the biochemical model of AtFH1’s actin-nucleating
mechanism. Formin1 is a non-processive formin which
means that it does not move with the growing barbed end
of a microfilament (Michelot et al., 2006). Rather, the barbed
end is spatially motile and grows away from the stable MF
nucleation site of formin attachment.
The extracellular region of AtFH1 has four highly conserved subdomains and the cell-wall anchoring role of
this region could explain why the amino acid sequences of
these subdomains are strongly conserved by purifying
selection within the plant kingdom. The extracellular
domain of AtFH1 that seems most responsible for immobilization of the protein is Domain C in construct AtFH1-TR3.
In the absence of this domain, AtFH1 was mobile within the
PM. Domain C is only comprised of 15 amino acids that,
intriguingly, include an SPPPP motif which is the signature
peptide of extensin, a class of cell-wall associated proteins
ª 2011 The Authors
The Plant Journal ª 2011 Blackwell Publishing Ltd, The Plant Journal, (2011), 66, 354–365
362 Alexandre Martinière et al.
Figure 6. A model of AtFH1 bridging between the cell wall and the actin cytoskeleton. AtFH1 (yellow) spans the plasma membrane (blue) connecting the actin
cytoskeleton (red) and the cell wall (indicated as cellulose microfibrils in brown). AtFH1 is an actin nucleating protein that binds the side of actin microfilaments via
its intracellular formin homology domains. When AtFH1 is overexpressed, dense sheets of actin form. This actin modelling function requires that AtFH1 link to both
the cell wall and to actin.
(Banno and Chua, 2000; Showalter et al., 2010) so it seems
reasonable that this motif confers the cell-wall interacting
function.
The biochemical nature of the cell wall anchoring interaction is not yet understood. It might be either a protein–
protein interaction or an interaction with other cell wall
components. AtFH1’s extracellular region contains a high
percentage of serine and proline residues, which could carry
O-linked oligosaccharide modifications. In this case, carbohydrates might also be responsible for the cell wall interaction.
Interestingly, AtFH4 and AtFH5 have specific patterns of
plasma membrane localization – AtFH4 strongly accumulates in cell-to-cell contact zones and AtFH5 accumulates at
the pollen-tube tip (Deeks et al., 2005; Cheung et al., 2010).
The fact that formins are targeted to a sub-domain of the PM
might reflect heterogeneity of cell wall composition.
The cell wall connection is required for AtFH1-induced
remodelling of the actin cytoskeleton
The bridging topology of AtFH1 makes us think that this
protein may participate in signal transduction between
outside stimuli and the actin cytoskeleton. Here, we show
that the extracellular region of AtFH1 – especially Domain C –
is required to effect a change in shape of the actin network
when AtFH1 is overexpressed (compare Figure 4g,h). Similarly, only AtFH1 with intact Domain C was able to strongly
reduce long distance movement of Golgi bodies (compare
Figure 5f,g) and, as stated in the previous section, Domain C
seems to be involved in anchoring of AtFH1 to the ECM.
These results, taken together, show that anchoring to the
ECM via Domain C is needed for inducing changes in the
actin network as observed in AtFH1-overexpressing cells.
Several molecular mechanisms can be proposed for
explaining the link between anchoring in the extracellular
space and actin nucleation or actin bundling within the cell.
Animal cell formin needs to form a dimer to be functional
(Harris et al., 2004). Isolated intracellular domains FH1 and
FH2 of AtFH1 are able to form dimers in in vitro assays, and
dimers are probably necessary for formin1 activity (Michelot
et al., 2005, 2006). Consequently, we can extrapolate that
the full protein is also able to form functional dimers. In the
plasma membrane, both non-functional monomer and
functional dimers may co-exist. We speculate that anchoring
of AtFH1 dimers in the cell wall has an effect on their
stabilization and conformation. In absence of anchoring,
the dimer might be less stable, less likely to form or might
not possess the active conformation and consequently the
protein will not induce the observed meshwork-like array of
microfilaments.
AtFH1 labelling forms a negative microtubule pattern
on the plasma membrane
Several animal formins have been described as interacting
directly with microtubules (Chesarone et al., 2010). This
phenomenon is involved in coordination of the two main
cytoskeletal networks: actin microfilaments and microtubules. Recently, it was reported that plant formin also
interacts with microtubules (Deeks et al., 2010; Li et al.,
2010). The microtubule shadow observed in the plasma
membrane when AtFH1 is overexpressed may also be the
result of a physical interaction between AtFH1 and microtubules (Figure 1c). But more likely the exclusion of AtFH1
from PM regions overlying the microtubules is because
the large cytoplasmic domain of AtFH1 is corralled by, and
cannot cross, cortical MTs which are also anchored to the
plasma membrane (Pesquet et al., 2010). The exact nature
and the biological significance of this phenomenon are yet
to be determined.
