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First published online April 10, 2009
doi: 10.1242/10.1242/dev.030098
í Friml3
1 Department of Biological Sciences, Graduate School of Science, University of
Tokyo, Bunkyo-ku, Tokyo 113-0033, Japan.
2 Molecular Membrane Biology Laboratory, RIKEN Advanced Science Institute, Wako,
Saitama 351-0198, Japan.
3 Department of Plant Systems Biology, VIB and Department of Plant Biotechnology
and Genetics, Ghent University, Technologiepark 927, 9052 Gent, Belgium.
Authors for correspondence (e-mails:
sanar{at}psb.vib-ugent.be;
fukuda{at}biol.s.u-tokyo.ac.jp)
Accepted 27 February 2009
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SUMMARY |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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Key words: Arabidopsis, VAN3, Vein, ARF-GAP, CVP2, VAB
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INTRODUCTION |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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; Moss and Vaughan,
1995; Moss and Vaughan,
1998). ARF proteins function through a cycle of GTP binding and
GTP hydrolysis, which leads to the GTP-bound active form and the GDP-bound
inactive form of the protein. This cycle is regulated by guanine nucleotide
exchange factors (GEFs) and GTPase-activating proteins (GAPs).
The ARF-GAP proteins have been categorized into three groups: ARF-GAP1
type, Git type and AZAP type in mammals
(Randazzo and Hirsch, 2004).
AZAPs are divided further into four subgroups based on their domain
structures: ACAPs, ASAPs, AGAPs and ARAPs. Because a single ARF protein
functions at multiple organelles, the intracellular site of ARF-GAP protein
action is important for conferring the site specificity of ARF function.
Whereas all ARF-GAP1 type proteins localize to the cis-Golgi, the AZAP-type
proteins function in the post-Golgi transport pathways. For example, ACAPs and
ASAPs localize to the plasma membrane (PM)/endosome
(Jackson et al., 2000), AGAPs
localize to the lysosome/endosome (Nie et
al., 2003), and ARAPs localize to the trans-Golgi/TGN/PM
(Miura et al., 2002). These
results suggest that the AZAP type ARF-GAP proteins are key players in the
regulation of membrane traffic in the post-Golgi transport pathways. Detailed
cell biological analysis has been performed in animal cells
(Jackson et al., 2000;
Kam et al., 2000;
Randazzo et al., 2000;
Miura et al., 2002;
Li et al., 2005) and these
results provide a theoretical framework for understanding the molecular
characterization of AZAPs in cells.
However, most of these data come from studies of cultured single animal cells. The fact that AZAPs exist only in multicellular organisms strongly suggests that AZAPs may have a variety of functions, depending on the different cell types, in multicellular organisms. Therefore, understanding the divergent roles of AZAPs in individual organisms is important. Genetic analysis with mutants of AZAPs and AZAP-related factors is one complementary approach to cultured cell-based analysis. However, no studies have reported on mutants of AZAPs in animals.
The genome of a model plant Arabidopsis thaliana encodes 15
ARF-GAPs, which can be classified into two groups: ARF-GAP1 and AZAP. Although
animal AZAPs are differentiated functionally into ACAPs, ASAPs, AGAPs and
ARAPs, Arabidopsis has no AZAPs other than the ACAPs, which comprise
four members. We and another group have found that mutants defective in an
Arabidopsis ACAP gene, VAN3/SFC, have disconnected formation
of the vascular bundles in leaves (Koizumi
et al., 2005; Sieburth et al.,
2006). In addition, quadruple mutants of all plant ACAPs showed an
enhanced vascular phenotype of VAN3/SFC
(Sieburth et al., 2006). These
observations indicate that plant ACAPs function in the continuous formation of
vascular bundles.
Plant vascular bundles comprise xylem and phloem tissues, which are
generated from the procambim. In leaves, the vascular bundle, called the vein,
forms a species-dependent complex network. The spatiotemporal regulation of
the vascular pattern has been considered an excellent model for studying the
plant developmental program. Some polarly distributed proteins were found
recently to function in generating continuous vascular patterns. For example,
xylogen is a glycoprotein that is secreted polarly from differentiating
vascular cells towards adjacent undifferentiated cells to induce vascular
differentiation and accomplish vascular connection
(Motose et al., 2004). Polar
auxin transport is a well-known crucial factor for continuous pattern
formation of veins (Przemeck et al.,
1996; Hardtke and Berleth,
1998; Mattsson et al.,
1999). Many reports show a close correlation between vascular
patterning and the distribution of cells with asymmetrically localized PIN
proteins, which are auxin efflux carriers
(Paciorek et al., 2005;
Wisniewska et al., 2006). PIN
proteins cycle between the endosome and the PM, and, interestingly, GNOM
ARF-GEF functions in the polar exocytosis of PIN proteins
(Geldner et al., 2001;
Geldner et al., 2003). These
results suggest that polar vesicle trafficking plays a pivotal role in the
coordinated vascular pattern formation.
