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The Type II Arabidopsis Formin14 Interacts with Microtubules
and Microfilaments to Regulate Cell Division
W OA
Yanhua Li,a Yuan Shen,a Chao Cai,a Chenchun Zhong,a Lei Zhu,b Ming Yuan,b and Haiyun Rena,1
a Key Laboratory of Cell Proliferation and Regulation Biology of Ministry of Education, College of Life Science, Beijing Normal
University, Beijing 100875, People’s Republic of China
b State Key Laboratory of Plant Physiology and Biochemistry, China Agricultural University, Beijing 100094, People’s Republic of
China
Formins have long been known to regulate microfilaments but have also recently been shown to associate with
microtubules. In this study, Arabidopsis thaliana FORMIN14 (AFH14), a type II formin, was found to regulate both
microtubule and microfilament arrays. AFH14 expressed in BY-2 cells was shown to decorate preprophase bands, spindles,
and phragmoplasts and to induce coalignment of microtubules with microfilaments. These effects perturbed the process of
cell division. Localization of AFH14 to microtubule-based structures was confirmed in Arabidopsis suspension cells.
Knockdown of AFH14 in mitotic cells altered interactions between microtubules and microfilaments, resulting in the
formation of an abnormal mitotic apparatus. In Arabidopsis afh14 T-DNA insertion mutants, microtubule arrays displayed
abnormalities during the meiosis-associated process of microspore formation, which corresponded to altered phenotypes
during tetrad formation. In vitro biochemical experiments showed that AFH14 bound directly to either microtubules or
microfilaments and that the FH2 domain was essential for cytoskeleton binding and bundling. However, in the presence of
both microtubules and microfilaments, AFH14 promoted interactions between microtubules and microfilaments. These
results demonstrate that AFH14 is a unique plant formin that functions as a linking protein between microtubules and
microfilaments and thus plays important roles in the process of plant cell division.
INTRODUCTION
Microtubules and microfilaments represent two major plant cell
cytoskeletal networks, both of which play important roles in
many aspects of the fundamental processes of plant cell growth
and development, including cell division, cell expansion, intracellular organization, and cell motility. There is no question that
microtubules and microfilaments constitute separate cytoskeletal systems and fulfill distinct functions. However, a growing
body of evidence also suggests that functional interactions
between microtubules and microfilaments are important for
specific cellular processes. In animal cells, microtubules have
been proposed to contribute to these processes in part by
mediating proper spatial distribution of microfilament structures.
For example, during cytokinesis, the microtubules of the mitotic
spindle have been shown to play an important role in the
positioning of the contractile microfilament ring assembly at
the cell cortex (Maddoxa and Oegema, 2003). During cell migration, microtubules have been implicated in steering the microfilament cytoskeleton in the proper direction, suggested by
1 Address
correspondence to [email protected].
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findings presented in this article in accordance with the policy described
in the Instructions for Authors (www.plantcell.org) is: Haiyun Ren
([email protected]).
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www.plantcell.org/cgi/doi/10.1105/tpc.110.075507
characterization of nerve growth cone guidance (Tanaka et al.,
1995; Dent and Gertler, 2003).
In plant cells, microtubules and microfilaments are often codistributed in the cortical area in interphase cells (Blancaflor, 2000)
and colocalize in structures, such as the preprophase band,
mitotic spindle, and phragmoplast, in mitotic cells (Hoshino et al.,
2003) and meiotic cells (Staiger and Cande, 1991). Pharmacological studies using microfilament- and microtubule-specific
drugs have demonstrated that microtubules interact with microfilaments. For example, cytochalasin, a microfilament-disrupting
drug, prevents the microtubule preprophase band from narrowing in Allium cepa (Eleftheriou and Palevitz, 1992). In addition,
long-term treatment with a low level of cytochalasin causes
transverse microtubule arrays to reorient into an oblique alignment in growing cotton (Gossypium hirsutum) hairs (Seagull,
1990). These results suggest that microfilaments can influence
microtubule organization. This relationship appears to be reciprocal, as microtubules have also been shown to affect microfilament organization (Collings and Allen, 2000). Recently, a few
proteins have been shown to interact with both microtubules and
microfilaments in plant cells (Petrášek and Schwarzerová, 2009).
However, the functional evidence for these interactions is still
lacking, and the molecular basis underlying interactions between
the microtubule and microfilament cytoskeletons in plants is
poorly understood.
Formins are known regulators of the microfilament cytoskeleton (Blanchoin and Staiger, 2008). However, it has been
noted recently that formins also behave as prominent regulators
of the microtubule cytoskeleton by modulating the dynamics of
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selected microtubules in interphase and mitosis (Bartolini and
Gundersen, 2009; DeWard and Alberts, 2009; Chesarone et al.,
2010). Some metazoan formins have also been shown to associate with microtubules through interactions with microtubuleassociated proteins, including CLIP170, APC, and end binding
protein 1 (EB1), via sequences located outside of the formin FH2
domain (Wen et al., 2004; Lewkowicz et al., 2008). Recent
evidence has suggested that many formins directly bind microtubules and affect their dynamic properties. For example, experiments using a constitutively active form of mouse formin
mDia2 lacking regulatory domains have provided an initial indication that this formin directly interacts with microtubules to alter
their dynamics (Bartolini et al., 2008). Subsequent studies have
shown that several animal formins, including mDia1, mDia2,
Formin1-Ib, INF1, and Capu, interact directly with microtubules
(Rosales-Nieves et al., 2006; Zhou et al., 2006; Bartolini et al.,
2008; Young et al., 2008). Furthermore, characterization of mDia2
has also suggested the possibility that formins may interact
with microfilaments while binding to microtubules (Bartolini
et al., 2008).
Recently, significant progress has been made toward understanding the biochemical properties and cellular functions of plant
formins. Plant formins can be divided into three classes (Cvrčková
et al., 2004; Grunt et al., 2008). However, all of the reported plant
formins, including the Class I Arabidopsis thaliana formins AFH1,
AFH3, AFH4, AFH5, AFH6, and AFH8 (Deeks et al., 2005; Ingouff
et al., 2005; Michelot et al., 2005; Yi et al., 2005; Ye et al., 2009) and
a Class II formin from Physcomitrella, For2 (Vidali et al., 2009), have
been characterized as microfilament regulators (Blanchoin and
Staiger, 2008), with the lone exception of AFH4. Recently, AFH4
was reported to bind microtubules through a newly discovered
GOE domain, when overexpressed in the leaf cells of Nicotiana
benthamiana (Deeks et al., 2010).
In this study, we identified and characterized a previously
undiscovered Arabidopsis formin, FORMIN14 (AFH14). We demonstrate that AFH14 plays an important role in regulating both
microtubule and microfilament arrays through association with
these networks in mitotic BY-2 and Arabidopsis suspensioncultured cells. Furthermore, we found that AFH14 is involved in
meiosis through regulation of microtubule structures required for
the generation of microspores. To further support these observations, AFH14 was also shown to interact with microtubules and
microfilaments and to affect the structures of these polymers in
vitro.
RESULTS
Identification of AFH14 and Preparation of
AFH14 FH1FH2 Domain Recombinant Protein and
AFH14-Specific Antibodies
AFH14 was identified based on sequence analysis of the Arabidopsis genome. The gene was found to contain 18 exons and 18
introns, spanning 6653 bp, and to encode a 3102-nucleotide
mRNA with a single open reading frame estimated to produce a
113.6-kD protein of 1033–amino acid residues. The AFH14
protein consisted of three functionally distinct subdomains: an
N-terminal PTEN (phosphatase tensin)–related domain, a Prorich FH1 domain, and a highly conserved C-terminal FH2 domain
(see Supplemental Figure 1 online). However, a 591-bp region
within the FH1 domain was absent in the amplified sequence,
differing from The Arabidopsis Information Resource prediction
(see Supplemental Figure 2 online).