EXPERIMENTAL PROCEDURES
Constructs and cloning procedures
The cDNA of At3g25500 (AtFH1), ABRC clone number U11142 was
used as a template for all AtFH1 constructs. For AtFH1 (nucleotide
1–3156), AtFH1-dFH (nucleotide 1–420) and AtFH1-107 (nucleotide
1–323) PCR fragments were cloned in pENTR-D-TOPO (Invitrogen).
ª 2011 The Authors
The Plant Journal ª 2011 Blackwell Publishing Ltd, The Plant Journal, (2011), 66, 354–365
AtFH1 links the cell wall and actin cytoskeleton 363
For AtFH1-dECM (nucleotide 1–72 fused to nucleotide 315–3156),
AtFH1-TR1 (nucleotide 1–102 fused to nucleotide 294–3156), AtFH1TR2 (nucleotide 1–102 fused to nucleotide 240–3156) and AtFH1-TR3
(nucleotide 1–102 fused to nucleotide 189–3156) overlapping PCR
were performed before the cloning via TOPO reaction. For CFP–
TuA6 or YFP-TuA6, TUBULINa6 was amplified from GFP6-TuA6
(Ueda et al., 1999) and cloned into the pENTR-D-TOPO vector. The
following Gateway (Invitrogen) binary vectors were selected to do
N- and C-terminal fusions of cloned PCR products with various
fluorescent proteins: pB7RWG2, pB7FWG2, pB7GWC2, pB7CWG2,
pB7GWY2, pB7YWG2 (Plant Systems Biology, Ghent University).
pB2GW7 was used to make the non-fluorescent protein tagged
AtFH1 (Plant Systems Biology, Ghent University). All final clones
were verified by sequencing.
Plant transformation and confocal microscopy
Nicotiana tabacum cv. Petit Havana plants were grown and used in
transient transformation experiments as described in Sparkes et al.
(2006). All the formin constructs, sialyl transferase signal anchor
sequence fused to mRFP (ST-mRFP) (Saint-Jore et al., 2002), GFPLTI6b (Runions et al., 2006) and Sec-mRFP (Samalova et al., 2006)
were infiltrated using Agrobacterium tumefaciens GV3101 at 0.1
OD600 (Sparkes et al., 2006). Cytoskeleton labelling probes; Lifeact–
GFP (Deeks et al., 2010), Lifeact–RFP (Berepiki et al., 2010), GFPFABD2 (Sheahan et al., 2004), GFP-mTn (Kost et al., 1998) and
C/YFP-TuA6, were infiltrated at 0.01 OD600 to prevent side effects
of overexpression (Runions et al., 2006). Two to three days after
transformation leaf tissue was observed with a Zeiss (http://
www.zeiss.com/) LSM 510 META confocal system.
Drugs treatments, cell wall labelling and plasmolysis assays
Leaf pieces (approximately 0.25 cm2) were perfused by immersion
in 25 lM latrunculin B (30–180 min) to remove the actin cytoskeleton (stock solution 2.5 mM in DMSO), 20 lM oryzalin (30 min) to
remove microtubules (stock solution 50 mM in DMSO), or with
357 lM Brefeldin A (BFA, 3 h for ST-mRFP and 6 h for AtFH1) to
block retrograde secretion. For protoplast experiments (see below),
primary-cell-wall regeneration was monitored by staining with
1 lg ml)1 of Calcofluor White M2R from a stock solution at
1 mg ml)1 in DMSO. For the plasmolysis assay, leaf sections
(approximately 0.25 cm2) were incubated with a hypertonic solution
of 0.5 M mannitol for 1 h.
Quantification of the relative mobile fraction
of formin constructs
The relative mobile fraction of formin fused to different fluorescent
proteins was assessed by Fluorescence Recovery After Photobleaching (FRAP). A 63· (1.4 NA) objective was used at a zoom setting
of 5. Pre-bleaching and post-bleaching excitation was done using
either the 488 nm line of an argon-ion laser (GFP) or the 514 nm
line (YFP) set at 50% output and 3% transmission. Five scans of the
entire field of view were made to establish the pre-bleach intensity
of the fluorescent protein and then a circular region of interest
(ROI) 8 lm2 in a median optical section of the fluorescent plasma
membrane was bleached. Ten iterations of the 488 or 514 nm laser
set at 100% transmission were used for bleaching. Recovery of
fluorescence was recorded during 115 sec with a delay of 1.5 sec
between frames. Images were 256 · 256 pixels and were made
with a scan speed of 0.46 sec per frame. We confirmed that the
energy of the 488 or 514 laser used to record post-bleach data had
no bleaching effect by recording a control region outside of the
bleaching ROI. Fluorescence intensity data were normalized using
the equation:
In ¼ ½ðIt Imin Þ=ðImax Imin Þ 100
Where In is the normalized intensity, It is the intensity at any time t,
Imin is the minimum post-photobleaching intensity and Imax is the
mean pre-photobleaching intensity.