Because the phenotype of the van3 mutant is largely restricted to
severe vascular discontinuity, it is thought that VAN3 is a pivotal factor
responsible for the vesicle transport governing the vascular continuity.
Genetic interactions between van3/sfc and gnom, and between
van3/sfc and other auxin-signaling mutants, suggest that VAN3/SFC
functions in auxin transport or auxin signaling or both
(Koizumi et al., 2005;
Sieburth et al., 2006).
Scarpella et al. (Scarpella et al.,
2006) reported that subcellular PIN1 polarity in pre-procambial
cells, the precursors of procambial cells, determines the route of auxin
transport and the subsequent pattern of veins. In the van3 mutant,
however, PIN1 polarity is established but is not maintained in pre-procambial
cells, resulting in discontinuous formation of procambial cells. These results
suggest that VAN3/SFC functions in vascular continuity by regulating the polar
localization of proteins such as PIN1. However, the molecular mechanism by
which VAN3 regulates the polar transport of vesicles is understood poorly. We
have suggested that VAN3 is localized to the TGN
(Koizumi et al., 2005), which
differs from the subcellular localization of animal ACAPs. This difference
does not allow us to interpret VAN3 function as being simply analogous to ACAP
functions in animals.
One advantage of using Arabidopsis as the model material is the
ease of genetic analysis. Several mutants are defective in vascular continuity
[e.g. mp, cvp and fkd
(Hardtke and Berleth, 1998;
Steynen and Schultz, 2003;
Carland and Nelson, 2004)]. To
understand the molecular function of VAN3 in vascular continuity, we searched
for factors that affect VAN3 function by analyzing the genetic interactions
between van3 and mutants showing the van3-like phenotype.
Screening proteins that interact with VAN3 has identified a dynamin, DRP1A,
that functions in vascular continuity in association with VAN3
(Sawa et al., 2005).
Therefore, we also rescreened VAN3-interactive proteins using the yeast
two-hybrid system. The analysis of factors obtained from these approaches
suggests that a specific phosphoinositide(s) plays a role in subcellular
protein targeting and enhancing of VAN3 ARF-GAP activity: this information
provides new insights into the upstream regulation of ACAPs.
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MATERIALS AND METHODS |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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Establishment of the transgenic plants expressing fluorescence tagged full length and mutated forms of VAN3
The 3x[(Gly)3Ser]-VENUS sequence was amplified by PCR from
the pCS2 plasmid (Nagai et al.,
2002). The VAN3-VENUS construct was made using a TT-PCR
method (Tian et al., 2004)
with the 3x[(Gly)3Ser]-VENUS sequence and the 8.6 kb
XbaI-SpeI genomic fragment of VAN3 that contained
the 1.1 kb upstream and the 1.1 kb downstream, as templates. The VAN3-VENUS
fragment was cloned into pCAMBIA1300 at XbaI and SpeI sites,
resulting in VAN3-VENUS translational fusion. For construction of VAN3
derivatives, essentially the same procedure was used. For subcellular
localization analysis of VAN3, transgenic plants harboring VAN3-VENUS were
crossed with lines expressing ST-sGFP, sGFP-SYP41 and sGFP-ARA7/RAB-F2b
(Wee et al., 1998;
Uemura et al., 2004;
Goh et al., 2007), which had
been developed by ourselves. For observing the signals of sGFP and VENUS,
Zeiss LSM 510 META system was used. sGFP
(Niwa et al., 1999) and VENUS
were excited with 488 nm line of Argon laser and emitted fluorescence was
detected with META detector from 505 to 580 nm. The reference for sGFP or
VENUS was obtained from cells expressing only the sGFP or VENUS fused protein.
We also obtained the reference for autofluoresence (background) from
non-transformed cells. Based on these references, sGFP and Venus signals were
separated by Emission Fingerprinting.