To characterize the chemical properties of AFH14 in vitro, a
6-His–tagged truncated recombinant protein (referred to as
FH1FH2) containing the FH1 and FH2 domains of AFH14 was
expressed and purified from bacterial cells. Polyclonal antibodies directed against the FH1FH2 recombinant protein were
then raised in mice. The molecular mass of the purified FH1FH2
proteins was estimated to be ;80 kD by SDS-PAGE. Immunoblot analysis revealed that the polyclonal antibodies effectively
recognized the recombinant protein (see Supplemental Figure
3A online). Because the FH1FH2 region is highly conserved
across formin family members, we expected that the polyclonal
antibodies were likely also to recognize other formins. Therefore,
these anti-AFH14 antibodies were exclusively used to examine
the biochemical characteristics of AFH14 in vitro. To explore
endogenous AFH14 localization in Arabidopsis cells, we raised a
monoclonal antibody directed against AFH14 in mice using a
fragment of the protein consisting of amino acids 340 to 502 (see
Supplemental Figure 1B online). The specificity of the monoclonal antibody was confirmed using protein extracts from 10-d-old
Arabidopsis seedlings (see Supplemental Figure 3B online).
AFH14 Localizes to the Preprophase Band, Spindle,
and Phragmoplast
To assess the localization of AFH14, a chimeric expression
construct harboring AFH14 fused to green fluorescent protein
(GFP) under the control of an inducible promoter was stably
transformed into wild-type tobacco (Nicotiana tabacum) BY-2
cells. Approximately 30 h after induction, cells were fixed and
microtubules were visualized using an anti-a-tubulin antibody.
The subcellular localizations of AFH14-GFP and microtubules
were then examined by confocal microscopy. In preprophase
cells, AFH14-GFP localized primarily to the preprophase bands
(Figure 1A). In metaphase and anaphase cells, AFH14-GFP was
concentrated along the entire spindle (Figures 1B and 1C). In
cells undergoing cytokinesis, AFH14 completely colocalized with
the two mirror halves of the phragmoplast (Figure 1D).
To confirm further the localization of endogenous AFH14,
antibodies directed against AFH14 and a-tubulin were used to
stain Arabidopsis suspension cells. In dividing cells, AFH14
colocalized with a-tubulin at the preprophase bands (Figure 1E),
spindles (Figure 1F), and phragmoplasts (Figure 1G) in agreement with the observations in BY-2 cells.
Overexpression of AFH14 Inhibits Turnover of the Mitotic
Apparatus and Increases the Resistance of Microtubules
to Oryzalin
To explore the contribution of AFH14 to the function of the mitotic apparatus, live cell imaging studies of mitotic progress were
performed. At 30 h postinduction with estrogen, there was a
pronounced increase in GFP signal in induced BY-2 cells
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control uninduced cells (Figure 2C). The mitotic index of cells
transformed with an empty vector carrying only GFP in the
presence or absence of estrogen treatment was 4.2 and 4.0%,
respectively, similar to uninduced cells from the AFH14-exressing line.
Based on the observations described above, we hypothesized
that overexpression of AFH14 may alter the resistance of microtubules to the microtubule-depolymerizing drug oryzalin. Treatment with 10 mM oryzalin had a strong effect on microtubules in
uninduced control cells, as 75.0% of spindles and 82.0% of
phragmoplasts were disrupted after 15 min of treatment (see
Supplemental Figures 4A and 4B online). Overexpression of
AFH14 partially suppressed the effect of oryzalin (Figures 2D and
2E), resulting in a decrease in the percentage of depolymerized
spindles and phragmoplasts to 52.0 and 58.2%, respectively
(Figure 2F). In addition to the effects of AFH14 on microtubule
stability, we also found that oryzalin-mediated microtubule depolymerization resulted in changes in the localization of AFH14GFP. GFP fluorescence was either associated with irregular
microtubule arrays or dispersed in the cytoplasm after oryzalin
treatment (see Supplemental Figures 5A and 5B online), further
confirming that AFH14 binds microtubules.
AFH14 Recruits Microfilaments to the Spindle
and Phragmoplast
Figure 1. AFH14 Is Associated with Microtubule Arrays in Plant Cells.
(A) to (D) AFH14-GFP colocalized with microtubule arrays (visualized
with tubulin antibody) at the preprophase band (A), metaphase spindle
(B), anaphase spindle (C), and cytokinetic phragmoplast (D) in BY-2 cells
overexpressing AFH14.
(E) to (G) AFH14 immunostaining revealed that endogenous AFH14
colocalized with microtubule arrays at the preprophase band (E), metaphase spindle (F), and cytokinetic phragmoplast (G) in wild-type Arabidopsis suspension-cultured cells.
Chromosomes were labeled by 49,6-diamidino-2-phenylindole (DAPI).
Bars = 5 mm.
expressing AFH14 in comparison to control cells without estrogen treatment. Interestingly, turnover of the spindles and phragmoplasts was severely inhibited in the BY-2 cells overexpressing
AFH14-GFP (Figures 2A and 2B). Failure of the turnover of
spindles and phragmoplasts resulted in a lack of progression
through mitosis in AFH14-expressing cells, which was reflected
by a high mitotic index score. Overexpression of AFH14 resulted
in an increased mitotic index of 8.8%, compared with 3.9% for
Significant evidence indicates that the classical formins play
important roles in microfilament organization. Therefore, fluorescently labeled phalloidin was used to visualize microfilaments
in BY-2 cells. In control uninduced cells, microfilaments formed a
cage around the spindle, and very few filaments remained in the
spindle (Figure 3A). In addition, the microfilament band in control
cells was thinner than the microtubule band in phragmoplasts
(Figure 3B). Interestingly, in cells overexpressing AFH14-GFP,
microfilaments codistributed with AFH14-GFP in spindles and
phragmoplasts (Figures 3C and 3D). Furthermore, similar to
results indicating resistance to microtubule depolymerizing
drugs, overexpression of AFH14 partially suppressed the effects
of treatment with the microfilament-depolymerizing drug latrunculin B (LatB). In BY-2 cells treated with 200 nM LatB for 50 min,
65.4% of AFH14-overexpressing cells exhibited intact mitotic
microfilament structures, compared with only 31.6% of control
cells (see Supplemental Figure 6 online). In addition, depolymerization of microfilaments with LatB did not appreciably affect the
spindle and phragmoplast pattern in cells expressing AFH14GFP (Figures 3E and 3F). Together, these results indicate that
AFH14 likely interacts with both microtubules and microfilaments
in vivo, although it binds preferentially to microtubules.
AFH14 Affects the Interaction between Microtubules
and Microfilaments
To determine whether connections between microtubules and
microfilaments were necessary for cell division in Arabidopsis
cells, codistribution of microtubules and microfilaments was
analyzed in wild-type suspension-cultured cells. Similar to observations in BY-2 cells, microtubules were arranged to form the
mitotic spindle concurrent with formation of a microfilament cage
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Figure 2. Overexpression of AFH14 Inhibits Turnover of the Mitotic Apparatus and Increases the Resistance of Microtubules to Oryzalin in BY-2 Cells.
Wild-type BY-2 cells were transformed with inducible AFH14-GFP vectors. After induction of expression, the cells were observed at the indicated time
points ([A] to [C]) or were treated with 10 mM oryzalin for 15 min ([D] to [F]). Mock-induced cells treated with DMSO served as negative controls.
(A) and (B) Serial images show representative cells arrested at metaphase (A) and cytokinesis (B). For each stage, at least 15 cells were observed and
exhibited similar results.
(C) The mitotic index was analyzed in AFH14-overexpressing cells (OX) and control cells. The mitotic index of OX cells was found to be significantly
different than that of control cells (t test, P < 0.05). Error bars indicate SE (n > 1500). Results represent data from three independent experiments.
(D) and (E) The metaphase spindle (D) and cytokinetic phragmoplast (E) remained partially intact after treatment with oryzalin in OX cells. Bar = 5 mm.
(F) The number of mitotic cells with depolymerized microtubules (MT) was analyzed in OX and control BY-2 cells. Significant differences in levels of
spindle and phragmoplast microtubule depolymerization between OX and control cells were observed, respectively (t test, P < 0.05). Error bars indicate
SE (n > 300). Results represent data from three independent experiments.
surrounding the spindle (Figure 4A). In cytokinesis, the microfilament band appeared thinner than the microtubule band in
phragmoplasts (Figure 4B).
To assess the effects of reduced AFH14 expression on microtubule and microfilament arrays, an artificial microRNA construct was prepared and placed under the control of an inducible
promoter. Uninduced cells without estrogen treatment were
used as a negative control, and the distribution of microfilaments and microtubules in these cells was similar to that of wildtype cells (see Supplemental Figure 7 online). For transformed
cells induced with 2 mM estrogen for 3 d, changes in cellular
phenotypes were classified as normal, mild, strong, or severe,
depending upon the severity of the resulting abnormalities.