Non-linear regression was used to model the normalized FRAP
data. In this case, a two-phase exponential association equation was
used:
Y ðtÞ ¼ A þ Bð1 þ expðK1 ÞðtÞ Þ þ Cð1 expðK2 ÞðtÞ Þ
Where Y(t) is normalized intensity, A, B, C, K1 and K2 are parameters
of the curve, and t is time.
For each treatment, 7–13 cells were analysed. The value of the
fluorescence intensity recovery plateau (t = 110 sec) was calculated
and used as an approximation of the relative mobile fraction.
Protoplast preparation
Fusion-protein-expressing tobacco leaf pieces were submerged in a
20· dilution of DOMESTOS commercial bleach solution (Unilever)
for 1 min and rinsed several times with distilled water. The leaf
tissue was then incubated for 15 h in 0.5 M mannitol, 1% cellulase
R10, 0.1% macerozyme, 1 mM CaCl2 and 10 mM MES pH 5.6. After
filtration, protoplasts were rinsed twice with 0.5 M mannitol, 10 mM
glucose, 1 mM CaCl2 and 10 mM MES pH 5.6. An aliquot of cells was
used immediately as a time 0 sample. Protoplasts were left at room
temperature in light for 24 and 48 h time points.
Golgi body mobility
For recording the mobility of Golgi bodies, ST-mRFP-expressing
cells were imaged (0.45 sec frame)1) for 49 sec. The data were
analysed with Volocity 4 (Improvision), according to Runions et al.
(2006). The meandering index (MI) was calculated as the ratio
between Golgi body displacement (length of the straight line
connecting start to end point) and track length (length of the actual
path followed). For each treatment 54–138 Golgi bodies were
followed for 10 different cells.
Calculation of selection acting on AtFH1–PAML analysis
The AtFH1 sequence from A. thaliana (NC_003074) was used to
retrieve coding sequences of AtFH1 orthologous genes in public
databases from Ricinus communis (XM_002532408), Populus
trichocarpa (XM_002310361), Vinis vitifera (XM_002274965), Medicago truncatula (AC171534), N. tabacum (AF213695), Oryza sativa
v. Japonicum (AP003349) and Sorghum bicolour (XM_002458874).
Nucleotide alignment was performed using TRANSLATOR X (Abascal
et al., 2010) using the default settings of the software. Alignment of
the whole gene was then partitioned into three alignments,
according to putative functional domains of the protein. Part 1 is
from the 5¢ part to the end of the putative extracellular domain of
AtFH1 (nt 1–323 in A. thaliana sequence). Part 2 is from the transmembrane domain to the beginning of the FH1 domain (nt 324–1560
in A. thaliana sequence). Part 3 is the 3¢ part of the gene and contains FH1 and FH2 domains (nt 1561–3156 in A. thaliana sequence).
Estimations of the nonsynonymous to synonymous substitution
rate (x = dN/dS) were performed using Phylogenetic Analysis by
Maximum Likelihood (PAML) with the CODEML program implemented in PAML 4.3 (Yang, 2007). Phylogenetic trees required for
PAML were then inferred from protein sequence data of each of the
three parts of the protein. To do so, the software GBLOCS (Talavera
and Castresana, 2007) was used to eliminate poorly aligned
positions and divergent regions of an alignment of protein
sequences. These positions may not be homologous or may have
been saturated by multiple substitutions and it is convenient to
ª 2011 The Authors
The Plant Journal ª 2011 Blackwell Publishing Ltd, The Plant Journal, (2011), 66, 354–365
364 Alexandre Martinière et al.
eliminate them prior to phylogenetic analysis (Castresana, 2000).
Phylogenies were inferred by Phyml (Guindon and Gascuel, 2003).
The GTR substitution model was selected assuming a proportion of
invariant sites (0.01). To account for rate heterogeneity across sites,
the gamma distribution with four rate categories was used.
ACKNOWLEDGEMENTS
The authors would like to thank Dr Michael Deeks for discussions
and comments on an early version of the manuscript and Dr Jens
Tilsner who provided us with the binary vector for Lifeact–RFP. This
research was funded by BBSRC Grant number BB/F014074/1.
SUPPORTING INFORMATION
Additional Supporting Information may be found in the online
version of this article:
Figure S1. Amino acid alignment of different Formin1 proteins from
the plant kingdom.
Figure S2. AtFH1 is localized at the plasma membrane.
Figure S3. AtFH1–GFP is targeted to the plasma membrane via the
secretory pathway.
Figure S4. Removing microtubules affects the pattern of AtFH1
localization at the plasma membrane.
Figure S5. Sub-cellular localization of the different AtFH1 extracellular region deletions.
Figure S6. AtFH1 overexpression induces a major re-organization of
the actin cytoskeleton.
Please note: As a service to our authors and readers, this journal
provides supporting information supplied by the authors. Such
materials are peer-reviewed and may be re-organized for online
delivery, but are not copy-edited or typeset. Technical support
issues arising from supporting information (other than missing
files) should be addressed to the authors.
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ª 2011 The Authors
The Plant Journal ª 2011 Blackwell Publishing Ltd, The Plant Journal, (2011), 66, 354–365