Agrobacterium-mediated transient expression of genes
The genomic fragment of VAB and the cDNA fragment of CVP2 were
cloned into a binary vector containing YFP: pH35GY
(Kubo et al., 2005). VAN3cDNA
was cloned into a binary vector containing GFP: pGWB2
(Nakagawa et al., 2007). These
DNA fragments were introduced into leaves of Nicotiana benthamiana
with Agrobacterium tumefaciens GV3101 (pMP90). Segregation of the
fluorescent signals between sGFP and VENUS was performed as described
above.
Whole-mount immunolocalization
Whole-mount immunolocalization of PIN1 was performed with anti-PIN1
(1:1000) and Cy3-conjugated anti-rabbit secondary antibodies (1:600) as
described by Friml et al. (Friml et al.,
2003).
Split-YFP assay
cDNA fragments of VAN3, VAL3 and VAB were used for the
split-YFP assay. The YFP was split between residues 154 and 155 into nYFP and
cYFP parts (Bracha-Drori et al.,
2004; Walter et al.,
2004). The chimeric gene was subcloned under the control of 35S
promoter and the NOS terminator. Co-introduction of VAN3-nYFP or
VAL3-nYFP with VAB-cYFP into protoplasts from
Arabidopsis cultured cells was performed as described previously
(Koizumi et al., 2005).
Yeast two-hybrid assay
The yeast two-hybrid assay was carried out as described previously
(Sawa et al., 2005). The cDNAs
for VAN3 and VAB were subcloned into pENTR D-topo
(Invitrogen) and then moved into pAD-GAL4-GWRFC and pBD-GAL4-GWRFC
(Sakai et al., 2008). Plasmids
containing VAN3 and VAB were introduced into AH109 strain
(Clontech).
Expression and purification of GST-tagged VAN3 and myristoylated AtARF1
DNA for VAN3, VAN3G321E and the GAP-ANK domain of VAN3 was
amplified by RT-PCR. These DNA fragments were cloned into the pGEX6p-1 vector
and expressed in Escherichia coli BL21 (DE3). The cells expressing
the fusion proteins were resuspended in lysis buffer [50 mM Tris (pH 8.0), 150
mM NaCl, 0.1% mercaptoethanol and protease inhibitor cocktail (GE Healthcare)]
and sonicated. Cell extracts were centrifuged at 10,000 g for
30 minutes, and the supernatant was loaded onto a glutathione-Sepharose 4B
column (GE Healthcare). After washing with 10 column volumes of buffer [50 mM
Tris (pH 8.0), 500 mM NaCl and 0.1% mercaptoethanol], GST-VAN3 was eluted with
elution buffer [20 mM reduced glutathione, 50 mM Tris (pH 8.0), 150 mM NaCl,
and 0.1% mercaptoethanol]. Myristoylated AtARF1 was purified from
Escherichia coli BL21 (DE3) that was co-transfected with expression
vectors for AtARF1 and yeast N-myristoyltransferase, as described
previously (Koizumi et al.,
2005).
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Primer information
Primer details can be provided on request.
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RESULTS |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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;
Koizumi et al., 2000;
Koizumi et al., 2005;
Sieburth et al., 2006). Many
of these alleles carry a nonsense or frameshift mutation before the catalytic
ARF-GAP domain, whereas a few missense mutations such as sfc-1 and
sfc-6 are mapped onto the ARF-GAP domain. However, no missense
mutation in other domains of VAN3 was identified
(Fig. 1A). fkd2 has a
defect in the venation pattern (Fig.
1B,D) and its mutation has been mapped close to the VAN3
locus (Steynen and Schultz,
2003). Complementation test between these genes demonstrate that
fkd2 was not rescued by van3-1, and its phenotype was
dominant over the strong van3 phenotype
(Fig. 1C-E), indicating that
FKD2 reduces VAN3 function. Sequencing of fkd2 revealed a
point mutation from G to A, which resulted in the change of amino acid from G
to E in the PH domain of VAN3 (Fig.
1A). We concluded that fkd2 is an allele of
van3, and we named fkd2 as van3-2. These results
suggest that the PH domain is necessary for VAN3 function to connect the leaf
vascular cells.