Approximately 25.5% of mitotic arrays in the treated cells
exhibited no visible phenotypic changes, and the mitotic apparatuses had normal microtubule and microfilament arrangements similar to control cells. Mild abnormalities, in which the
spindle and phragmoplast microtubules seemed intact, but the
microfilament cages around the spindles and microfilaments in
phragmoplasts were not present, were observed in 27.7% of the
cells (Figures 4C and 4D). An additional 14.9% of cells exhibited
strong phenotypic changes, in which only a remnant of mitotic
microtubules were observed, accompanied by severely dispersed microfilaments (Figure 4E). In the remaining 31.9% of
cells that showed severe phenotypic abnomalities, severely
depolymerized microtubules and microfilaments were observed
(Figure 4F). None of these phenotypes were observed in control
uninduced cells or in cells transformed with an empty vector
cultured in the presence or absence of estrogen.
AFH14 Loss-of-Function Plants Exhibit Defective
Microspore Generation
The AFH14 tissue expression pattern was analyzed by transforming Arabidopsis with an AFH14 promoter:b-glucuronidase
(GUS) reporter gene transcriptional fusion construct. Three independent GUS reporter lines were examined, and all showed
the same pattern of expression. In seedlings, high levels of GUS
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To characterize the function of AFH14 in vivo, two distinct
Arabidopsis mutant alleles harboring T-DNA insertions in AFH14
were identified. The afh14-1 allele harbored a T-DNA insertion in
the second exon, and the afh14-2 allele harbored an insertion in
the 39 untranslated region (see Supplemental Figure 9 online).
Further characterization revealed that both mutants exhibited a
similar phenotype with respect to decreased pollen counts
(Figure 5A). To explore further the underlying reasons for this
phenotype, meiosis was compared between the wild-type and
mutant plants. In wild-type plants, all tetrads observed had four
spores, and the tetrahedral microspore arrangement appeared
normal (Figure 5B). By contrast, the afh14-1 and afh14-2 lines
produced dyads, triads, polyads, and abnormal tetrads with an
irregular microspore distribution patterns (Figures 5C to 5F; see
Supplemental Figures 10B to 10E online). These phenotypes
accounted for 40.0% of all meiotic tetrads. In addition, no tetrads
from afh14-1 and afh14-2 had the swollen domain present in all
wild-type tetrads (Figure 5B; see Supplemental Figure 10A
online).
AFH14 Affects the Arrangement of Microtubule Arrays in
Microspore Formation
Figure 3. AFH14 Binds Microfilaments in BY-2 Cells, but Localization of
AFH14 Is Not Dependent on Microfilaments.
(A) In control BY-2 cells, microfilaments (visualized with phalloidin)
formed a cage around the spindle with few microfilaments present in
the spindle. Control cells were mock induced by treatment with DMSO.
(B) Microfilaments were clearly present in the region of the phragmoplast
microtubules in control BY-2 cells.
(C) and (D) AFH14-GFP codistributed with microfilaments in the spindle
(C) and phragmoplast apparatus (D) in AFH14-GFP–overexpressing (OX)
cells.
(E) and (F) Depolymerization of microfilaments with LatB had no appreciable effect on spindle (E) or phragmoplast (F) localization of AFH14GFP. Bar = 5 mm.
activity were detected in the shoot apex, leaf primordia, guard
cells, trichomes, and roots, with the exception of the root tip (see
Supplemental Figures 8A to 8D online). In the inflorescence, GUS
activity was strong in all floral buds (see Supplemental Figures 8E
and 8F online). However, GUS activity could not be detected in
the wild-type floral buds (see Supplemental Figure 8G online).
This pattern is consistent with the expression of AFH14 reported in publicly available expression array data (http://www.
genevestigator.ethz.ch/).
We first analyzed the subcellular localization of AFH14 during
meiosis in pollen mother cells (PMCs) by immunofluorescence
microscopy. As shown in Figure 6, the AFH14 protein was bound
to spindle microtubules at metaphase I and II and to phragmoplast microtubules at anaphase I and advanced cytokinesis
(Figures 6B, 6C, 6E, and 6G). Although the radial microtubule
system (RMS) was present surrounding the nuclei during prophase I, telophase I, early cytokinesis, and in tetrads, AFH14 was
dispersed throughout the cytoplasm in these phases (Figures 6A,
6D, 6F, and 6H).
Next, the microtubule arrays were compared in wild-type and
mutant plants during the process of microspore generation.
These results showed that microtubule arrays behaved irregularly in afh14-1 and afh14-2 PMCs. For example, metaphase I
spindles exhibited a typical fusiform configuration in wild-type
cells (Figure 7A) but became visibly smaller in afh14-1 cells
(Figure 7D). At telophase I, two RMSs formed properly around the
nucleus in wild-type cells (Figure 7B) but displayed a skewed
configuration in afh14-1 cells (Figure 7E). In addition, the two
adjacent spindles present during metaphase II were perpendicular to one another in wild-type cells (Figure 7C) but were parallel
to one another in 58.4% of afh14-1 cells (n = 24) (Figure 7F).
Furthermore, in wild-type cells, four RMSs surrounded four nuclei
symmetrically during early cytokinesis (Figure 7G), whereas in
afh14-1 cells, the microtubules became slightly waved (Figure
7J). Also, during advanced cytokinesis, the phragmoplast microtubules were concentrated between two adjacent nuclei
(Figure 7H), but this was not the case in 100% of afh14-1 cells
observed (n = 36) (Figure 7K). Finally, in wild-type cells, the tetrad
developed into four microspores with microtubules radiating
from each nucleus (Figure 7I). However, the afh14-1 mutants
produced some dyads that included two nuclei in each spore,
although microtubules still radiated from each nucleus in some
afh14-1 cells (Figure 7L). These observations indicate that
mutations in AFH14 have a deleterious effect on microtubule
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was quantified by SDS-PAGE and densitometry (Figures 8A and
8C). In the absence of microtubules, FH1FH2 was found predominantly (>90%) in the supernatant (Figure 8A, lane 17), indicating
that the recombinant protein alone did not sediment. However,
with the addition of microtubules, FH1FH2 was largely found in the
pellet with the microtubules. Therefore, we conclude that FH1FH2
can bind and cosediment with taxol-stabilized microtubules in
vitro, behavior typical of a microtubule binding protein.
We next analyzed the binding activity of FH1FH2 for microfilaments using a high-speed cosedimentation assay. As shown in
Figures 8B and 8D, addition of increasing concentrations of
FH1FH2 to microfilaments resulted in concentration-dependent
increases in the amount of FH1FH2 present in the pellets. These
results indicate that, in addition to microtubules, FH1FH2 also
binds microfilaments in vitro.
Additional experiments assessing the effects of FH1FH2 on
actin polymerization were also performed using the high-speed
cosedimentation assays. These results showed that, in samples
containing FH1FH2, more microfilaments were present in the
pellets (with a concomitant decrease in the supernatants) than
when FH1FH2 was not added during the first 5 min of the assay
(see Supplemental Figure 12 online), indicating that FH1FH2 can
bind and nucleate microfilaments.
The AFH14 FH1FH2 Domain Bundles Microtubules
and Microfilaments
Figure 4. AFH14 Modulates Interactions between Microtubules and
Microfilaments in Arabidopsis Suspension-Cultured Cells.
(A) and (B) Representative images showing the distributions of microfilaments and microtubules in wild-type cells. Cortical microfilaments
formed a cage around the spindle in wild-type cells (A). Microfilaments
were clearly present in regions near the phragmoplast microtubule plus
ends in wild-type cells (B).
(C) to (F) Wild-type Arabidopsis suspension-cultured cells were transformed with inducible amiR-AFH14 RNA interference vectors. After
induction, microfilaments and microtubules were observed by fluorescence microscopy. Cells with the mild phenotype lacked the microfilament cage around the spindle (C) and the microfilaments located in the
region of the phragmoplast (D). Only dispersed microfilaments and a few
remnants of microtubules were observed in cells that exhibited the
strong phenotype (E). Both the microfilaments and microtubules were
dispersed in cells with the severe phenotype (F).
Bars = 5 mm.
arrangement. The afh14-2 mutant displayed similar microtubule
array defects to those observed for afh14-1 (see Supplemental
Figure 11 online).
The FH1FH2 Domain of AFH14 Directly Binds Both
Microtubules and Microfilaments in Vitro
Because AFH14 colocalized with microtubules and microfilaments in vivo, the direct binding activity of FH1FH2 to the two
cytoskeletal polymers was analyzed in vitro. To assess microtubule binding activity, taxol-stabilized microtubules were incubated with varying concentrations of the recombinant protein,
and the amount of tubulin and FH1FH2 in the resulting pellets
Because the AFH14 FH1FH2 recombinant proteins were found
to bind directly to microtubules and microfilaments in vitro, we
further analyzed the effects of the FH1FH2 protein on microtubule and microfilament organization. First, confocal microscopy
was used to examine the effects of 1 mM FH1FH2 on rhodamineconjugated microtubules. In the absence of FH1FH2 and in the
presence of heat-denatured FH1FH2, microtubules were scattered individually throughout the solution (see Supplemental
Figure 13A online). However, in the presence of active FH1FH2,
the microtubules organized almost exclusively into densely
packed bundles, with few microtubules found outside of the
bundles (see Supplemental Figure 13B online). Consistent with a
role for FH1FH2 in this process, FH1FH2 staining was observed
in dot-like structures along the lengths of the microtubules (see
Supplemental Figure 13C online). To investigate further this
microtubule bundling activity, 200 mM NaCl was added to the
reaction to detach the fusion protein from the microtubule
bundles. Two hours after NaCl treatment, the large aggregated
bundles disassembled completely into single microtubules (see
Supplemental Figure 13F online). In addition to confocal microscopy, electron microscopy was performed with negative staining
to confirm that, in the presence of heat-denatured FH1FH2,
microtubules were present as separate individual filaments (see
Supplemental Figure 13D online), and, in the presence of active
FH1FH2, the individual microtubules were tightly bunched along
their entire lengths to form microtubule bundles (see Supplemental Figure 13E online). In addition to these results, FH1FH2
was also shown to stabilize microtubules against cold-induced
disassembly (see Supplemental Figure 14 online).
Similar experiments were performed with Alexa 488-phallodinlabeled microfilaments, which, when incubated with 1 mM active
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Figure 5. Microspore Formation Is Defective in afh14 Arabidopsis Plants.
(A) Pollen counts from afh14-1, afh14-2, and wild-type anthers. Both mutant Arabidopsis plants produced significantly less pollen than did wild-type
plants (t test, P < 0.05). Error bars indicate SE (n > 100). Results include data from three independent experiments.
(B) A representative tetrad from a wild-type anther. The arrow indicates the swollen domain.
(C) to (E) Representative dyad (C), triad (D), and polyad (E) from the afh14-1 line.
(F) Representative irregular tetrahedral arrangement of afh14-1 microspores. Bar = 5 mm.
FH1FH2, appeared to become bundled (see Supplemental Figure 13H online). Similar to observations in microtubules, dot-like
structures corresponding to active FH1FH2 staining were visualized alongside the filaments (see Supplemental Figure 13I
online). Electron microscopy confirmed that individual microfilaments were bundled together in the presence of active FH1FH2
(see Supplemental Figure 13K online). By contrast, in the absence of the recombinant protein or in the presence of denatured
recombinant protein, only individual microfilaments were observed (see Supplemental Figures 13G and 13J online). Also
similar to microtubules, treatment with 200 mM NaCl promoted
disassembly of the microfilament bundles (see Supplemental
Figure 13L online). The observed bundle formation and association of FH1FH2 were not due to nonspecific antibody binding,
as microtubules and microfilaments stained with the fluorescently labeled secondary antibodies alone failed to exhibit dotlike structures along the length of the polymers (data not shown).
The AFH14 FH1FH2 Domain Bundles and Cross-Links
Microtubules with Microfilaments
To determine whether the AFH14 FH1FH2 domain could interact with microtubules and microfilaments concomitantly, binding experiments were performed using 0.5 mM preformed
microtubules, microfilaments, and 100 nM active FH1FH2. As
shown in Figure 9A, microfilaments assembled into bundles in
the presence of FH1FH2. When preformed taxol-stabilized
microtubules were added to reactions containing microfilaments and FH1FH2, microtubule bundles began to form after 5
min and, at the same time, some microfilament bundles became looser (Figure 9B). After 40 min and 4 h incubations,
increased numbers of microtubule bundles were observed,
whereas microfilament bundles completely disassembled into
individual filaments (Figures 9C and 9D). These observations
suggest that the microtubules induce detachment of FH1FH2
from the microfilament bundles.
In contrast with microfilament disassembly, when preformed
microfilaments were added to reactions containing bundled
microtubules and FH1FH2 (Figure 9E), the microtubule bundles
remained intact, and microfilament bundling was not observed
after 5 min, 40 min, or 4 h incubations (Figures 9F to 9H). This
result suggests that microfilaments cannot strip FH1FH2 from
microtubule bundles. In addition, when preformed microtubules
and microfilaments were mixed together in the absence of
FH1FH2, both cytoskeletal polymers remained as individual
filaments scattered randomly throughout the solution (Figure
9I). However, addition of FH1FH2 promoted microtubule bundling, but not microfilament bundling (Figure 9J). These results
indicate that the FH1FH2 protein preferentially bundles microtubules over microfilaments.
Interestingly, when higher concentrations of FH1FH2 were
added to reaction mixtures (up to 1 mM), both microtubules and
microfilaments formed bundles within 30 min (Figure 9K), and in
some cases, the two types of bundles colocalized (Figure 9L).
When an excess of anti-AFH14 antibodies was added to these
reactions, nearly all of the bundles disassembled into individual
filaments (Figure 9M), indicating that a dynamic exchange may
exist between free FH1FH2 in the solution and bound FH1FH2 on
microtubule and microfilament bundles. Negative staining electron microscopy was used to confirm the microtubule and
microfilament bundling and cross-linking activities of FH1FH2
(Figures 9O to 9R). The statistic analysis of the thickness of the
filaments and filament bundles corresponding to the result of
negative staining electron microscopy are shown in Supplemental Figure 15 online.
Similar experiments were also performed to test the activity of
the FH2 domain alone in binding or bundling microtubules and
microfilaments. Similar to the FH1FH2 recombinant protein,
treatment with 1 mM active FH2 resulted in binding to microtubules and microfilaments, decoration of filament bundles with
FH2 immunoreactivity, and microtubule and microfilament bundling (see Supplemental Figure 16 online).
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DISCUSSION
AFH14 Serves as a Linker Protein Coordinating Microtubule
and Microfilament Assembly in Plant Cells
Formins have been primarily shown to associate with actin
and regulate actin dynamics by nucleating, capping, severing,
elongating, and bundling microfilaments (Ingouff et al., 2005;
Michelot et al., 2005; Yi et al., 2005; Vidali et al., 2009; Ye et al.,
2009). Recently, various studies have suggested that formins can
also directly regulate microtubule organization and stability in
animal (Wen et al., 2004; Yasuda et al., 2004) and yeast cells
(Delgehyr et al., 2008). However, all formins identified to date in
plant cells have been characterized as microfilament regulators
(Blanchoin and Staiger, 2008), with the exception of the most
recently reported formin, AFH4, which can bind microtubules in
addition to microfilaments (Deeks et al., 2010). In this study, we
found that a type II formin from Arabidopsis, AFH14, appears to
play essential roles in regulation of both microtubules and
microfilaments. First, in vitro experiments revealed that the
FH1FH2 fragment of AFH14 binds to purified microtubules and
microfilaments directly and induces the formation of microtubule
or microfilament bundles. Additional in vitro experiments conducted in the presence of both microtubules and microfilaments
indicate that FH1FH2 binds preferentially to microtubules. This
result differs from the function of the well-characterized animal
formin, mDia2, which has a much higher in vitro binding affinity
for microfilaments than for microtubules (Bartolini et al., 2008).
mDia2-mediated promotion of microfilament polymerization and
stabilization of microtubules are functionally separate events,
and stable dimerization of mDia2 is not necessary to generate
stable Glu microtubules (Bartolini et al., 2008). With respect to
AFH4, it has been shown that the GOE domain located between
the transmembrane and the FH1 domain binds directly to microtubules (Deeks et al., 2010). By contrast, our results show that
the FH2 domain of AFH14 is critical for microtubule and microfilament binding. Although we have not directly analyzed the
requirement for dimerization of AFH14 in bundling of microtubules or microfilaments, our results showed that microtubules
compete with microfilaments to bind FH1FH2 and that an excess
of FH1FH2 promotes bundling of microtubules with microfilaments. Combining these observations with the fact that FH1FH2
has a highly conserved domain structure and has been suggested to function as a tethered dimer (Kovar, 2006; Michelot
et al., 2006), we propose that FH1FH2 or FH2 dimers of AFH14
Figure 6. Localization of AFH14 in Arabidopsis PMCs during Meiosis.