Subcellular localization of VAN3 in planta
We have reported from studies of the transient expression of VAN3-VENUS
that VAN3 may localize both to subpopulations of the TGN and to unknown
organelles in Arabidopsis suspension cells
(Koizumi et al., 2005). Here,
we re-examined the in situ localization of VAN3 using van3-1 mutant
lines that had been rescued by pVAN3::VAN3-VENUS
(Fig. 2A). To identify the
subcellular localization of VAN3-VENUS, we introduced GFP tagged-organelle
marker genes into the VAN3-VENUS plants by crossing.
The fluorescent signal of VAN3-VENUS showed dot-like localization in leaf
vascular cells (see Fig. S1A-C in the supplementary material). This dot-like
localization of VAN3-VENUS was observed in all tissues, including immature
leaves, petioles and the root stele, in which VAN3 was expressed (see Fig. S2
in the supplementary material). This result indicates that the dot-like
localization of VAN3 is common feature in VAN3-expressing cells. Next, we
investigated colocalization of the VAN3-VENUS and various organelle markers.
Because signals from leaf vascular cells were too weak to determine reliable
colocalization with leaf vascular cells, we used petiole cells and root
vascular cells in which enough fluorescence signals could be obtained.
Dot-like fluorescence of VAN3-VENUS colocalized with some but not all
fluorescence signals of the TGN marker GFP-SYP41
(Uemura et al., 2004)
(Fig. 2B-E). VAN3-VENUS did not
colocalize with the Golgi body marker ST-GFP
(Wee et al., 1998)
(Fig. 2F) or with the PVC
marker GFP-ARA7/RAB-F2b (Goh et al.,
2007) (Fig. 2G). As
in the subcellular localization of the VAN3-VENUS in Arabidopsis
suspension cells, VAN3-VENUS also localized to organelles that were not
recognized by known organelle markers (Fig.
2D,E, arrows). These results clearly indicate that VAN3 resides
both in a subpopulation of the TGN and in unknown organelles. Interestingly,
VAN3-VENUS was not distributed uniformly but formed discriminate domains in
the TGN marked by GFP-SYP41 (Fig.
2D,E, arrowheads). This result suggests that the TGN segregates
into distinct membrane domains that display different biochemical
compositions.
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Role of At5PTase for maintaining vascular continuity
We have reported that the PH domain of VAN3 binds preferentially to
PtdIns(4)P (Koizumi et al.,
2005). PtdIns(4)P is produced by the phosphorylation of
PI and the dephosphorylation of PtdIns(4,5)P2 through the
action of PI4 kinases and inositol 5' phosphatases, respectively.
CVP2, which encodes an inositol polyphosphate 5' phosphatase
(At5PTase), may function in the VAN3-related signaling pathway because of the
phenotypic similarity between cvp2 and van3-2 mutants
(Carland et al., 1999;
Carland and Nelson, 2004).
Therefore, we analyzed the genetic interaction of CVP2 and
VAN3 in terms of vascular continuity.
Vascular discontinuity caused by the cvp2 mutation was restricted in the marginal region of cotyledons and leaves (Fig. 1F), whereas van3-1 caused many vascular islands (VIs) in leaves (Fig. 1C). We presumed that this difference is due to the presence of a redundant paralogue of CVP2. Search of the Arabidopsis genomic database showed a candidate of the paralogue, At2g32010, which was named CVL1 (CVP2-like gene). CVL1 encodes a protein comprising 582 amino acids, of which 459 amino acids were identical to those of CVP2. Although cvl1-1 and cvl1-2 single mutants did not cause any vascular defect (data not shown), the cvp2cvl1-1 double mutant showed an enhanced phenotype of cvp2 (Fig. 1G), which is reminiscent of van3-1 (Fig. 1C). The same result was also obtained by using cvl1-2 allele (data not shown). These results suggest that both CVP2 and CVL1 function redundantly in the continuity of veins, although CVP2 is predominant.
Judging from the number of VIs, the phenotype of van3-2 was
enhanced by introducing the cvp2 mutation
(Fig. 1D,H), whereas the
phenotype of van3-1 was not affected by the cvp2 mutation
(data not shown). Genetic analysis of triple mutants between sfc-9,
cvp2 and cvl1-1 also identified that the strength of vascular
phenotype was essentially the same as sfc-9 (see Fig. S3A-C in the
supplementary material). It has recently been reported that VAN3 is involved
in the maintenance of continuous PIN1 expression domains in leaf primodia
(Scarpella et al., 2006). To
investigate whether CVP2/CVL1 is also involved in this process, we performed
PIN1 antibody staining in leaf primodia. Similar to the result obtained by
Scarpella et al. (Scarpella et al.,
2006), in sfc-9, files of cells expressing PIN1 were
first produced continuously and then became discontinuous during leaf
development, showing a defect of the maintenance of continuous PIN1 expression
(see Fig. S4M-P in the supplementary material). cvp2cvl1-1 showed the
same defect in maintenance of continuous PIN1 expression (see Fig. S4D-H in
the supplementary material) and the PIN1 expression pattern of
sfc-9cvp2cvl1-1 was essentially the same as that of sfc-9
and cvp2cvl1-1 (see Fig. S4Q-T in the supplementary material). These
results suggest that VAN3 and CVP2/CVL1 act on the pathway at the same point
to regulate the continuity of the leaf vasculature.
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CVP2-dependent subcellular localization of VAN3
VAN3 binds preferentially to PtdIns(4)P, a possible product of
CVP2, presumably through its PH domain
(Koizumi et al., 2005). The PH
domain also binds to soluble IP3
[Ins(1,4,5)P3], which is hydrolyzed by CVP2. Therefore,
CVP2 may affect VAN3 function by regulating the cellular levels of
PtdIns(4)P and/or IP3. Because the PH domain is related to
the subcellular localization of proteins, we examined whether CVP2
mutation would affect the subcellular localization of VAN3. At first, we
observed the localization of VAN3-sGFP in the leaf vasculatures of
cvp2cvl1-1 mutants. Although it was difficult to focus on detail
because of the weak fluorescence of VAN3-sGFP, it seems that localization of
VAN3 is disturbed in cvp2cvl1-1 mutants (see Fig. S1D,E in the
supplementary material). To clarify this issue, we performed subcellular
localization analysis using the root vascular cells that give us reliable
signals. In cvp2 mutants, dot-like structures derived from
sGFP-tagged VAN3 was not largely different from wild-type background
(Fig. 4D,E). By contrast,
combinational mutations between CVP2 and CVL1 induced the
mis-location of VAN3-sGFP from dot-like structures to the cytoplasm
(Fig. 4F). To explore the
cooperation of VAN3 and CVP2 further, we used a translational product of
van3-2 (VAN3G321E), which has a mutation within the PH
domain. Subcellular localization of VAN3G321E-sGFP mutants changed
dramatically from the dot-like to cytoplasmic localization, accompanied by few
dot-like structures in cvp2 (Fig.
4H), but essentially the same in the wild type
(Fig. 4G). These results
clearly indicate that functional CVP2 or CVL1 is necessary for localization of
VAN3 onto the TGN membranes.
Identification of new player function in phosphoinositide signaling-mediated vascular continuity
To isolate the new player acting on vascular continuity through
VAN3-mediated phosphoinositide signaling, we performed a yeast two-hybrid
screening and obtained At3g63300 as a candidate. We confirmed its interaction
by the yeast two-hybrid assay (see Fig. S5 in the supplementary material) and
named it as a VAN3-binding protein (VAB), which possessed a PH domain and an
unknown domain, DUF828 (Fig.
5A,B).
To determine whether VAB is associated with VAN3 in situ, we first examined
the colocalization of VAB and VAN3 by transient co-expression of VAB-eYFP and
VAN3-sGFP in Nicotiana benthamiana leaf epidermis. In epidermal
cells, VAB-eYFP and VAN3-sGFP colocalized as dot-like structures
(Fig. 5C-E). Functionality of
VAB-eYFP was confirmed by introducing pVAB::VAB-sGFP in vab
mutants (data not shown). To identify the direct interaction between VAN3 and
VAB in vivo, we performed a bimolecular fluorescence complementation (BiFC)
analysis (Bracha-Drori et al.,
2004; Walter et al.,
2004). As a control, we used the VAN3-like 3 (VAL3) protein, which
is a member of four plant ACAPs, including VAN3. No YFP fluorescence was
detected in Arabidopsis protoplasts in which VAB-cYFP (C terminal
fragment of YFP) was co-expressed with VAL3-nYFP (N terminal fragment of YFP)
(Fig. 5H,I). By contrast,
strong dot-like fluorescent signals were observed in protoplasts co-expressing
VAN3-nYFP and VAB-cYFP (Fig.