Localization of AFH14 was assessed by confocal microscopy in wildtype PMCs during prophase I (A), metaphase I (B), anaphase I (C),
telophase I (D), metaphase II (E), early cytokinesis (F), advanced cytokinesis (G), and the tetrad (H). AFH14 was dispersed throughout the
cytoplasm in prophase I (A), telophase I (D), early cytokinesis (F), and
in the tetrad (H). The RMS was present around the nuclei in these
cells. AFH14 colocalized with spindle microtubules in metaphase I (B)
and II (E). AFH14 colocalized with the phragmoplast microtubules in
anaphase I (C) and in advanced cytokinesis (G). Images were derived
from stacks of three or four sections containing the appropriate information. Bar = 5 mm.
A Plant MT and MF Interacting Formin
Figure 7. Arrangement of Microtubule Arrays in Meiosis Is Affected in
afh14 Male Meiocytes.
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Increasing evidence has shown that interactions between the
microtubule and microfilament cytoskeletons are common phenomena in plant cells. Coordination of microtubules and microfilaments in the cortical cytoplasm plays a key role during
directional expansion of cells as well as during positioning of
the plane of cell division. Microtubules and microfilaments
cooperate during cell division to coordinate the mitotic and
cytokinetic apparatus (Panteris and Galatis, 2005; Yasuda
et al., 2005; Collings, 2008; Petrášek and Schwarzerová, 2009).
However, little information is available at the protein level to
understand the mechanisms by which microtubules and microfilaments interact. The cotton calponin homology domain
(KCHs)–containing kinesins, Gh-KCH1 and Gh-KCH2, which
have been shown to bind both microtubules and microfilaments
in vitro, colocalize with microtubules and microfilaments in the
cortical cytoplasm and in the midzone of the phragmoplast in
dividing root tip cells (Xu et al., 2009), and the cytoskeletal dual
binding activity of Gh-KCH2 has been demonstrated in Arabidopsis protoplasts. Os-KCH1, recently identified in rice (Oryza
sativa), is associated with cortical microtubules and microfilaments in vivo and in vitro. These kinesins may play a role in both
cell elongation and cell plate formation. Microtubule-associated
190-kD polypeptide (MAP190) from BY-2 cells, a protein that
cosediments with both microtubules and microfilaments in vitro,
has been shown to associate with the spindle and the phragmoplast during cell division (Igarashi et al., 2000; Hussey et al.,
2002), but the in vitro cross-linking ability of MAP190 is not clear.
Microtubule-associated protein SB401, which has been shown
to bind both microtubules and microfilaments in vitro, colocalizes
with cortical microtubules in pollen tubes (Huang et al., 2007).
Immunostaining was used to assess tubulin localization in wild-type ([A]
to [C] and [G] to [I]) and afh14-1 ([D] to [F] and [J] to [L]) male meiocytes
in metaphase I ([A] and [D]), telophase I ([B] and [E]), metaphase II ([C]
and [F]), early cytokinesis ([G] and [J]), advanced cytokinesis ([H] and
[K]), and in tetrads ([I] and [L]). In metaphase I, the spindle in afh14-1
plants (D) was smaller than that of wild-type plants (A). In telophase I, two
RMSs were present surrounding the nuclei in wild-type male meiocytes
(B), but the RMS configuration was skewed in afh14-1 male meiocytes
(E). In metaphase II, the spindles were perpendicular in wild-type plants
(C) and parallel in afh14-1 plants (F). In early cytokinesis, microtubules
radiate from the four nuclei in both wild-type (G) and afh14-1 plants (J). In
advanced cytokinesis, phragmoplast microtubules were concentrated
between the two nuclei in wild-type male meiocytes but were absent in
the afh14-1 mutant (K). Four normal microspores were present in a tetrad
from wild-type plants (I). Two 2n microspores were present in afh14-1
dyad (L). Bar = 5 mm.
are probably involved in the bundling of the filaments. In addition,
for FH1FH2 and FH2, dots distributing along the bundled cytoskeletons seem uneven, which implies that AFH14 might form
different sizes of multimers like the mouse formin mDia1 does (Li
and Higgs, 2003) Furthermore, the microtubule bundling domain
may overlap with the domain required for microfilament bundling. Bundling of microtubules with microfilaments has previously been observed for Cappuccino, a related formin in
Drosophila melanogaster (Rosales-Nieves et al., 2006). These
results seem to indicate that formins are evolutionarily conserved
proteins that mediate interactions between microtubules and
microfilaments in eukaryotic cells.
Figure 8. FH1FH2 Recombinant Protein Binds Taxol-Stabilized Microtubules and Polymerized Microfilaments in Vitro.
(A) and (B) Taxol-stabilized microtubules (A) or polymerized microfilaments (B) were incubated with varying concentrations of recombinant
FH1FH2 protein. After centrifugation, supernatants (S) and pellets (P)
were analyzed by SDS-PAGE and Coomassie blue staining.
(C) Quantification of scanned SDS-PAGE gel from (A).
(D) Quantification of scanned SDS-PAGE gel from (B).
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The Plant Cell
Figure 9. FH1FH2 Recombinant Protein Preferentially Binds Microtubules and Links Microfilaments and Microtubules in Vitro.
(A) to (M) To compare the binding of FH1FH2 to microtubules and microfilaments, 0.5-mM preformed microtubules (red) and/or 0.5-mM microfilaments
(green) were incubated with 100 nM ([A] to [J]) or 1 mM ([K] to [M]) FH1FH2, and polymerization was assessed by confocal microscopy.
(A) Microfilaments exhibited considerable bundling 60 min after addition of FH1FH2.
(B) to (D) Addition of preformed taxol-stabilized rhodamine-conjugated microtubules to the reaction in (A) resulted in the formation of microtubule
bundles within 5 min (B). Microtubule bundling increased and microfilament bundling decreased at 40 min (C) and 4 h (D) after the addition of
polymerized microtubules.
(E) Microtubules exhibited significant bundling 60 min after the addition of FH1FH2.
(F) to (H) Microtubules remained bundled and microfilaments were randomly scattered throughout the solution 5 min (F), 40 min (G), and 4 h (H) after the
addition of preformed microfilaments to the reaction in (E).
(I) In the absence of FH1FH2, microfilaments and microtubules were randomly scattered throughout the solution.
(J) Microtubule bundles were present at 5 min, 30 min, and 4 h after the addition of FH1FH2 to the reaction in (I), but microfilaments were not bundled.
(K) and (L) Excess FH1FH2 protein (1 mM) induced bundling of both microtubules and microfilaments at 30 min (K) and 4 h (L) after addition.
(M) Addition of excess antibodies directed against FH1FH2 to the reaction described in (K) caused the majority of cytoskeletal bundles to dissociate
into single filaments.
(N) to (R) After incubation of 0.5 mM globular actin (G-actin) with preformed taxol-stabilized rhodamine-conjugated microtubules or microtubule bundles
(0.5 mM) for 20 min, microfilaments and microtubules were examined by electron microscopy. White arrows indicate microtubules, and black arrows
A Plant MT and MF Interacting Formin
Consistent with this observation, SB401 binds preferentially to
microtubules in the presence of both microtubules and microfilaments and has been shown to target specific organelles
to microtubules (Huang et al., 2007). However, it is not clear
whether this protein also binds microfilaments in vivo. Two
microtubule plus end–interacting proteins, EB1 and CLASP,
may be involved in root hair tip growth through interactions
with microfilaments. Despite the identification of these proteins
as in vitro cross-linkers of the microtubule and microfilament
networks, significant evidence is still needed to bridge the gap
between in vitro biochemical interactions and in vivo functional
cellular interactions. Intriguingly, AFH14 decorates almost all
mitotic and cytokinetic cytoskeletal apparatuses but does not
localize to cortical cytoskeletal networks in interphase cells,
which is different from the localization of PTEN-containing
formins in moss (Vidali et al., 2009). In combination with our
genetic, cellular, and biochemical studies, these results strongly
suggest that AFH14 may have a function distinct from cytoskeletal linkers involved in interactions between microtubules and
microfilaments in the cell cortex (Collings et al., 1998; Schwab
et al., 2003; Preuss et al., 2004; Frey et al., 2009).
AFH14 Plays an Important Role in Cell Division
During mitosis and cytokinesis, microtubule arrays form the
basis for mitotic and cytokinetic apparatuses, including the
preprophase band, the mitotic spindle, and the phragmoplast.
It is well accepted that the microfilament cytoskeleton is strongly
linked to these microtubule structures during cell division, although subtle differences in microfilament distribution have been
identified (Panteris, 2008). While some studies have shown that
microfilaments codistribute with microtubules in the preprophase band (Kakimoto and Shibaoka, 1987; Palevitz, 1987;
Traas et al., 1987; Zachariadis et al., 2001, 2003), others have
shown that cortical actin filaments do not always follow the
pattern of the preprophase microtubule band (Cho and Wick,
1990, 1991; Panteris et al., 1992, 2007; Collings and Wasteneys,
2005). In the mitotic spindle, microfilaments are usually present
(Schmit and Lambert, 1987; Seagull et al., 1987; Traas et al.,
1987; Lloyd and Traas, 1988; Panteris et al., 1992; Cleary, 2001;
Collings et al., 2001), but their absence from the spindle has also
been reported (Liu and Palevitz, 1992; Baluska and Hasenstein,
1997; Vitha et al., 2000; Voigt et al., 2005). It is generally accepted
that, in the phragmoplast, actin filaments lie parallel to microtubules (Panteris, 2008). This paradigm is consistent with a previous finding showing that microfilaments localize to the central
region of the phragmoplast in BY-2 cells (Yoneda et al., 2004).
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In agreement with previous results, our experiments in both
Arabidopsis and BY-2 cells showed that microfilaments form a
cage around spindle and that the length of phragmoplast microfilaments is shorter than that of microtubules. However, in
AFH14-overexpressing cells, the microfilaments and microtubules appeared to be similar in length and aligned evenly with
one another. By contrast, in cells with decreased AFH14 expression, the connections between microtubules and microfilaments
were disturbed at several different levels. These results indicate
that proper microfilament structure is important for stabilizing the
spindle apparatus. We hypothesize that downregulation of
AFH14 disturbs the interaction between microtubules and microfilaments, which initially causes microfilament instability and
is shortly followed by alterations in microtubule structures important for mitosis and cytokinesis. Consistent with this idea, in
animal cells, mDia2-mediated regulation of the microfilament
cytoskeleton has been suggested to be required for proper
distribution of stable microtubules (Bartolini and Gundersen,
2006).
The precise nature of microfilament distribution relative to
microtubule distribution has remained somewhat unclear due to
discrepancies in the observed distribution of microfilaments.
These discrepancies could be explained by differences in techniques used for visualization (Maupin and Pollard, 1986; Staiger
and Schliwa, 1987; Liu and Palevitz, 1992) and differences in
cell types (Derksen et al., 1986; Schmit and Lambert, 1987;
Haraguchi et al., 1997; Hasezawa et al., 1998; Silverman-Gavrila
and Forer, 2000). In addition, we propose that some of these
discrepancies may also be due to differences in the timing of the
images, as it is very difficult to obtain images at the exact same
time point due to the extreme dynamics of the two cytoskeletal
structures, particularly microfilaments. In this regard, functional
studies characterizing cross-linking proteins that coordinately
regulate the two cytoskeletal networks may help us to understand better the precise relationship between cytoskeletal microtubules and microfilaments.
Through analysis of two mutant Arabidopsis lines harboring
T-DNA insertions within AFH14, afh14-1, and afh14-2, we found
that AFH14 is involved in microspore generation. These mutant
plants exhibited a variety of meiosis-related phenotypes, including dyads, triads, polyads, and irregular tetrahedral tetrads with
abnormal microspore distributions. Dyads and triads may contribute primarily to the reduction of the total amount of pollen.
These phenotypes imply that AFH14 is involved in the regulation
of male meiotic division. Meiotic microtubule arrays have been
described in a number of different species. During premeiotic
interphase, microtubules form random networks throughout the
Figure 9. (continued).
indicate microfilaments. White lines and numbers indicate the diameter of filaments or filament bundles.
(N) Addition of G-actin to a mixture of microtubules and denatured FH1FH2 resulted in a random (not bundled) distribution of microfilaments and
microtubules.
(O) to (Q) Addition of G-actin to a solution containing microtubule bundles induced by 400 nM FH1FH2 resulted in elongation of microfilaments (O) or
microfilament bundles (P) from the microtubules.
(Q) Representative image of microfilament bundles arising from microtubule bundles.
(R) A representative image of a microfilament cross-linked with a microtubule.
(A) to (H) have the same magnification; bars = 5 mm. (I) to (M) have the same magnification; bars = 5 mm. Bars = 50 nm in (N) to (R).
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The Plant Cell
cytoplasm. In telophase I, two RMSs are formed around the
nuclei. In metaphase II, the spindles lie perpendicular to one
another, and in early cytokinesis, RMSs radiate from the four
nuclei. During advanced cytokinesis, microtubules are not only
radially distributed around the entire nucleus but also form
phragmoplast microtubules located between the two nuclei
(Hogan, 1987; Brown and Lemmon, 1988; Traas et al., 1989;
Staiger and Cande, 1991). We observed similar structures during
tetrad formation in wild-type plants. However, in afh14-1 plants,
abnormal microtubule structures were observed that included
skewed RMSs in telophase I, parallel spindles in metaphase II,
and the absence of phragmoplast microtubules in advanced
cytokinesis. Compared with the strong disturbance of mitosis
in suspension-cultured cells after downregulation of AFH14,
T-DNA insertion mutants show relatively weak deviations in
meiosis. The difference is probably due to gene redundancy or
off-target silencing in RNA interference experiments.
The best documented and described meiotic abnormalities
indicate that the RMS, the orientation of the spindle, and the
cytokinetic microtubule systems affect the distribution of microspore nuclei in tetrads (Brown and Lemmon, 1988; Bretagnolle
and Thompson, 1995; Magnard et al., 2001). For example, the
RMS determines spindle positioning, and the orientation and
position of the spindles reflect the positions of the microspore
nuclei in the meiotic coenocyte (Heslop-Harrison, 1971; Dover,
1972; Brown and Lemmon, 1988). A multiplanar second division
spindle in Lonicera japonica causes a tetrahedral arrangement
of nuclei, whereas a uniplanar division in Impatiens sultani results
in a tetragonal nuclear arrangement (Brown and Lemmon, 1988).
During advanced cytokinesis, the structure of the phragmoplast
microtubules maintains the microspore nuclear distance and
positioning (Hogan, 1987). Extensive studies have characterized the mechanism of dyad formation and have found that
parallel spindle orientation is the most common cause of
dyads (Bretagnolle and Thompson, 1995; Genualdo et al., 1998;
Consiglio et al., 2004; Jiang et al., 2009).
Microtubule–microfilament interactions have been proposed
to be indispensable for male meiotic division. In support of this
idea, in metaphase I and II, microfilaments codistribute with the
spindle microtubules. During the second division, microfilaments
intimately connect with the radiating microtubules and guide
outgrowth of the phragmoplast (Schmit and Lambert, 1987;
Traas et al., 1989; Staiger and Cande, 1991; Preuss et al., 2004;
Yasuda et al., 2005). Drug-based experiments have shown that
fragmentation of microfilaments leads to the disappearance of
spindles and phragmoplasts. In addition to the disappearance of
these microtubule-based structures, microfilaments normally
concentrated around the chromosomes are absent (Traas
et al., 1989). Microtubules also affect microfilament distribution,
for example, the maize (Zea mays) dv and ms17 mutants display
defects in microtubule distribution and a dramatic reorganization
of microfilaments (Staiger and Cande, 1991). In animals, the
mitotic microfilament cortex has been shown to guide spindle
orientation in response to extracellular stimuli (Kunda and Baum,
2009). Consistent with this observation, dyad production appears to be related to deviations in the spatial configurations
of microtubules and microfilaments in plants (Genualdo et al.,
1998). Furthermore, treatment with cytochalasin D, a potent
inhibitor of actin polymerization, often results in tetrads with
disoriented cell walls (Magnard et al., 2001).