5F,G). This result suggests that VAN3 and VAB form a specific
complex in vivo.
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Because VAB possesses the PH domain and interacts with VAN3, we hypothesized that the VAN3-VAB complex functions in recognizing the PtdIns(4)P-enriched domain of the membrane, resulting in the recruitment of VAN3 to the correct membrane domain. To address this issue, we examined the subcellular localization of VAN3 in vab-1 mutants. The vab-1 mutation affected the subcellular localization of VAN3G321E-sGFP (Fig. 4I), as did the cvp2 mutation, although the subcellular localization of VAN3-sGFP did not change dramatically in vab-1 mutants (Fig. 4J). In addition, the vab-1 mutation enhanced the mislocation of VAN3-sGFP in the cvp2 mutant background (Fig. 4K; see Fig. S1D,F in the supplementary material). These results suggest that VAB is involved in the recruitment of VAN3 into the PtdIns(4)P-enriched membrane domains of the TGN.
Phosphoinositides-dependent ARF-GAP activity of VAN3
Next, we examined the effect of phospholipids on the ARF-GAP activity of
VAN3. To achieve this, GST-tagged VAN3, its derivatives and myristoylated
AtARF1 were produced in E. coli and purified as shown in the
Materials and methods. ARF-GAP activity was quantified using the change in
intrinsic Trp fluorescence during the transition from the GTP- to the
GDP-bound state of myristoylated AtARF1
(Antonny et al., 1997).
Although phospholipids such as phosphatidylcholine (PC), phosphatidylserine
(PS) and PI have no effects (see Fig. S6A in the supplementary material), the
ARF GAP activity of VAN3 was stimulated strongly by PtdIns(4)P, and
moderately by and PtdIns(4,5)P2 and phosphatidic acid (PA)
(Fig. 6A). By contrast,
application of a nonhydrolyzable analogue GMPPNP instead of GTP or application
of GST instead of GST-VAN3 did not reduce the fluorescence (see Fig. S6B,C in
the supplementary material). These results indicate that the ARF GAP activity
of VAN3 is dependent on specific phospholipids and that, among the
phospholipids, PtdIns(4)P promotes the ARF-GAP activity of VAN3 most
efficiently. This finding is consistent with our previous report that the VAN3
PH domain binds to PtdIns(4)P more strongly than to other
phospholipids (Koizumi et al.,
2005).
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Finally, we examined the effect of IP3 on ARF-GAP activity of
VAN3 because the CVP2 mutation elevates the level of IP3
(Carland and Nelson, 2004) and
enhances the phenotype of van3-2. Addition of IP3 in the
GAP assay solution inhibited the ARF-GAP activity of VAN3 in the presence of
PtdIns(4)P. The inhibitory effect of IP3 on the ARF-GAP
activity of VAN3 became stronger with higher concentrations of IP3
(Fig. 6A). This result raises
the possibility that CVP2 also upregulates ARF-GAP activity of VAN3 by
suppressing the cellular level of IP3.
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DISCUSSION |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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). However,
any animal ACAP is not localized to the TGN
(Jackson et al., 2000;
Peter et al., 2004). The
differences in subcellular localization between animal and plant ACAPs may
relate to differences in binding specificity to phosphatidyl inositols,
PtdIns(4)P for VAN3 versus PtdIns(4,5)P2 for
animal ACAPs (Jackson et al.,
2000). Thole et al. (Thole et
al., 2008) recently found using EYFP-FAPP1 as a
PtdIns(4)P staining agent that PtdIns(4)P is localized
primarily in the PM but also some internal membranes in Arabidopsis
root hairs. In addition, PtdIns(4)P is produced at the TGN by the
PI4Kβ1 in Arabidopsis
(Preuss et al., 2006). Our
results support the presence of PtdIns(4)P in the TGN and the PM, and
its function in the localization of TGN-residing proteins. In this context, it
is interesting that VAN3 is localized in the subdomain of the TGN membrane,
which might suggest the presence of the PtdIns(4)P-rich subdomain in
the TGN membrane. Future EM analysis will be required to prove the existence
of PtdIns(4)P-rich subdomain in the TGN.