In summary, coordinate regulation of microtubules and microfilaments is thought to be important for the formation of regular
microtubule configurations necessary for mitosis and meiosis.
Based on direct genetic and biochemical evidence, we have
demonstrated that AFH14 plays a key role in spatial regulation of
cell division by mediating interaction between microtubules and
microfilaments.
METHODS
Plant Materials, Growth Conditions, and Transformations
Arabidopsis thaliana afh14-1 (SALK_058886) and afh14-2 (SALK_038277)
lines were obtained from the Salk Institute for Biological Studies. Wildtype and mutant plants were of the Columbia-0 ecotype. Plants were
grown under similar conditions as described previously (Yi et al., 2005).
Arabidopsis thaliana suspension cultures and tobacco BY-2 (Nicotiana
tabacum Bright Yellow-2) cells were grown as described by Smertenko
et al. (2004). Agrobacterium tumefaciens (strain GV3101) was used for
transformation of Arabidopsis and BY-2 cells. Cells were transformed as
previously described (Yu et al., 2006). Exogenous gene expression in
transgenic cell lines was induced by the addition of 2 mM or 100 nM 17bestradiol (Sigma-Aldrich) to Arabidopsis or BY-2 culture medium, respectively. Mock induction treatments consisting of DMSO alone were
used as negative controls for all experiments.
Screening of T-DNA Insertion Mutants and RT-PCR Analysis
Genomic DNA extraction was performed using a plant genomic DNA kit
according to the manufacturer’s recommended protocol (Tiangen).
T-DNA insertions in afh14-1 and afh14-2 were detected by PCR using
the following primer sets: RP1, 59-TGTTGTTGTTGTCGTCTCAGG-39,
and LP1, 59-GCCTCCCAAAGAATATCCAAG-39; and RP2, 59-CTCGATGTCCTTTTGAGCAAG-39, and LP2, 59-AATCGCAATGCTTATTCG
AAG-39). LP1, LP2, and LBa1 (http://signal.salk.edu/tdnaprimers.2.html)
were used to detect the wild-type gene.
For RT-PCR, total RNA was isolated from the 10-d-old wild-type,
afh14-1, and afh14-2 plants using a plant RNA extraction kit (Autolab),
and 2 mg RNA was used for reverse transcription using a commercial kit
(Invitrogen). Primers for PCR were as follows: afh14-1 (forward, 59-TGAGCGTAGAACTCTTGAGATTGTGC-39, reverse, 59-AAATAAAAACTCCCTTTGTGCCGAA-39), afh14-2 (forward, 59-AATAAACGCTGCTACTAA
AGAGGTG-39, reverse, 59-CTTCTGGGTTCTATGTAAGTTCAAAT-39),
and ACTIN2 (forward, 59-AGGGCTGTTTTTCCCAGTGTTG-39, reverse,
59-GTGCTGTGATTTCTTTG CTCATACG-39).
Cloning, Recombinant Protein Expression, and Construct Design
PCR products were amplified using the primers shown below and were
digested with the indicated restriction enzymes prior to introduction into
different expression vectors. The AFH14 cDNA was amplified using the
following primers: 59-GGCG CGCCATGTCTTTGTTAAGTAGATTCTTTTACA-39 (forward) and 59-GGTACCTGTTCTATGTCTATGGATCTGCTG-39 (reverse). AscI and KpnI (underlined) were used for digestions.
GFP was amplified from pCAMBIA1300-GFP-Fabd2 using 59-GGTACCATGGGTAAAGGAGAAGAACTTTT-39 or 59-CTCGAGATGGGTAAAGGAGAAGAACTTTT-39 (forward; for the fusion or GFP alone, respectively)
and 59-ACTAGTTCACTTTTTGTATAGTTCATCCAT-39 (reverse), and KpnI
or XhoI and SpeI were used for digestion, respectively. The ligated
AFH14:GFP and GFP fragments were individually inserted into the pER8
A Plant MT and MF Interacting Formin
13 of 17
plant transformation vector, which contains the G10-90 promoter (Zuo
et al., 2000). AFH14 promoter was amplified from Arabidopsis genomic
DNA using the following primers: 59-GAGCTCGGTGAAATCCTAGAATTGAGGGTT-39 (forward) and 59-CCATGGTGGAAGAACCTTGTTACAATAAGAAGACGG-39 (reverse). SacI and NcoI were used for digestion. The
AFH14 promoter fragment was then inserted into the pCAMBIA1305.1
vector (CAMBIA), which contains the GUS gene. The FH1FH2 and
FH2 fragments were amplified from the AFH14 cDNA with the following
primers 59-GGTACCCCATCAGATCCACCATCTTCT-39 and 59-GGTACCATGGCAGGAAGG GGGAGAG-39 (forward; respectively) and 59-GCGGCCGCTGTTCTATGTCTATGGATCTGCTG-39 (reverse). KpnI and NotI
were used to digest the fragments, which were then inserted individually into pET30-a expression vectors (Novagen) for protein expression in the Escherichia coli BL21(DE3) strain. Recombinant protein
was affinity purified as previously described (Sambrook and Russell,
2001).
An artificial microRNA construct was designed according to published
protocols (Schwab et al., 2006). The amiR-AFH14 construct included
the miR319a precursor as a backbone (http://www.weigelworld.org), a
designed duplex 21-nucleotide sequence (miRNA, 59-TATTACGTCACATTCTGCGTG-39, and miRNA*, 59-CAAGCAGAATGTGTCGTAATT-39)
unique to Arabidopsis AFH14 cDNA, and XhoI and SpeI restriction sites at
the 59 and 39 ends, respectively. After digestion with XhoI and SpeI, the
amiR-AFH14 fragment was introduced into the pX7 vector (Guo et al.,
2003). All constructed plasmids were confirmed by DNA sequencing.
To assess microtubule binding activity of FH1FH2, 2 mM preformed
taxol-stabilized microtubules were mixed with 2, 4, 6, 8, 10, 12, 14, or 18
mM FH1FH2 recombinant protein in 100 mL PEMT buffer for 30 min at
room temperature. The samples were centrifuged at 25,000g for 30 min at
258C. A volume of 480 or 304 mL 13 loading buffer was added to the
resulting pellet or to 80 mL of supernatant, respectively.
For both assays, all samples were resolved on 12% SDS-PAGE
gels. Gels were stained with Coomassie Brilliant Blue R 250 (Dingguo).
FH1FH2-bound microtubules and microfilaments were quantified by
scanning gels with UMAX PowerLook 2100XL (Umax). Protein concentrations were measured three times using a 318C microplate reader
(Sanke).
Antibody Preparation and Immunoblotting
Staining and Immunolabeling for Fluorescence Microscopy
The FH1FH2 fusion protein was used as an antigen to raise polyclonal
antibodies in mouse, which was used to assess FH1FH2 chemical activity
on microtubules and microfilaments for in vitro experiments. To assess
endogenous AFH14 localization in Arabidopsis cells, a monoclonal antibody (AbMART) was raised in mouse using a specific fragment consisting
of AFH14 amino acids 340 to 502.
Total protein extraction from 10-d-old Arabidopsis seedlings and
subsequent immunoblotting with ECL Plus Western Blotting Detection
reagents (GE Healthcare) was performed as previously described (Fan
et al., 2004; Hiwatashi et al., 2008).
For immunolabeling, Arabidopsis and BY-2 cells were fixed in 4% (w/v)
paraformaldehyde in PEM buffer (50 mM PIPES, 5 mM EGTA, 5 mM
MgSO4, 0.1 M mannitol, pH 6.9) for 30 min at room temperature. The cells
were then rinsed three times with PEM buffer and treated with a solution
of 0.5% pectolyase and 1% cellulase R-10 in PEM buffer for 10 min at
room temperature. Afterward, the cells were incubated for 40 min in 3%
(w/v) BSA in PEM buffer, followed by incubation with primary antibodies
overnight at 48C in a dark, moist chamber. Primary antibodies used for
immunostaining included mouse AFH14 monoclonal antibodies (1:100
dilution) and rat monoclonal anti-a-tubulin YL1/2 antibodies (AbD Serotec;
1:200 dilution). The specimens were then washed three times for
10 min each in PEM buffer. For double immunolabeling, anti-mouse
tetramethyl rhodamine isothiocyanate-conjugated antibodies (crossadsorbed against rat and other IgGs; catalog number 715-025-151)
and anti-rat fluorescent isothiocyanate-conjugated antibodies (crossadsorbed against mouse and other IgGs; catalog number 112-095-167)
were used for secondary antibody labeling. Finally, the cells were washed
four times with PEM buffer and stained with DAPI (1 mg/mL) or 110 nM
rhodamine-phalloidin (Invitrogen) if needed for 10 min prior to visualization by confocal laser scanning microscopy. As a negative control, each
sample was incubated with only one of the primary antibodies, followed
by incubation with a mixture of both secondary antibodies used in the
experiment. No detectable signal was recorded in fluorescent channels
that did not correspond to the primary antibody.