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;
Xue et al., 2007). Although
little is known about the function of phospholipid signaling in venation
formation, only 5PTases appear to be involved in this process
(Xue et al., 2007). A mutation
in CVP2 encoding a plant type1 5PTase causes weak vascular
discontinuity (Carland and Nelson,
2004). A mutation in At5Pase13 also causes a defect in
cotyledon vein formation (Lin et al.,
2005). We have identified van3-2 weak allele mutants, the
phenotype of which is enhanced upon introduction of the CVP2
mutation. In addition, double mutants between CVP2 and CVL1
phenocopied the van3-1 strong allele. We also found that double
knockout of CVP2 and CVL1 caused mislocalization of VAN3
from the TGN form to the cytosolic form. These results imply that the normal
function of VAN3 in the right position is essential for the continuous
formation of vascular tissues. In addition, these results suggest that one or
more products from CVP2 and CVL1 5PTases are necessary for the correct
localization of VAN3.
In animals, type 1-5PTases hydrolyze only soluble inositol phosphates such
as IP3, whereas type 2-5PTases hydrolyze
PtdIns(4,5)P2 and PtdIns(3,4,5)P3, as
well as IP3 (Majerus et al.,
1999). In the Arabidopsis genome, at least 15 genes are
predicted to encode 5PTases, and these are classified into 11 type 1-like and
four type 2-like genes (Berdy et al.,
2001). Of the type 2-like PTases, two (AtPTase14 and FRAGILE
FIBER3) can hydrolyze PtdIns(4,5)P2 and
PtdIns(3,4,5)P3, as well as IP3, and the other
two (AtPTase12 and AtPTase13) can hydrolyze only IP3
(Zhong et al., 2004;
Zhong and Ye, 2004). At5PTase1
and At5PTase2, type 1-like 5PTases, exhibit only IP3-hydrolyzing
activity (Berdy et al., 2001;
Gunesekera et al., 2007).
However, a cooperative mutation in both At5PTase1 and
At5PTase2 caused both an increase in IP3 level and a
decrease in PtdIns(4)P level
(Gunesekera et al., 2007).
Interestingly, although CVP2 has been classified to the type 1-like 5PTase
family, CVP2 has been revealed recently to possess the ability to hydrolyze
both IP3 and PtdIns(4,5)P2 (Francine M. Carland
and Timothy Nelson, personal communication). Therefore, an increase in
IP3 level but also a decrease in PtdIns(4)P level is
expected to occur in the double knockout mutant of CVP2 and
CVL1. Indeed, an increased level of IP3 was observed in
cvp2 (Carland and Nelson,
2004). Association between the PH domain and the membrane is
prompted by some inositol phospholipids and inhibited by some soluble
inositols, such as IP3 (Lemmon
and Ferguson, 2000). From these results and the observation that
VAN3 binds strongly to PtdIns(4)P
(Koizumi et al., 2005), we may
think that VAN3 is recruited onto the PtdIns(4)P-rich subdomain of
the TGN by specific binding of its PH domain to PtdIns(4)P.
IP3 may suppress the binding. The facts that AT5PTase13
mutation caused defects in venation continuity and that double mutation of
At5PTase1 and At5PTase2 caused increase in IP3
level and a decrease in PtdIns(4)P level suggest that not only
CVP2/CVL1 but also other 5PTases are also involved in this process. The weaker
phenotype of cvp2cvl1 than that of sfc-9 is not inconsistent
with this notion. Therefore, CVP2 and CVL1, probably together with other
5PTase may contribute to the enhancement of TGN localization of VAN3
(Fig. 7).
VAB is a plant-specific protein containing the PH domain and functions in vascular continuity together with VAN3
Using the yeast two-hybrid system, we identified VAB as a novel
VAN3-interacting protein, the homologue of which is not found in animals. The
interaction between VAB and VAN3 in situ was strongly supported by (1)
colocalization of the two proteins to the TGN, (2) attachment of the two
proteins observed with the split YFP and (3) collaboration for vascular
continuity demonstrated by genetic analysis. VAB is a plant-specific protein
with a PH domain and an unknown domain, DUF828, and forms a small family with
the other seven homologues in the Arabidopsis genome. The subcellular
localization of VAN3G321E was changed in the vab mutation
background, as did the cvp2 mutation, suggesting that VAB is involved
in the proper targeting of VAN3. In addition, the vab mutation
enhanced the mislocation of VAN3-sGFP in the cvp2 mutant background.
Interestingly, VAL1, a VAN3 homologue, was not associated with VAB protein,
suggesting the specific binding of VAB with VAN3. It is known that many PH
domain-containing proteins have an inherent affinity for the membrane surface
that is insufficient by itself to drive their membrane localization, and the
formation of a protein complex with other proteins increases the avidity for
the membrane, resulting in the membrane recruitment
(Lemmon and Ferguson, 2000).
This suggests that VAN3 and VAB form a complex to bind to the
PtdIns(4)P-enriched membrane domain of the TGN, which was also
supported by the result of the split-YFP experiment.
PtdIns(4)P and IP3 control both the ARF-GAP activity and the subcellular localization of VAN3
Measurement of ARF-GAP activity indicated that PtdIns(4)P enhances
and IP3 inhibits ARF GAP activity of VAN3. This is the first
indication of phosphatidylinositol-dependent ARF-GAP activity in plants. In
animals, the GAP activity of ASAPs and ACAPs is stimulated by binding of
PtdIns(4,5)P2 and PA to their PH domains
(Randazzo, 1997;
Brown et al., 1998;
Jackson et al., 2000;
Kam et al., 2000). However,
our results showed the PtdIns(4)P-dependent activation of the VAN3
GAP activity. This result is consistent with the specific binding of
PtdIns(4)P to the VAN3 PH domain
(Koizumi et al., 2005). This
difference might relate to differences in the subcellular localization between
animal ACAPs and VAN3. The accelerating and inhibitory effects on
PtdIns(4)P and IP3 for VAN3 ARF-GAP activity,
respectively, should be related to the activity of CVP2 (probably and also
CVL1), which increase the PtdIns(4)P level and decrease the
IP3 level, as discussed above.
Upon these results, we present the model that antagonistic action of PtdIns(4)P and IP3 regulates both the subcellular localization and ARF-GAP activity of VAN3 (Fig. 7). CVP2 and CVL1 contribute to the increase in PtdIns(4)P levels and the decrease in IP3 levels. Both the activity and the localization of VAN3 may be coupled by specific binding of their PH domains to the PtdIns(4)P-rich domain in the TGN, and IP3 may suppress the binding and activation of VAN3. VAB helps VAN3 to bind to PtdIns(4)P-enriched membrane domains of the TGN. According to this model, in the cvp2cvl mutant, the reduction of PtdIns(4)P level on the membranes and an increased level of cellular IP3 would decrease the affinity of the PH domain of VAN3 for the membrane, resulting in the mislocalization of VAN3, as well as a decrease in ARF GAP activity.
Molecular function of VAN3 in continuous formation of vascular tissues
In conclusion, our findings strongly suggest that phosphoinositide
signaling plays a pivotal role in continuous vascular formation by regulating
the subcellular localization and ARF-GAP activity of VAN3. Polar auxin
transport is an essential factor that induces vascular pattern formation
(Sachs, 1981;
Scarpella et al., 2006). Sachs
(Sachs, 1981) proposed the
`auxin signal flow canalization hypothesis': auxin flow, starting initially
with diffusion, induces the formation of the polar auxin transport cell
system. This, in turn, promotes auxin transport and leads to canalization of
the auxin flow along a narrow file of cells. This continuous file of cells
differentiates into a strand of vascular cells. According to this hypothesis,
the positive-feedback system of auxin transport is a key event, but its
mechanism is still mysterious. This hypothesis agrees with accumulating data
on PIN proteins (Benková et al.,
2003). In the sfc root, the treatment with brefeldin A
results in the accumulation of PIN1, an auxin efflux carrier, in smaller and
more numerous compartments than in the wild type
(Sieburth et al., 2006). This
suggests that VAN3/SFC may be involved in PIN1 targeting. Conversely, a recent
report shows that auxin regulates phosphoinositide signaling
(Xue et al., 2007). Therefore,
it is possible that auxin promotes VAN3 function through CVP2-mediated
phosphoinositide signaling, and, in turn, that VAN3 promotes auxin transport
through the polar localization of PIN proteins to establish the vascular
continuity. We are currently investigating this hypothesis.
Supplementary material
Supplementary material for this article is available at
http://dev.biologists.org/cgi/content/full/136/9/1529/DC1
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ACKNOWLEDGMENTS |
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|
Footnotes |
|---|
* Present address: Department of Biology, University of Pennsylvania 415
South University Avenue, Philadelphia, PA 19104, USA 
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REFERENCES |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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