Inflorescences were fixed in Carnoy’s solution (6:3:1, ethanol:chloroform:glacial acetic acid) at room temperature for at least 24 h. To observe
the tetrads, the inflorescences were stained in Carbol fuchsin for 48 h. An
anther was then placed on a slide and gently rubbed between the cover
glass and slide under the light microscope. This process resulted in the
release of tetrads from the anther.
Preparation of PMCs for immunofluorescence microscopy was performed by as previously described (Jiang et al., 2009). Briefly, inflorescences were fixed in methanol:acetone (4:1, v/v) for 3 h, waving at 100g,
and washed two times with PEM buffer. Anthers were then gently flattened
on poly-L-lysine–coated slides, resulting in the release of PMCs from
the anthers. Subsequently, slides were dried for 10 min. A thin layer of
Preparation and Polymerization of Actin and Tubulin
Rabbit muscle actin was purified according to published methods
(Pardee and Spudich, 1982). Actin was centrifuged at 100,000g for 30
min using a TLA-110 rotor (Beckman) at 48C. Actin (10 mM) was polymerized in 13F buffer (5 mM Tris-HCl, 0.5 mM DTT, 0.5 mM ATP, 50 mM
KCl, and 1 mM MgCl2, pH 7.0) at 48C for 16 h or at 378C for 2 h in the
presence of 100 nM Alexa-488 phalloidin (Molecular Probes).
Tubulin and NHS-rhodamine–labeled tubulin were centrifuged at
30,000g for 30 min at 48C. Tubulin was polymerized to produce taxolstabilized microtubules in PEMT buffer (100 mM PIPES, 2 mM EGTA,
1 mM MgCl2, and 0.1, 1, or 10 mM taxol, pH 6.9). The resulting mixture was
centrifuged at 12,000g at 258C for 10 min. Pellets were washed with
PEMT buffer containing 10 mM taxol and resuspended gently in PEMT
buffer containing 10 mM taxol to a final concentration of 10 mM tubulin.
Microtubule and Microfilament Cosedimentation Assays
FH1FH2 protein was centrifuged at 100,000g at 48C for 1 h. To analyze
microfilament binding activity of FH1FH2, 5 mM microfilaments and 0.1,
0.2, 0.3, 0.4, 0.5, 0.6, 0.8, or 1.0 mM FH1FH2 recombinant protein in a
100-mL total volume were incubated for 30 min at 378C. The samples were
centrifuged at 100,000g for 30 min. The resulting pellets were then
resuspended in 120 mL 13 loading buffer, and 16 mL 53 loading buffer
was added to 80 mL of the resulting supernatant.
Assessment of Microtubule and Microfilament Bundling Activity
Taxol-stabilized microtubules and microfilaments prepared as described
above were incubated without or with 1 mM active or heat-denatured
FH1FH2 or FH2 at room temperature in a 10 mL total volume for 1 h. To
determine whether FH1FH2 changed the stability of microtubules in vitro,
solutions containing microtubules and active or heat-denatured FH1FH2
were incubated at 48C for 30 min or 2 h. A 1-mL sample was then placed
on a slide and visualized by confocal microscopy or fluorescence microscopy (Zeiss). Microtubules or microfilaments were also observed by
negative staining under an electron microscope (Hitachi H600).
14 of 17
The Plant Cell
agarose/gelatin (0.75% low melting agarose, 0.75% gelatin, and 0.3%
sucrose) was spread on the male meiocytes. PMCs were then soaked in
PEM buffer containing 2% Drilease (Dingguo) for 1 h at 378C in the
absence of light. Drilease was removed by washing in PEM buffer. Slides
were then incubated in 1% Triton X-100/10% DMSO for 1 h at room
temperature, followed by three washes. Afterward, the slides were
incubated with AFH14 antibody and anti-a-tubulin antibodies (1:100
dilution) overnight, washed, and incubated with the appropriate secondary antibody (1:200 dilution) for 2 h. Finally, nuclei were stained by
incubation with DAPI for 40 min as described above.
Drug Treatments
Cytoskeletal inhibitors used at the following concentrations: 10 mM
oryzalin (Sigma-Aldrich) and 200 nM LatB (Invitrogen). After 30 h of
induction, cells were incubated in fresh medium containing oryzalin or
LatB for 15 or 50 min, respectively. Cells were then fixed and labeled with
rhodamine-phalloidin or anti-a-tubulin as described above.
Microscopy
Confocal microscopy and image acquisition were performed using a
Zeiss LSM 510 META confocal microscope. Zeiss 363 oil objectives
(Plan-Apochromat; numerical aperture of 1.4) were used for visualization.
Serial confocal optical sections were taken at a step size of 0.5 to 1.0 mm.
Images are presented as either single sections or stacks of neighboring
sections. All images were processed using Adobe Photoshop 7.0 software.
Supplemental Figure 11. Microtubule Array Arrangement Is Defective in afh14-2 Plants.
Supplemental Figure 12. The FH1FH2 Recombinant Protein Nucleates Microfilaments in Vitro.
Supplemental Figure 13. Microtubule and Microfilament Organization in the Presence of FH1FH2.
Supplemental Figure 14. FH1FH2 Inhibits Cold-Induced Disassembly of Microtubules in Vitro.
Supplemental Figure 15. The Thickness of Filaments and Filament
Bundles.
Supplemental Figure 16. Microtubule and Microfilament Organization in the Presence of the FH2 Recombinant Protein.
ACKNOWLEDGMENTS
We thank Xiaohua Li (Baylor Institute for Immunology Research) for
critical readings. We also thank Bo Liu (University of California at Davis)
for detailed discussions, Chuanmao Zhang (Peking University) for helping with electron microscopy images, and Liwen Jiang (Chinese University of Hong Kong) for providing Arabidopsis suspension cells. This
work was supported by the National Basic Research Program of China
(2007CB108700 and 2006CB100100) and by the National Natural Science Foundation of China (30970174, 30325005, and 30870211).
Received March 24, 2010; revised July 17, 2010; accepted July 28, 2010;
published August 13, 2010.
Accession Numbers
Sequence data from this article can be found in the Arabidopsis Genome Initiative or GenBank/EMBL databases under accession numbers
At1g31810/HM016081 (AFH14).
Supplemental Data
The following materials are available in the online version of this article.
Supplemental Figure 1. Predicted Domain Organization of the
AFH14 Protein.
Supplemental Figure 2. Differences between the Amplified and
Predicted AFH14 Coding Sequences (CDSs).
Supplemental Figure 3. FH1FH2 and AFH14 Are Specifically Recognized by Polyclonal and Monoclonal AFH14 Antibody, Respectively.
Supplemental Figure 4. Oryzalin Depolymerizes Microtubules in
Uninduced Control BY-2.
Supplemental Figure 5. AFH14-GFP Localization Is Irregular or
Dispersed in BY-2 Cells Treated with Oryzalin.
Supplemental Figure 6. Overexpression of AFH14-GFP Increases
the Stability of Mitotic Microfilaments.
Supplemental Figure 7. Phragmoplast Microfilaments Codistribute
with Microtubules in Uninduced Cells.
Supplemental Figure 8. Analysis of the AFH14 Expression Pattern in
Arabidopsis Tissues and Organs.
Supplemental Figure 9. Schematic Showing the AFH14 T-DNA
Insertion Locus.
Supplemental Figure 10. Microspore Formation Is Defective in
afh14-2 Arabidopsis Plants.
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The Type II Arabidopsis Formin14 Interacts with Microtubules and Microfilaments to Regulate
Cell Division
Yanhua Li, Yuan Shen, Chao Cai, Chenchun Zhong, Lei Zhu, Ming Yuan and Haiyun Ren
Plant Cell; originally published online August 13, 2010;
DOI 10.1105/tpc.110.075507
This information is current as of June 17, 2017
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